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ENVIRONMENTAL STRESS AND ADAPTATION TO STRESS

Ascorbic Acid Deficiency Activates Cell Death and Disease Resistance Responses in Arabidopsis¹

Centro de Investigaciones en Química Biológica de Córdoba/Consejo Nacional de Investigaciones Científicas y Técnicas, Departamento de Química Biológica, Facultad de Ciencias Químicas, Universidad Nacional de Córdoba, Ciudad Universitaria, Cordoba 5000, Argentina (V.P., M.E.A.); and Crop Performance and Improvement Division (E.O., G.K., S.M., S.K., C.H.F.) and Wheat Pathogenesis Program, Plant-Pathogen Interactions Division (J.A.), Rothamsted Research, Harpenden, Hertfordshire AL5 2JQ, United Kingdom

	ABSTRACT

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
Programmed cell death, developmental senescence, and responses to pathogens are linked through complex genetic controls that are influenced by redox regulation. Here we show that the Arabidopsis (Arabidopsis thaliana) low vitamin C mutants, vtc1 and vtc2, which have between 10% and 25% of wild-type ascorbic acid, exhibit microlesions, express pathogenesis-related (PR) proteins, and have enhanced basal resistance against infections caused by Pseudomonas syringae. The mutants have a delayed senescence phenotype with smaller leaf cells than the wild type at maturity. The vtc leaves have more glutathione than the wild type, with higher ratios of reduced glutathione to glutathione disulfide. Expression of green fluorescence protein (GFP) fused to the nonexpressor of PR protein 1 (GFP-NPR1) was used to detect the presence of NPR1 in the nuclei of transformed plants. Fluorescence was observed in the nuclei of 6- to 8-week-old GFP-NPR1 vtc1 plants, but not in the nuclei of transformed GFP-NPR1 wild-type plants at any developmental stage. The absence of senescence-associated gene 12 (SAG12) mRNA at the time when constitutive cell death and basal resistance were detected confirms that elaboration of innate immune responses in vtc plants does not result from activation of early senescence. Moreover, H₂O₂-sensitive genes are not induced at the time of systemic acquired resistance execution. These results demonstrate that ascorbic acid abundance modifies the threshold for activation of plant innate defense responses via redox mechanisms that are independent of the natural senescence program.

While the chemical nature of ROS dictates that they are potentially harmful to cells, plants use ROS as second messengers in signal transduction cascades regulating diverse processes such as mitosis, tropisms, and cell death. It is now well accepted that ROS accumulation is crucial to plant development as well as defense (Foyer and Noctor, 2005a). ROS signal transduction will ensue only if ROS escape destruction by cellular antioxidants that determine the lifetime and specificity of the signal. Ascorbic acid (AA) and glutathione are the major redox buffers of the plant cells, and they themselves are also signal-transducing molecules that can either signal independently or further transmit ROS signals (Fig. 1). They are thus intrinsic to redox homeostasis and redox-signaling events (Foyer and Noctor, 2005b).

View larger version (21K): [in this window] [in a new window]   Figure 1. The involvement of key redox couples in the expression of PR proteins in plant cells. The major soluble reductant couples are arranged according to their midpoint potentials, with principal protein components with which they interact indicated. DHAR, Dehydroascorbate reductase; Fdox, oxidized ferredoxin; Fdred, reduced ferredoxin; FTR, ferredoxin-thioredoxin reductase; GR, glutathione reductase. Solid arrows indicate known interactions, while broken arrows indicate putative components or redox couples implicated in the signal transduction cascade.

In addition to redox buffering and ROS detoxification (Foyer and Noctor, 2005a, 2005b), AA fulfills many essential roles in plant biology (Arrigoni and de Tullio, 2000). In particular, AA modulates growth through regulation of the cell cycle (Potters et al., 2002, 2004) and through regulation of elongation growth (Fry, 1998; Tokuna et al., 2005). AA and hydroxyl radicals participate in the oxidative scission of structural polysaccharides, promoting cell wall loosening (Fry, 1998). In addition, AA is a substrate for secretory peroxidases involved in cell wall stiffening and its presence limits the formation of phenolic radical intermediates in wall peroxidase reactions (de Pinto and De Gara, 2004). AA-dependent dioxygenases participate in the synthetic pathways of a number of key plant hormones that also influence growth (Arrigoni and de Tullio, 2000). For example, AA is required for the activity of 9-cis-epoxycarotenoid dioxygenase, an enzyme catalyzing the formation of xanthoxin, the precursor of abscisic acid.

A number of Arabidopsis (Arabidopsis thaliana) mutants that have low AA (vitamin C; vtc) have been isolated (Conklin et al., 2000). Since the function of most of the genes modified in these mutants is unknown, we have concentrated our efforts on analyzing vtc1 and vtc2, which are better characterized. Most importantly, the vtc1 and vtc2 phenotypes are caused by low AA alone, as shown by studies involving the expression of the animal AA biosynthetic enzyme L-gulono-1,4-oxidase, which restores wild-type AA levels and the wild-type phenotype to these mutants (Radzio et al., 2003). vtc1 is relatively well studied, harboring a point mutation in the AA biosynthetic enzyme GDP-Man pyrophosphorylase (Conklin et al., 2000). This mutant was instrumental in the characterization of the pathway of AA synthesis in plants (Conklin et al., 1999). In contrast, vtc2 (At4g26850) harbors a less-characterized mutation in an unknown protein (for gene database information, see, for example, http://mips.gsf.de/projects/plants).

The vtc1 mutation confers sensitivity to ozone and other abiotic stresses, such as freezing and UV-B irradiation (Conklin et al., 1996), but it enhances pathogen resistance (Barth et al., 2004). We have previously shown that AA deficiency in the vtc1 mutant led to the differential expression of 171 genes, a substantial number of which encode pathogenesis-related (PR) proteins such as PR1, PR2, and PR5 (Kiddle et al., 2003; Pastori et al., 2003). Moreover, PR proteins accumulated more rapidly in vtc1 than the wild type when challenged with Pseudomonas syringae (Barth et al., 2004). However, when grown under optimal growth conditions, vtc1 shows no evident indications of increased oxidative stress in terms of tissue H₂O₂ content or the redox state of key indicator pools such as AA (Veljovic-Jovanovic et al., 2001).

Like AA (1–5 µmol mg^–1 fresh weight or µmol mg^–1 chlorophyll), glutathione is an abundant plant metabolite (100–300 nmol mg^–1 fresh weight or nmol mg^–1 chlorophyll) that has many diverse and important functions (Noctor and Foyer, 1998), including signal transduction (Noctor et al., 2002; Gomez et al., 2004). As an indicator of the general cellular thiol-disulfide redox balance, the reduced glutathione (GSH)-oxidized glutathione (GSSG) couple is well suited to the role of redox sensor. Cytosolic thiol-disulfide status appears to be important in regulating the expression of PR genes through the nonexpressor of PR protein 1 (NPR1) pathway (Mou et al., 2003). NPR1 is an intrinsic component of the salicylic acid (SA)-triggered systemic acquired resistance (SAR) response to biotic attack. The redox dependence of the NPR1 pathway implies that biotic or abiotic stimuli that perturb the cellular redox state can up-regulate defense genes via the NPR1 pathway (Mou et al., 2003). Such redox-linked effects explain, for example, PR gene expression in response to UV-B exposure (Green and Fluhr, 1995) and in catalase-deficient mutants (Chamnongpol et al., 1996). Figure 1 illustrates the redox relationships between the different components implicated in the regulation of NPR1 movement from the cytosol to the nucleus to elicit SAR and PR gene expression.

The redox poise of the plant cell at any moment in time is largely dictated by three redox pools, namely, pyridine nucleotides, AA, and glutathione (Fig. 1). The high cellular AA content dictates that it is the major low-M_r antioxidant and redox buffer of plant cells (Foyer, 2004). It is therefore logical to pose the question of whether low redox buffering capacity alone, as occurs during AA deficiency, is sufficient to trigger a redox-sensitive pathway leading to enhanced basal resistance. To address this question, we have analyzed SAR and pathogen resistance in two AA-deficient Arabidopsis mutants, vtc1 and vtc2. We provide evidence that a decrease in overall cellular redox-buffering capacity in these mutants, resulting from a diminished AA pool, together with an enhanced GSH-to-GSSG ratio, reduces the threshold for local PCD and causes movement of NPR1 into the nucleus, triggering systemic resistance responses in the absence of enhanced ROS levels or external cues.

	RESULTS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 

AA Deficiency Retards Growth and Senescence and Enhances Leaf GSH-to-GSSG Ratios

In this study, the Columbia (Col-0) wild type and the vtc1 and vtc2 mutants were grown in a 10-h photoperiod in controlled environment chambers with filtered air to remove atmospheric ozone. In these conditions, the vtc1 (Fig. 2) and vtc2 (data not shown; see Radzio et al., 2003) mutants were smaller at equivalent stages in development than wild-type plants. This phenotype was clearly evidenced at 2 weeks and was maintained throughout the plant growth cycle (Fig. 2). As shown in Table I, the vtc1 and vtc2 rosettes accumulate biomass at a much slower rate than the wild type. The vtc1 leaves had about 70% less AA than the wild type throughout development. The vtc2 mutants studied here had even lower leaf AA content for most of the growing period, with leaf AA reaching maximum values of about 25% of those of the wild type at 4 weeks (Table I). In addition, the vtc1 leaves had 1.3 to 1.6 times more glutathione than the wild type. Moreover, the GSH-to-GSSG ratios of vtc1 rosette leaves were almost double those measured in the wild type at all stages of development. The vtc2 rosette leaves also tended to have more leaf glutathione than the wild type, with much higher GSH-to-GSSG ratios later in rosette development (Table I). Wild-type, vtc1, and vtc2 leaves had similar amounts of chlorophyll and leaf protein at equivalent time points in the growth phase up to 10 weeks of growth. Thereafter, leaf chlorophyll and soluble protein declined in wild-type plants, but in vtc1 and vtc2 they remained at similar values to those observed at 8 weeks (Table I). On the other hand, the vtc1 plants were delayed in flowering compared to the wild type (Fig. 3), as did vtc2 (see Fig. 2). Together, these features show that vtc1 and vtc2 rosettes senesce later than the wild-type leaves.

View larger version (75K): [in this window] [in a new window]   Figure 2. The delayed development phenotype of the vtc1 (vtc1) and vtc2 (vtc2) mutants compared to the wild type (WT). The number of weeks after sowing is indicated.

View larger version (11K): [in this window] [in a new window]   Figure 3. The appearance of the apical meristem is delayed in the vtc1 mutant compared to the wild type. The days to flowering were compared in vtc1 () and wild-type plants (). The experiment was performed using 100 plants and repeated three times with similar results.

Macroscopic spontaneous lesions were not observed in naive vtc1 leaves from plants grown on either short or long days (Fig. 2), but a few whole chlorotic leaves per plant appeared after 10 weeks, and these, by eye, might easily be mistaken for a symptom of senescence. However, in contrast to Col-0 rosette leaves (Fig. 4A), we consistently detected the presence of individual dead cells early in the development of vtc1 (Fig. 4B) and vtc2 (Fig. 4C) rosette leaves at the microscopic level. Small foci of collapsed cells were found all over the leaf surface and preferentially in the mesophyll layer of vtc1 and vtc2 leaves (Fig. 4, B and C). By 8 weeks, the areas of dead cells had expanded in vtc2 leaves to form larger foci with clear patches of dead cells detectable by autofluorescence or lactophenol blue staining (Fig. 4C). Even the young (4–6 weeks) vtc1 and vtc2 rosettes had a small number of dead cells as detected by autofluorescence (Fig. 4, B and C) or lactophenol blue staining (Figs. 4 and 5). Individual dead cells were also detected in wild-type leaves, but their appearance was delayed compared to vtc1 and vtc2 (Figs. 4 and 5).

View larger version (79K): [in this window] [in a new window]   Figure 4. The appearance of individual dead cells (microlesions) in the rosette leaves of wild-type (A), vtc1 (B), and vtc2 (C) plants during development. Cell death was monitored using autofluorescence (top) or lactophenol blue staining (bottom) of leaf tissues. To aid clarity, examples of individual dead cells are marked with fine arrowheads, while large patches of dead cells are marked by thick arrowheads in the bottom images.

View larger version (11K): [in this window] [in a new window]   Figure 5. The number of dead cells present in vtc1 and wild-type leaves through development. Values were calculated from leaves stained with lactophenol blue as shown in Figure 4, and data represent the means ± SE of five separate experiments involving 25 samples per line.

To evaluate how vtc1 controls cell death expansion once it is initiated by an exogenous stimulus, we challenged leaf tissues with high doses of the biotrophic bacterium P. syringae pv tomato DC3000 (Pst). This virulent pathogen proliferates in the intercellular spaces of leaf tissues of naive wild-type plants causing disease with spreading, chlorotic lesions (Whalen et al., 1991). However, proliferation is restricted when plants orchestrate SAR (Uknes et al., 1992; Cameron et al., 1994). We monitored the development of pathogen-induced lesions in 8-week-old wild-type and vtc1 and vtc2 plants infiltrated with a high titer of bacteria (5x10⁶ cfu/mL). Large patches of necrotic tissue were evident 5 d postinfection in inoculated wild-type leaves, while only a few small groups of dead cells (without spreading necrosis) were found in inoculated vtc1 tissues (Fig. 6). Thus, vtc1 mutation does not lead to cell death propagation once it is initiated by Pst inoculation. Like vtc1, vtc2 leaves exhibited individual collapsed cells in the young (4–6 weeks) rosettes. Spontaneous cell death is therefore more prolific in vtc1 than vtc2 leaves, whereas cell death patches were more abundant in vtc2 leaves in which AA content is much lower than vtc1 at the 6- and 8-week stages (Table I). Individual dead cells were also detected in wild-type leaves, but their appearance was delayed compared to vtc1 (Figs. 4 and 5).

View larger version (137K): [in this window] [in a new window]   Figure 6. Pathogen-induced macroscopic and microscopic cell death symptoms are greatly decreased in Pst-infected vtc1 leaves compared to Pst-infected wild-type leaves. A and B, Lactophenol blue staining of 7-week-old wild-type and vtc1 leaves before (A) and after (B) pathogen inoculation (5 x 106 cfu/mL). The black arrows in A indicate the presence of individual dead cells in vtc1 but not wild type (WT) in the absence of inoculation. Attached leaves were infiltrated with Pst at single-point sites (black arrowheads; B [top]), and pathogen-induced cell death was analyzed at 5 d postinfiltration. Middle section, The whole leaves show that patches of dead cells occur around inoculation sites in wild type but not in vtc1. Magnification of the infiltration sites (B [bottom]) shows massive cellular collapse (indicated by fat arrowheads) in the wild-type plants, whereas only a few small groups of dead cells in vtc1 (thin arrows). Bars, 50 µm.

View larger version (59K): [in this window] [in a new window]   Figure 7. The vtc1 and vtc2 mutants display enhanced resistance to infection by Pst. Leaves from 6-week-old plants were locally infiltrated with Pst (5 x 106 cfu/mL), and symptoms were analyzed at the infiltration sites (indicated by yellow arrows) at 4 d postinfection. Visible disease symptoms, displayed by Pst-infected wild-type (Col-0) leaves, were much reduced in vtc1 and vtc2 plants (A). Moreover, Pst growth curves in planta, quantified in leaf discs excised at the indicated times from the infiltration sites inoculated with a low titer of Pst (105 cfu/mL), show that bacterial growth is restricted in vtc1 and vtc2 leaves compared with wild-type (Col-0) controls. Data represent the mean ± SE of three independent experiments (B).

View larger version (14K): [in this window] [in a new window]   Figure 8. Growth of Pst is increasingly restricted as vtc1 rosettes develop. Leaves from 2-, 4-, 6-, 8-, and 11-week-old plants were infiltrated with Pst (105 cfu/mL). Pst growth in planta was quantified in leaf discs excised from infiltration sites at 5 d postinfiltration. Data represent the mean ± SE of three independent experiments.

By week 6, both mutants displayed constitutive expression of PR genes (Fig. 9). However, neither vtc mutants nor wild-type plants displayed substantial constitutive expression of the antioxidant glutathione S-transferase (GST) gene at this time (Fig. 9). PR gene expression was first detected in naive vtc1 plants after 6 weeks of growth and was clearly evident by week 8 (Fig. 10). In contrast, PR expression was active only at the time of senescence in wild-type plants (about 16 weeks; Fig. 10). Neither Col-0 nor vtc1 plants showed expression of the senescence marker gene senescence-associated gene 12 (SAG12) during the developmental period in which leaves were analyzed (2–11 weeks) in these experiments (Fig. 10). The absence of SAG12 expression in vtc1 confirms the conclusion drawn from the enhanced duration of levels of high protein and chlorophyll that vtc1 rosettes do not develop faster or senesce earlier than the wild-type plants.

View larger version (94K): [in this window] [in a new window]   Figure 9. Six-week-old vtc1 (left) and vtc2 (right) rosette leaves constitutively express PR transcripts. RNA-blot analyses of wild-type, vtc1, and vtc2 naive leaf tissues were hybridized with Arabidopsis probes (Uknes et al., 1992). The 5S rRNA probe was used as the gel-loading control.

View larger version (30K): [in this window] [in a new window]   Figure 10. Accumulation of defense-related markers. The abundance of PR5 and SAG12 mRNA was determined on RNA blots prepared from naive leaves of 2-, 4-, 6-, 8-, and 11-week-old vtc1 and wild-type (Col-0) plants. Gel loading was controlled by hybridization with the 5S rRNA probe.

The mature vtc1 and vtc2 leaves had the same numbers of cells as the wild type, but they were smaller (Fig. 11). The vtc1 leaf cells stopped growing early in development at about week 6 (Fig. 12). Transgenic wild-type and vtc1 plants expressing green fluorescent protein (GFP) fused to NPR1 (Col-0 GFP-NPR1 and vtc1 GFP-NPR1, respectively) were constructed and the fluorescence profile was assessed throughout the development of homozygous transformed plants. The amount of GFP fluorescence observed in the cytosol of mesophyll or epidermal cells was very low in all conditions. However, the stomatal guard cells of all genotypes always showed nuclear localization of NPR1 as observed by high fluorescence (Fig. 13), as reported previously (Mou et al., 2003). When the fluorescence images are overlaid bright-field images of the cells (Fig. 14, bottom), then nuclear localization of the fluorescent spots within the guard cell pairs is evident. A higher resolution image of an individual stomatal guard cell pair is shown in the inset to Figure 14 (bottom right) to clearly illustrate this point.

View larger version (95K): [in this window] [in a new window]   Figure 11. Light microscopy sections of naive leaves showing the comparative cell structure of 10-week-old plants. Wild-type leaves (A) have larger cells than vtc1 (B) or vtc2 (C) leaves. Bar, 100 µm.

View larger version (11K): [in this window] [in a new window]   Figure 12. The effect of rosette age on cell area in vtc1 (), vtc2 (), and wild-type () leaves.

View larger version (54K): [in this window] [in a new window]   Figure 13. Detection of the cellular localization of GFP fused to NPR1 in transformed 4-week-old wild-type (Col-0-NPR1; top) and vtc1 plants (vtc1-NPR1; middle). Top and middle sections show fluorescence only in stomatal guard cells. The bottom section shows that SA causes fluorescence to appear in mesophyll nuclei as well as in guard cells. The bottom image shows fluorescence in 4-week-old Col-0-NPR1 24 h after spraying with 1 mM SA.

View larger version (81K): [in this window] [in a new window]   Figure 14. The effect of rosette age on the cellular localization of GFP fused to NPR1 in transformed Col-0 (Col-0-NPR1) and vtc1 (vtc1-NPR1) plants. Fluorescence in the nuclei of mesophyll (arrowheads) and stomatal (fine arrows) guard cells is indicated for leaf samples of 6- and 8-week-old plants. For the vtc1-NPR1 8-week-old sample, the bright-field image (bottom left) of the leaves has been overlaid with the GFP fluorescence (bottom right) to illustrate the cellular localization of the fluorescence. The inset shows a higher focus image of an individual stomatal guard cell pair in the epidermis in the overlaid image.

	DISCUSSION

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
As in animals, plant PCD is a physiological process that is indispensable for correct development and regulates homeostasis through the elimination of unneeded cells. In contrast to necrosis, PCD is triggered by intrinsic factors or extrinsic defense cues and is executed through genetic signals that lead to a cell suicide program through the activation of defined autolytic processes. Current concepts suggest that pathogen-induced PCD is triggered by an interaction of signaling molecules, such as ROS and nitric oxide, and simultaneous suppression of ROS detoxification (Delledonne et al., 2001; Apel and Hirt, 2004). ROS-mediated cell suicide events are genetically controlled (Wagner et al., 2004) and ROS are required in many (but not all) systems leading to pathogen-induced PCD (Apel and Hirt, 2004). The intrinsic pathway involves two of the most important ROS-generating organelles of the plant cell, the chloroplasts and the mitochondria (Chen and Dickman, 2004; Yao et al., 2004). The aim of this study was to elucidate the mechanisms whereby AA deficiency in the vtc1 and vtc2 mutants modifies pathogen-induced PCD and enhances pathogen resistance. In addition, the experiments were designed to explore the role of AA in the coordinate control of growth and defense. We used the vtc1 and vtc2 mutants because the slow-growth phenotype observed in these mutants can be reversed by expression of L-gulono-1,4-oxidase, which restores wild-type AA levels and the wild-type phenotype to these mutants (Radzio et al., 2003). These results suggest that low AA alone causes the observed phenotypic characteristics of the vtc1 and vtc2 mutants. The results presented in this article allow us to draw the following conclusions.

AA Deficiency Stimulates the Innate Immune Response via Activation of Localized PCD Rather Than Early Senescence

PCD events in plants that can be triggered by elicitors, such as harpin (Boccara et al., 2001), or by pathogens (Schon et al., 2005) can involve an early loss of photosynthesis and ROS accumulation in chloroplasts (Joo et al., 2005). Moreover, photosynthetic electron transport inhibitors block lesion formation in the lesion mimic mutant (LMM), lsd1. Similarly, ROS-mediated lesion development is favored by high light and low CO₂, implicating photorespiratory H₂O₂ production in the PCD response (Mateo et al., 2004). Under optimal growth conditions, photosynthesis is largely unaffected in the vtc1 and vtc2 mutants (Pastori et al., 2003). However, photosynthetic CO₂ assimilation and other related parameters are much more sensitive to inhibition by environmental stress and high light in vtc mutants compared to the wild type (Mulle-Moule et al., 2002). Here we show AA deficiency limits overall cell size, an effect that becomes apparent only after the first weeks of plant development. Total leaf cell number and size were measured every 2 weeks throughout rosette development. These data show that the wild-type and mutant leaves contain more or less the same number of cells, but that final cell size is smaller in the vtc1 and vtc2 mutants than in the wild type. Although leaf AA content is low throughout development, cell area is affected only in older rosettes (age more than 6 weeks; Fig. 12). This result is consistent with known effects of AA on cell wall expansion and cross-linking (Fry, 1998; de Pinto and De Gara, 2004). Interestingly the cessation of growth coincides with the appearance of microscopic lesions (cell death events). Moreover, NPR1 is detectable in the nuclei and key features of SAR appear (PR gene expression and increased resistance to virulent pathogens) only after this point, suggesting an inverse correlation between cell growth and activation of innate immune responses.

In Arabidopsis, the age of the leaf is considered to be a predictor of the timing of senescence, the sequential senescence of rosette leaves coinciding with the time of maximum inflorescence development (Hensel et al., 1993). Based on SA accumulation and the early expression of SAGs, Barth et al. (2004) attributed the enhanced pathogen resistance in vtc mutants to early senescence. During senescence, leaf metabolism is shifted toward the breakdown of macromolecules, particularly leaf soluble protein and chlorophyll. While the vtc mutant leaves accumulate increasing numbers of microlesions through development, they do not show early senescence. Evidence supporting this conclusion is as follows: (1) the vtc leaves retain chlorophyll and protein longer than those of the wild type; (2) the vtc mutants flower later than the wild type; and (3) SAG12 mRNA was absent at the time when microlesions developed. Taken together, these results confirm that cell death in vtc mutants does not result from activation of a senescence program. Since low AA does not trigger early senescence, other effects of AA deficiency must trigger the events leading to enhanced basal resistance.

The data shown in Figure 9 indicate that the expression of H₂O₂-induced genes, such as GST, is not constitutively induced in vtc1. Moreover, when we analyzed the expression profile of H₂O₂-sensitive genes in vtc1 using a database search, the vtc1 transcriptome signature (Pastori et al., 2003) showed no characteristic features of H₂O₂-mediated gene expression (data not shown). Then we compared the effect of AA feeding on the vtc1 transcriptome (Pastori et al., 2003) with that produced in Arabidopsis cells by H₂O₂ application (Desikan et al., 2001). The data for both the H₂O₂ application (Desikan et al., 2001) and AA feeding microarrays were downloaded from the Stanford Microarray Database (SMD) site and analyzed as described by Desikan et al. (2001) to ensure uniform data analysis, the rationale being that any genes controlled by H₂O₂ in vtc1 would show reversed expression patterns after AA feeding. AA feeding also modified only 6% of the genes with altered expression patterns following H₂O₂ treatment (Table II). H₂O₂ application led to changes in the expression of 175 expressed sequence tags, of which 113 were induced (26 low expressing, 87 high expressing) and 62 were repressed (Desikan et al., 2001). Of the 106 genes repressed in vtc1 by AA (Pastori et al., 2003), three of these were also changed by H₂O₂ (two were repressed and one was induced). Of the 112 genes induced by AA (Pastori et al., 2003), Desikan et al. (2001) also reported nine of these, but all were induced. Thus, genes whose expression was modulated by both H₂O₂ and AA showed similar patterns of expression rather than inverse trends. Taking into account effects due to random patterns of gene expression, these results show that the AA and H₂O₂ transcriptome signatures are different, with no common genes with patterns of expression consistent with predicted direct oxidant/antioxidant effects (Table II).

In order to elucidate whether the molecular mechanisms leading to activation of pathogen resistance and SAR responses in the mutants were modulated by altered redox-buffering controls, we first measured the total amount of glutathione and the GSH-to-GSSG ratios in vtc1, vtc2, and wild-type leaves. The mechanisms through which the SA signal pathway functions are not completely understood, but the ankyrin repeat protein NPR1 is one of the key regulators of SA-dependent gene expression (Cao et al., 1994; Delaney et al., 1995). It has recently been demonstrated that NPR1 is converted from an inactive oligomer to an active monomer as a result of cellular redox changes induced by SA during SAR (Mou et al., 2003). The monomer form of NPR1 moves into the nucleus, where it activates expression of defense genes such as PR1 via redox interaction with TGA transcription factors (Després et al., 2003). Both monomerization of NPR1 oligomers and reduction of monomeric NPR1 involve thiol-disulfide exchange reactions with the participation of glutathione and specific forms of thiol-active proteins, such as thioredoxins, as shown in Figure 1 (Vanacker et al., 2000; Mou et al., 2003; Laloi et al., 2004). The total amount of glutathione and the GSH-to-GSSG ratios were increased in vtc1 leaves compared to wild-type leaves along development. These data suggest that vtc1 leaves have key adjustments in the glutathione pool that would favor the monomerization and nuclear translocation of NPR1, resulting in the enhancement of the basal resistance level of the cell. Modulation of cellular glutathione content transmits information through diverse signaling mechanisms, including the establishment of an appropriate redox potential for thiol-disulfide exchange and release of calcium to the cytosol (Gomez et al., 2004). However, the changes in glutathione reported here, since 2 weeks of growth, are not alone sufficient to trigger PR gene expression, which occurs only after 6 to 8 weeks of growth when NPR1 is increasingly located in the nucleus.

The Onset of SAR Is Linked to the Cessation of Expansion Growth and the Appearance of Microlesions But Not to ROS Enhancement in vtc Mutants

Our results suggest that another factor associated with low AA is triggered during development and that this, together with favorable GSH-to-GSSG ratios, stimulates SAR. AA deficiency in vtc1 and vtc2 activates cell death and the development of microlesions from about 4 weeks and prevents cell expansion from 4 to 6 weeks onward. The onset of SAR is linked in time to the cessation of expansion growth and the appearance of microlesions in vtc1 and vtc2. Nearly 40 Arabidopsis LMMs that develop HR-like lesions on their leaves have been characterized to date. Some, but not all, LMMs develop enhanced disease resistance (Lorrain et al., 2003). Among the LMM mutants having enhanced resistance to virulent infections, cpn1 (copine), hrl1 (HR-like lesions), and hlm1 (HR-like lesion mimic) show low-level or microscopic cell death (Jambunathan et al., 2001; Devadas et al., 2002; Balagué et al., 2003) similar to that observed in vtc1 leaves. We have found no evidence that vtc1 and vtc2 stimulate low-level cell death by ROS enhancement. We conclude that the threshold of sensitivity to ROS produced during the normal course of cell development and growth is decreased because of low redox buffering capacity. Localized ROS accumulation occurs in a wide range of hormone-dependent developmental signaling processes, as well as in cell wall cleavage and associated cell wall growth in a diverse range of developing and expanding organs, such as embryonic axes (Puntarulo and Cederbaum, 1988), roots (Joo et al., 2001; Foreman et al., 2003), germinating seeds (Schopfer et al., 2001), expanding leaves (Rodriguez et al., 2002), and coleoptiles (Schopfer et al., 2002). The absence of apoplastic AA in vtc1 means that ROS generated during normal developmental signaling will have a longer lifetime than in wild-type plants. In addition, zinc finger proteins and transcription factors involved in PCD and SAR are probably activated in these conditions by virtue of their modulation by cellular redox poise.

In summary, the results demonstrate that AA deficiency positively modulates plant biotic defense cascades leading to greater disease resistance as reported by Barth et al. (2004), but here we show that this effect is not caused by early senescence, as suggested by these authors. In contrast, our results show that vtc rosettes exhibit nuclear localization of the NPR1-GFP and SAR in association with localized PCD events. The observed overall smaller leaf cell size in the vtc leaves is consistent with the known roles of AA in cell wall formation. However, this cannot be the sole explanation of the smaller cell area in the vtc lines because cell expansion is affected only in older rosettes, even though AA content is low throughout leaf development. It is possible that AA levels also modulate DELLA protein activity and hence growth by modulation of gibberellic acid and abscisic acid levels, as discussed previously (Pastori et al., 2003). The inverse relationship between growth and defense that is frequently observed in plants is crucial in any consideration of plant productivity, but it is poorly understood. The results presented suggest that cellular AA may be one component that influences this relationship acting either downstream or alongside ROS accumulation to coordinate plant growth and defense processes.

	MATERIALS AND METHODS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 

Plant Growth

Seeds of the wild-type Arabidopsis (Arabidopsis thaliana L. Heynh. accession Col-0) and the vtc1-1 and vtc2-2 mutants were obtained from the Nottingham Arabidopsis Stock Centre (NASC) and from Nick Smirnoff (University of Exeter). These plants were grown in a controlled environment chamber essentially as described previously (Veljovic-Jovanovic et al., 2001). Plants were grown for 12 weeks at 22°C under low (250 µmol m^–2 s^–1) light with a 10-h photoperiod and 70% humidity. The air in the chamber was filtered to remove atmospheric ozone and with strict hygiene.

Biomass, Chlorophyll, and Metabolite Analyses

Whole rosettes were harvested in the growth chamber and biomass recorded (fresh weight). Extraction and assay of AA, glutathione, protein, and chlorophyll were performed as described previously (Vernon, 1960; Veljovic-Jovanovic et al., 2001).

Pathogen Inoculation

The virulent Pseudomonas syringae pv tomato strain DC3000 (Pst) was grown overnight in King's B medium with 10 µg/mL tetracycline and 100 µg/mL rifampicine, and bacterial suspensions were washed twice in 10 mM MgCl₂, diluted to appropriate concentrations and used for infiltration into fully expanded leaves at two sites on the abaxial surface, as previously described (Alvarez et al., 1998). For bacterial growth curves, two leaf discs (6-mm diameter) were cut around the Pst-inoculation sites from each leaf. Discs from different inoculated leaves were pooled at each time point and homogenized in 10 mM MgCl₂ to liberate the bacteria. Serial dilutions of the homogenates were plated (in duplicate) on King's B medium supplemented with 10 µg/mL tetracycline and 100 µg/mL rifampicine. Colonies on the plates were counted after incubation at 28°C for 24 h.

Northern-Blot Analysis

Total RNA was prepared for RNA gel-blot hybridization analysis by use of standard protocols as described previously (Alvarez et al., 1998). Arabidopsis probe templates were used for detection of GST-1 (At1g02930), SAG12 (At5g45890), PR-1 (At2g14610), PR-2 (At3g57260), and PR-5 (At1g75040) transcripts (Uknes et al., 1992).

Plant Transformation

Competent Agrobacterium tumefaciens (strain GV3101; pMP90) were prepared using the protocol of McCormac et al. (1998). Arabidopsis (Col-0/vtc1) plants were transformed with Agrobacterium containing the NPR1-GFP construct (pBI 1.4t backbone; Kinkema et al., 2000) using a simple dip transformation technique (Clough and Bent, 1998). Primary transformed seedlings were selected on Murashige and Skoog agar plates containing kanamycin (2.1 g/L Murashige and Skoog salts, 0.7% bacto-agar, and 50 µg/mL kanamycin, pH 5.7).

Microscopy

GFP fluorescence in leaves expressing the NPR1 reporter constructs was determined using a Zeiss Axiophot microscope. Leaf samples were mounted in water and illuminated using an excitation and emission wavelength of 490 and 510 nm, respectively, and viewed using the appropriate filters (BP 470/20, FT 493, BP 505–530; Zeiss).

Histochemistry

Leaf sections were fixed and embedded in Spurr's medium as described by Jones et al. (2002). Semithin sections (0.5 µm) were stained with toluidine blue and examined with a Leica DMR light microscope. Quantitative determinations were developed using an imaging analysis Leica QM500 as described previously by Olmos and Hellín (1998).

Dead cells were identified using lactophenol blue (Fluka) staining followed by destaining in saturated chloral hydrate as previously described (Waspi et al., 2001). Leaves were then mounted on microscopic slides in 40% glycerol and analyzed by bright-field or blue-light incident fluorescence microscopy. Numbers of dead cells per leaf were counted and 40 independent measurements were made per time point. Autofluorescence detection was performed as described previously (Waspi et al., 2001).

	ACKNOWLEDGMENTS

 
We thank Xinnian Dong for the generous gift of the NPR1-GFP construct.

Received June 28, 2005; returned for revision August 12, 2005; accepted August 12, 2005.

	FOOTNOTES

 
¹ This work was supported by the Biotechnology and Biological Sciences Research Council (C.F., G.K., and J.A.); grants from Agencia Nacional de Promoción Científica y Tecnológica (BID 1201/OC–AR PICT 01–10123) and Fundación Antorchas and Secretaria de Ciencia y Tecnologia/Universidad Nacional de Córdoba (to M.E.A.); the Department of Biotechnology, Government of India, for a Biotechnology Overseas Associateship Award (to S.K.); the Royal Society (U.K.) for a short-term fellowship (to S.M.); CONICET for a fellowship (to V.P.); and the Spanish Government for a Mobility Grant of Researcher, Ministerio de Educacion y Ciencia (PR2004–0361; to E.O.).

² Present address: Centro de Edafologia y Biologia Aplicada del Segura/Consejo Superior de Investigaciones Científicas, Department of Plant Physiology, P.O. Box 164, 30080 Murcia, Spain.

³ Present address: Biotechnology Division, Institute of Himalayan Bioresource Technology, P.O. Box 6, Palampur–176 061 (HP), India.

The authors 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.plantphysiol.org) are: Christine H. Foyer (christine.foyer{at}bbsrc.ac.uk) and María E. Alvarez (malena{at}mail.fcq.unc.edu.ar).

Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.067686.

^* Corresponding author; e-mail christine.foyer{at}bbsrc.ac.uk; fax 0044–1582–763010.

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