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Keywords:

  • ASCORBATE PEROXIDASE 2;
  • reactive oxygen species;
  • photosynthesis;
  • wounding;
  • jasmonic acid;
  • chitosan

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

ASCORBATE PEROXIDASE 2 (APX2) encodes a key enzyme of the antioxidant network. In excess light-stressed Arabidopsis leaves, photosynthetic electron transport (PET), hydrogen peroxide (H2O2) and abscisic acid (ABA) regulate APX2 expression. Wounded leaves showed low induction of APX2 expression, and when exposed to excess light, APX2 expression was increased synergistically. Signalling pathways dependent upon jasmonic acid (JA), chitosan and ABA were not involved in the wound-induced expression of APX2, but were shown to require PET and were preceded by a depressed rate of CO2 fixation. This led to an accumulation of H2O2 in veinal tissue. Diphenyl iodonium (DPI), which has been shown previously to be a potent inhibitor of H2O2 accumulation in the veins of wounded leaves, prevented induction of APX2 expression probably by inhibition of PET. Thus, the weak induction of APX2 expression in wounded leaves may require H2O2 and PET only. As in other environmental stresses, wounding of leaves resulted in decreased photosynthesis leading to increased reactive oxygen species (ROS) production. This may signal the induction of many ‘wound-responsive’ genes not regulated by JA-dependent or other known JA-independent pathways.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Plants react to physical damage by locally and systemically activating groups of genes that may contribute to wound repair and defence against opportunistic pathogens. The physiological and biochemical responses of a leaf to wounding have been shown to include changes in transpiration and CO2 assimilation rates because of closure of stomata (Peña-Cortés et al., 1995), rapid changes in leaf water potential and turgor pressure (Malone, 1992; Malone and Alarcon, 1995), the propagation of electrical signals associated with rapid changes in membrane potential (Peña-Cortés et al., 1995) and the release of oligosaccharide and peptides (Bowles, 1998). These signals initiate both local and systemic wound-signalling pathways transduced by jasmonic acid (JA), ethylene, nitric oxide and abscisic acid (ABA; Orozco-Cárdenas and Ryan, 2002; Orozoco-Cárdenas et al., 2001; Peña-Cortés et al., 1995; Rojo et al., 1999; Titarenko et al., 1997). In Arabidopsis thaliana, at least two wound signal transduction pathways, JA-dependent and -independent, have been described that are linked by their antagonistic responses to ethylene (Ellis and Turner, 2002; McConn et al., 1997; Nishiuchi et al., 1997; Rojo et al., 1998, 1999; Titarenko et al., 1997).

Hydrogen peroxide (H2O2), a reactive oxygen species (ROS), is also implicated in foliar wound responses (Orozco-Cárdenas and Ryan, 1999; Orozoco-Cárdenas et al., 2001). H2O2 may act as both a local and diffusible signalling molecule, activating defensive or acclimatory processes including excess light, ozone fumigation, heat stress and infection by pathogens (Alvarez et al., 1998; Bowler and Fluhr, 2000; Dat et al., 1998; Fryer et al., 2003; Karpinski et al., 1997, 1999; Pellinen et al., 1999; Tenhaken et al., 1995; Willekens et al., 1997). Pre-exposure of plants to this ROS can trigger protective functions that cause plants to acclimate to a range of stress conditions (Bowler and Fluhr, 2000; Karpinski et al., 1999; Kovtun et al., 2000).

The tissue-specific sources of H2O2 may be important in allowing discrimination between its roles (Mullineaux and Karpinski, 2002). In both local and systemic responses to wounding and excess light, H2O2 has been shown to accumulate in bundle sheath tissue of stressed leaves and this is associated with induction of defensive or acclimatory processes (Fryer et al., 2003; Karpinski et al., 1999; Orozco-Cárdenas and Ryan, 1999; Orozoco-Cárdenas et al., 2001). H2O2 has been suggested to mediate the ABA-regulated closure of stomata and could be either produced in guard cells for this function or diffuse in from other foliar locations (Murata et al., 2001; Pei et al., 2000; Zhang et al., 2001).

The subcellular origin of H2O2 may also be important in eliciting specific signalling pathways (Mullineaux and Karpinski, 2002). In response to pathogens and wounding, membrane-bound NADPH oxidase activity, cytosol-located oxidases and apoplastic peroxidases have been suggested to generate H2O2 (Allan and Fluhr, 1997; Bolwell et al., 1999, 2002; Orozoco-Cárdenas et al., 2001). In excess light-exposed leaves, an important source of ROS may be the Mehler reaction (Asada, 1999). The Mehler reaction is the reduction of O2 by electrons donated from photosystem I (PSI) and is one of several mechanisms for dissipating excess excitation energy (Asada, 1999). Furthermore, conditions, such as drought, chilling and infection by pathogens, that inhibit photosynthesis can result in excess excitation energy in the photosynthetic apparatus at low and moderate light intensities, which in more normal circumstances would not pose a problem to the plant (Long et al., 1994).

Ascorbate peroxidase (APX; EC 1.11.1.11) catalyses the reduction of H2O2 to water using ascorbate as the electron donor (Asada, 1999). The ASCORBATE PEROXIDASE 2 (APX2) gene encodes a cytosolic isoform of the enzyme, and its expression either is not detectable, or occurs only at extremely low levels in the absence of stress (Fryer et al., 2003; Karpinski et al., 1997, 1999; Panchuk et al., 2002; Santos et al., 1996). The induction of APX2 expression occurs at the level of de novo synthesis of transcript and is associated with foliar responses to excess light and heat stresses (Fryer et al., 2003; Karpinski et al., 1997, 1999; Panchuk et al., 2002). APX2 expression is limited to bundle sheath cells in leaves exposed to excess light (Fryer et al., 2003). The signals that initiate APX2 expression in excess light-stressed leaves are redox changes in photosynthetic electron transport (PET), H2O2 accumulation derived from Mehler reaction and a transient change in leaf water status (Fryer et al., 2003; Karpinski et al., 1999). The link between the induction of APX2 expression and leaf water status has been suggested to be mediated by ABA (Fryer et al., 2003).

Preliminary published observations showed that the expression of a fusion of the APX2 promoter to the firefly luciferase gene (APX2LUC) in transgenic Arabidopsis (Karpinski et al., 1999) was enhanced in the veins closest to the wounded tissue of detached leaves partially exposed to excess light (Mullineaux et al., 2000). This raised the possibility that the signalling pathway controlling the expression of APX2 could be integrated into one or more wound-signalling pathways or could share common components, or be responsive to the same effector molecules. However, the further investigations reported here led to a different and unexpected conclusion. We suggest that, in wounded leaves in the light, there is an increased diversion of photosynthetic electron flux to O2 in the vascular regions, leading to redox changes in PET and an increased production of H2O2 that, in turn, triggers induction of APX2 expression. This has important implications for how ‘wound-specific’ gene expression is viewed.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Wounding induces APX2 expression

The induction of APX2 expression in wounded leaves still attached to the plant could be detected by induction of expression of APX2LUC (Figure 1a; Karpinski et al., 1999). Luciferase activity could be imaged in the central vein and apical region of both the wounded leaf and adjacent unwounded leaves (Figure 1a). APX2 transcript could be detected as little as 45 min after wounding (Figure 1b). Under these conditions, the wounded plants expressed the JA-dependent wound-inducible VEGETATIVE STORAGE PROTEIN 1 (VSP1) and JA-independent wound-inducible BASIC CHITINASE (CHIB) genes (Figure 1c; Ellis and Turner, 2002; Rojo et al., 1999).

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Figure 1. Induction of APX2 expression in wounded leaves.

(a) Imaging of luciferase activity from an APX2LUC transgenic Arabidopsis line (Karpinski et al., 1999) 2 h after wounding (right panel). Note that there was no detectable activity immediately after wounding (left panel). The wounding was carried out by crimping with forceps at the points shown by red bands in the right panel. Wounded tissue can also be seen in the left panel at the start of the experiment as slightly darkened bands (arrowed). Plants were sprayed with 1 mm d (−)-luciferin and imaged using a Berthold Luminograph LB 980 CCD camera with an aperture setting of 1.8 (see Experimental procedures). The image of the rosette was taken using background luminescence detectable up to about 1 min after placing the plant under the camera. The light from the luciferase-catalysed reaction was imaged after 15 min in the dark and superimposed on the rosette image. The colour bar code gives the range of luciferase activity from background to 1000 RLU (blue) to 1800 relative light unit (RLU) (yellow).

(b) Effect of combined excess light and wounding on levels of APX2 transcript. Individual plants, undamaged or wounded as in (a), were subjected to a fivefold excess light treatment for one of the time points indicated, and their RNAs were extracted and analysed by blotting of RNA gels and probing with an APX2-specific cDNA (see Experimental procedures). High light-stressed (wounded and undamaged) samples were always transferred to the same membrane and probed simultaneously. The ACTIN probe was used as a loading control. The detection of APX2 mRNA from wounded leaves was not possible in RNA gel blots (data not shown). For early detection of APX2 transcript after wounding, 3′ RACE PCR was used to amplify cDNA followed by Southern blotting, gene-specific probes and detection using a phosphorimager (see Experimental procedures). Here, cDNA derived from APX3 transcript is used as a loading control and to check for presence of cDNA in early time points.

(c) Levels of VSP1 and CHIB transcripts in wounded, excess light treated, and combined treatments. The levels of transcripts were assayed using a quantitative RT-PCR procedure, normalized for loading using 18S RNA standard and expressed relative to untreated controls.

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APX2 expression in wounded leaves subjected to excess light

ASCORBATE PEROXIDASE 2 expression has been shown to be induced in leaves subjected to excess light treatments at times as little as 7 min after the onset of the challenge (Karpinski et al., 1997). To determine if the regulation of APX2 expression in wounded leaves shared any similarities to that in leaves subjected to excess light stress, the effect of these combined stresses on expression was compared with that in undamaged leaves subjected to a fivefold excess light stress. Using RNA gel blots, APX2 transcript was detected 15 min earlier and it accumulated to a >10-fold higher level in wounded, excess light-stressed leaves than in their undamaged counterparts (Figure 1b). It should be noted that, in wounded leaves, induction of APX2 transcript could only be detected by using reverse transcription-polymerase chain reaction (RT-PCR) and was undetectable on RNA gel blots (data not shown and Figure 1b). Therefore, it was concluded that the combined effect of wounding and excess light had a synergistic effect on the level of induced APX2 transcript, compared with either excess light or wounding stress alone. Both VSP1 and CHIB expression were induced by excess light treatment, but no synergistic effect could be discerned of the combined stresses on their transcript levels (Figure 1c).

Treatment of detached illuminated APX2LUC leaves with the PET inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) abolished wound-induced luciferase activity (Figure 2). These observations confirmed the requirement for PET for wound-induced APX2 expression.

image

Figure 2. The inhibition of wound-induced APX2LUC expression by the photosynthesis inhibitor DCMU.

Leaves were vaccum infiltrated for 3 min with 10 µm DCMU and then incubated in the same solution for 3 h. At the end of this period, the leaves were wounded with one crimping action across the middle of the leaf, incubated on wetted blotting paper at growth PPFD for 2 h and sprayed with luciferin, and the luciferase activity was imaged as described in the legend of Figure 1(a). As controls, water-infiltrated leaves were wounded in the same way. As a further positive control of DCMU action, the expression of luciferase activity in an excess light-stressed leaf (10-fold; 40 min) and its inhibition by the DCMU treatment are shown and were as reported previously by Karpinski et al. (1999). The colour bar code gives the range of luciferase activity from background to 1000 RLU (blue) to 1800 RLU (yellow).

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Wound- and excess light-induced APX2 expression is not controlled by known wound-signalling pathways in Arabidopsis

Arabidopsis has at least two wound-signalling pathways: JA-dependent and JA-independent (Ellis and Turner, 2002; Rojo et al., 1999; Titarenko et al., 1997). Communication between these pathways is mediated by ethylene generated in damaged tissues (Rojo et al., 1999). The mutant coronatine-insensitive 1-1 (coi1-1; Feys et al., 1994) is blocked in the perception of JA, and COI1 may play a role in cross-talk between the JA pathway and those directed by ethylene and ABA (Ellis and Turner, 2002; Feys et al., 1994). The coi1-1 mutant showed no difference in the abundance of APX2 transcript in wild-type plants when wounded or subjected to excess light (Figure 3a). VSP1 transcript induction was blocked in wounded, but not in high light-stressed coi1-1 plants (Figure 3b). Similarly, wounded or high light-treated mutants impaired in their perception of ethylene, ethylene insensitive 2-1 (ein2-1; Guzman and Ecker, 1990) and ethylene resistant 1-3 (etr1-3; Bleecker et al., 1988), or JA, jasmonic acid resistant 1-1 (jar1-1; Staswick et al., 1992), showed no inhibition of APX2 expression (data not shown).

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Figure 3. Transcript levels of VSP1 and APX2 in transgenic and mutant plants subjected to wounding or exposure to high light.

(a) Quantitative RT-PCR of APX2 cDNA prepared from pooled RNA from six untreated (tracks 1 and 2), wounded (tracks 3 and 4) or high light-stressed (tracks 5 and 6) plants. Mutant coi1-1 individuals (tracks 1, 3 and 5) were compared with phenotypically wild-type plants (tracks 2, 4 and 6). This was achieved by carrying out the experiments on all 4–5-week-old individuals that segregated for the coi1-1 mutation and had come from a self-pollinated coi1-1/COI1 parent (Ellis and Turner, 2002). Plants were sampled for leaf material 2 h after wounding, and their RNA were extracted (see Experimental procedures). At the end of the experiment, the plants were placed under 18-h photoperiod conditions to induce bolt formation, and flowers from the plants were scored for fertility (wild-type or coi1-1 heterozygous) or infertile (coi1-1 homozygous; Ellis and Turner, 2002; Feys et al., 1994). After being scored, RNA from individuals was pooled into a coi1-1 or wild-type phenotype samples. The 18S rRNA was used as an internal control. The picture shows a single experiment typical of two independent experiments and sets of RNA isolations.

(b) Quantitative RT-PCR detection of VSP1 transcript. PCR of APX3 transcript was used here as non-responsive control to wounding. The detection of these transcripts was from the same RNA samples as described in (a).

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APX2LUC Arabidopsis rosettes sprayed with either JA or chitosan, an elicitor of the JA-independent pathway (Rojo et al., 1999), did not induce luciferase activity under ambient light conditions (Table 1). The effects of JA and chitosan in the leaves were confirmed by CHIB and VSP1 gene induction (Table 1). Taken together, these data indicated that the wound induction of APX2 expression was not regulated by either of the known JA-dependent or -independent pathways in Arabidopsis and, in contrast to wounding, VSP1 induction of expression in excess light-stressed leaves also was not JA dependent.

Table 1.  Jasmonic acid and chitosan have no effect on APX2LUC expression
TreatmentRLU g−1 FW (±SD)VSP1 transcript level (relative units)CHIB transcript level (relative units)
  1. In vitro luciferase activity (RLU g−1 FW) and relative transcript levels for VSP1 and CHIB from 4-week-old, short-day-grown APX2LUC transgenic plants, sprayed with 50 µm JA or chitosan (250 µg ml−1) until run-off was observed, on leaves incubated for 2 h. Controls were water-sprayed plants or wounded as in legend of Figure 1. Transcript levels were determined using a quantitative RT-PCR procedure (see Experimental procedures). Values are means of three independent experiments (n = 3 for each treatment).

Control18700 (±4500)11
Wounded92900 (±22460)1.9 (±0.3)3.4 (±0.6)
JA9790 (±783)2.5 (±0.4)6.1 (±0.8)
Chitosan5020 (±552)0.7 (±0.3)3.0 (±0.5)

PET in wounded leaves

The induction of APX2 expression in excess light-stressed leaves requires PET and H2O2 (Fryer et al., 2003; Karpinski et al., 1997, 1999). Therefore, it was considered that electron transport may also be affected in wounded leaves in such a way as to lead to APX2 induction similar to that in excess light-stressed leaves. A large decrease in the operating efficiency of PSII was imaged within 5 min at the wounded sites in leaves (Figure 4a), indicating that PET was rapidly inhibited. Over a 6-h period after wounding, a partial recovery in overall PSII efficiency occurred (Figure 4b). By this time, APX2LUC expression (as luciferase activity) had been induced (Figure 4b). Treatment of leaves with JA or chitosan had no significant effects on PET, as determined by the PSII operating efficiency (Fm′ − F′/Fm′ was 0.49 (±0.04), 0.44 (±0.05) and 0.46 (±0.03) for leaves (n = 6) treated for 2 h with water, JA or chitosan, respectively).

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Figure 4. PET in wounded leaves.

(a) Spatial and temporal monitoring of operating efficiency of PSII electron transport ((Fm′ − F′)/Fm′; Barbagallo et al. (2003) in wounded leaves were obtained by imaging chlorophyll a fluorescence parameters (see Experimental procedures). The wound sites can clearly be seen in damaged halves of leaves as having much lower values for (Fm′ − F′)/Fm′. The range of (Fm′ − F′)/Fm′ values was mapped to a colour palette as shown. The distribution of values of this parameter within this parameter range is also shown.

(b) Averaged values of operating efficiency of PSII electron transport ((Fm′ − F′)/Fm′) were calculated from chlorophyll a fluorescence measurements using a modulating fluorimeter in which the detector was placed over crimp wounded tissue (see Experimental procedures) at the times indicated (solid line). Wounded and control undamaged plants were maintained at their growth PPFD throughout the experiment. Data are the means of measurements from a single wounded leaf (as in Figure 1a) from each of eight plants for each time point from a total of three independent experiments. SEs are shown. Luciferase activity (dotted line) from expression of APX2LUC in a similar experiment is shown for ease of reference to expression data elsewhere in the paper.

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H2O2 and stomatal responses in wounded leaves

A further criterion for the induction of APX2 expression in the bundle sheath tissue of excess light-stressed leaves is a rapid and transient change in leaf water status (Fryer et al., 2003). As rapid changes in both leaf water potential and turgor are known to occur in response to leaf wounding (see Introduction), it was reasoned, as in excess light-stressed leaves, that this may also be a cue for the induction of APX2 expression in wounded leaves.

Changes in transpiration rate and stomatal conductance of the leaves were measured following wounding (Figure 5a). At suitable time points (points 1, 2 and 3 in Figure 5a), leaves were removed from the gas exchange chamber and luciferase activity (Figure 5b) was imaged, and then the leaves were de-stained to reveal H2O2 accumulation as insoluble 3,3′-diaminobenzidine (DAB) precipitate (Figure 5c; see Experimental procedures). In wounded leaves, stomatal conductance and transpiration declined within 15 min of leaf wounding, reaching a minimum after 20 min, whereas control leaves exhibited an increase in both stomatal conductance and transpiration during this period (Figure 5a). The decreases in stomatal conductance in wounded leaves were coincident with a decrease in CO2 assimilation, the detection of luciferase activity (Figure 5b) and accumulation of brown DAB stain in veinal tissue indicative of increased levels of H2O2 in these tissues (Figure 5c). Leaves attached to the plant showed the same pattern of stomatal conductance as shown in Figure 5(a) (data not shown).

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Figure 5. APX2 expression and H2O2 in wounded leaves is preceded by a change in leaf water status and CO2 fixation.

(a) Changes in transpiration rate, stomatal conductance and CO2 fixation of leaves wounded by one crimping action (see reflected light image below). The leaves were kept at growth PPFD (250 µmol m−2 sec−1) and an atmospheric vapour pressure of 0.5 kPa. Data are the means of five leaves; SEMs were <15%. Wounded and control leaves were infiltrated with luciferin and DAB before the start of the experiment.

(b) Development of luciferase activity after wounding leaves with a single crimping action. The images were taken of leaves at 0, 30 and 60 min after wounding (sampling points 1, 2 and 3 in (a)) using a Peltier cooled CCD camera (see Experimental procedures). The predominantly blue colour represents the lowest detectable values for luciferase with this camera system. A reflected light image of a leaf is shown to the left of the images of luciferase activity.

(c) Staining of H2O2 in the region of damage in the wounded leaf using DAB. Note the appearance of new staining around the damaged area of the leaf. The staining in the mid-vein may be associated with wounding after detachment from the plant (Fryer et al., 2003).

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The involvement of leaf water status in the wounding-mediated induction of APX2 expression was further investigated using the ABA-insensitive mutants abscisic acid insensitive (abi)1-1 and abi2-1. These mutants are impaired in ABA-induced stomatal closure and in the ABA-directed production and perception of H2O2 in guard cells, respectively (Koornneef et al., 1984; Leung et al., 1994; Meyer et al., 1994; Murata et al., 2001).

Both mutants induced APX2 expression in response to wounding at a level equivalent to that of wild-type controls (Figure 6). Wounded leaves of both mutants exhibited a wilting phenotype, implying that stomata failed to close in response to this challenge. It was not possible for us to obtain reliable stomatal conductance data from these mutants because of foliar damage incurred on placing the leaves in the porometer. Also, the changing position of mutants' leaves as they wilted in both the gas exchange chamber and in the open laboratory resulted in changes in light absorption and gas exchange, which made interpreting leaf physiological responses to wounding problematic (data not shown).

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Figure 6. Steady-state levels of APX2 transcript in wounded leaves (as in Figure 1a) of the wild type (Accession Landsberg erecta) and mutants abi1-1 and abi2-1.

Each RNA sample was pooled from three experiments using four plants that had been wounded 2 h previously. After 3′ RACE PCR, blotting and probing with a specific APX2 probe, the hybridized filter was placed on a phosphorimager to visualize the bands. The degree of hybridization was calculated by subtracting pixel density at each band from background using the densitometry software provided on the phosphorimager. The data are presented relative to cDNA levels from the wild-type wounded plant. APX3 transcript levels were used as a known non-responsive control to indicate variation in loading (as in Figure 1b). The data used to calculate relative values had SEs of <21%

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Sources of H2O2 for induction of APX2 expression

It has been proposed that NADPH oxidase(s) could be responsible for the generation of H2O2 in the peri-veinal tissue of wounded leaves (see Introduction). This proposal comes in part from the observation that the wound-induced accumulation of H2O2 can be inhibited by pre-treatment of leaves with diphenyl iodonium (DPI; Orozco-Cárdenas and Ryan, 1999; Orozoco-Cárdenas et al., 2001). Therefore, the effect of DPI was tested on the wound-induced APX2 expression and on PET.

DPI inhibited the wound-induced expression of APX2LUC in fully expanded leaves by >80% after 6 h (Figure 7a). This inhibition of APX2LUC expression by DPI was associated with a decline in the PSII operating efficiency indicative of decreases in the rate of PET (Figure 7b). These data suggest that DPI is a potent inhibitor of PET in wounded leaves. This conclusion was further supported by the inhibition of the rate of photosynthetic O2 evolution over a range of photosynthetically active photon flux (PPFDs) in DPI-treated leaves (Figure 7c). Furthermore, the rate of O2 evolution in DPI-treated leaves saturated below a PPFD of 600 µmol m−2 sec−1 in DPI-treated leaves, whereas control leaves required in excess of 800 µmol m−2 sec−1 for saturation.

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Figure 7. Effect of DPI on the induction of APX2LUC expression (a), operating efficiency of PSII in wounded leaves (b) and photosynthetic O2 evolution (c).

(a, b) DPI (50 µm) or water was injected into the mid-vein of leaves while still attached to the plant 1 h before wounding (as in Figure 1a) at time zero. Three plants per time point were used with a single wounded leaf taken from each rosette. The data shown are means (±SE) combined from two experiments. Duplicate assays for luciferase activity (see Experimental procedures) were performed from cell-free extracts prepared from the wounded leaves from which the operating efficiency of PSII, estimated from (Fm′ − F′)/Fm′, had been determined as described in the legend of Figure 3 and Experimental procedures.

(c) The rate of photosynthetic O2 evolution in leaf discs (Karpinski et al., 1997) prepared from DPI-treated leaves (after 4 h incubation) of APX2LUC plants was compared with that in control water-treated leaves at a range of PPFDs. The data are the means (±SE) of three independent experiments, each conducted with a single fully expanded leaf from four plants for each treatment (n = 12). The differences observed between DPI- and water-treated leaves are statistically significant (P < 0.001).

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Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The observation that wounded leaf tissue showed enhanced expression of APX2 under ambient and excess light conditions (Figure 1) raised the possibility that wound-signalling pathways in Arabidopsis might share common components with excess light-signalling pathways (Mullineaux et al., 2000). However, the evidence presented here indicates that neither the JA-dependent nor -independent wound-signalling pathways known in Arabidopsis (Leon et al., 2001; McConn et al., 1997; Nishiuchi et al., 1997; Rojo et al., 1998, 1999; Titarenko et al., 1997) are implicated in APX2 expression in wounded leaves (see Results). This is despite both pathways being operative in our experimental conditions, as evidenced by the induction of VSP1 and CHIB expression, respectively (Figure 1b). While not examined in detail, it was interesting to note that high light-mediated induction of VSP1 expression was not JA dependent (Figures 1b and 3b), adding further weight to the argument that the JA-signalling pathway plays no direct role in governing responses of gene expression to high light stress.

The highest levels of APX2 expression were always observed in leaves subjected to both wounding and excess light (Figure 1c). Wounded leaves treated with the photosynthetic inhibitor DCMU did not show any induction of APX2 expression (Figure 2). This led us to hypothesize that the mechanism of APX2 induction in wounded Arabidopsis leaves was qualitatively the same as that described previously for excess light-stressed leaves, depending on redox changes in PET that generate H2O2 and augmented by ABA (Fryer et al., 2003; Karpinski et al., 1999). This hypothesis would mean that induction of APX2 expression would differ between wounded and excess light-stressed leaves only in the degree of response. The inhibition of wound-induced APX2 expression by DCMU (Figure 2) is consistent with this hypothesis. PET has been shown previously to impact on the expression of some, but not all, genes associated with wound responses in plants. For example, the induction of a VSP gene in soybean by JA was shown to be inhibited by DCMU (Sadka et al., 1994), whereas the wound induction of proteinase inhibitor 2 (pin2) expression in potato leaves was shown to be light dependent but not inhibited by DCMU (Peña-Cortés et al., 1992, 1995).

While this hypothesis was a useful framework for further experimentation, it should be noted that the synergistic effect of excess light and wounding on APX2 expression (Figure 1b) might suggest that unknown factors could be influencing expression of the gene when multiple stresses challenge the plant. However, these factors are not likely to involve the other known wound-responsive pathways as they have no impact on APX2 expression (Table 1).

A key point to establish in this study was whether or not PET undergoes a change in activity in wounded leaves consistent with its association with the induction of APX2 expression (Fryer et al., 2003; Karpinski et al., 1997, 1999). The operating efficiency of PSII had declined markedly in the vicinity of wound sites within 5 min (Figure 4a) after damage and then began a slow, but incomplete, recovery over the following 6 h (Figure 4b). This implied that PET in wounded leaves, while remaining at a lower rate compared with undamaged leaves, was increasingly supplying electrons to metabolic sinks in the hours following leaf damage. In excess light-challenged leaves, one of the important components for the induction of APX2 expression is the accumulation of H2O2 in bundle sheath tissue derived from PET but not photorespiration (Fryer et al., 2003; Karpinski et al., 1997, 1999). In wounded leaves, the timing of the decline and recovery of PET, the accumulation of H2O2 and APX2LUC expression in the bundle sheath tissues is consistent with this explanation (Figures 4 and 5). In high light-stressed leaves, the expression of APX2, the accumulation of H2O2 and the changes in PET have been shown to occur in bundle sheath cells as part of acclimation to high light (Fryer et al., 2003; Karpinski et al., 1999). Equally, in wounded leaves, bundle sheath cells have been shown to accumulate H2O2, which has been suggested to be a secondary messenger for the induction of a class of wound-responsive defence genes (Orozco-Cárdenas and Ryan, 1999; Orozoco-Cárdenas et al., 2001). Our observations confirm that wounded Arabidopsis leaves do accumulate H2O2 in veinal tissue (Figure 5c). It has been shown previously that direct treatment of leaves with H2O2, in ambient light conditions, can induce APX2 expression to about 10% of that caused by high light (Karpinski et al., 1999). Therefore, we conclude that H2O2 accumulation in the chloroplast may be a signal for APX2 induction in damaged leaves in the light.

JA has been proposed to mediate the accumulation of H2O2 in wounded tomato leaves (Orozco-Cárdenas and Ryan, 1999; Orozoco-Cárdenas et al., 2001). In our experiments, no evidence could be found for JA involvement in the wounding induction of APX2, which would indicate an accumulation of H2O2 (see above). Furthermore, JA-treated leaves showed no perturbation in PET (see Results), which might indicate that, despite there being a JA-stimulated accumulation of H2O2 in such plants (Orozco-Cárdenas and Ryan, 1999), by itself this was not sufficient to induce APX2 expression. Thus, for APX2 induction in wounded leaves, additional signals may be required such as the accompanying changes in PET. Supporting this conclusion is that, in contrast to JA, H2O2 treatment of leaves does incur changes in PET consistent with the latter's role in the regulation of APX2 expression (Karpinski et al., 1999). Therefore, if JA triggers accumulation of H2O2 in wounded Arabidopsis leaves in a manner similar to that reported in tomato leaves (Orozco-Cárdenas and Ryan, 1999), it does not achieve this by influencing PET.

In wounded tomato leaves, the accumulation of H2O2 in bundle sheath cells is inhibited by DPI (Orozco-Cárdenas and Ryan, 1999; Orozoco-Cárdenas et al., 2001). DPI is an inhibitor of NADPH oxidases and other flavoenzymes (Cross and Jones, 1986; Moulton et al., 2000; O'Donnell et al., 1993). This observation has led to the proposal that wound-induced H2O2 accumulation may be catalysed by a bundle sheath-located NADPH oxidase (Orozoco-Cárdenas et al., 2001). Similar conclusions about the origin of H2O2 have been made from DPI treatments of stomatal guard cells, wounded fruit mesocarp tissue and elicitor-treated suspension culture cells (Levine et al., 1994; Murata et al., 2001; Tenhaken et al., 1995; Watanabe and Sakai, 1998; Zhang et al., 2001). There is considerable biochemical and molecular genetic evidence for the existence of at least the gp96phox subunit of NADPH oxidases in plants, homologous to that in mammalian phagocytes (Keller et al., 1998; Tenhaken et al., 1995; Torres et al., 1998). However, DPI has been shown to inhibit other haem-containing enzymes such as xanthine oxidase, nitric oxide synthase (Delledonne et al., 1998; Moulton et al., 2000) and cell wall peroxidases (Bolwell et al., 1999). Some of these enzymes have been implicated as sources of ROS at least for the hypersensitive response (Allan and Fluhr, 1997; Bolwell et al., 1999, 2002). Furthermore, DPI and other iodonium compounds are potent inhibitors of mitochondrial electron transport (Cross and Jones, 1986; Ellis et al., 1989), and mitochondrial-derived ROS have been implicated in controlling gene expression at least in ozone-fumigated birch leaves (Pellinen et al., 1999).

The data presented here indicate that the effects of DPI may include inhibition of photosynthesis, complicating the interpretation of data when using this compound on leaves. DPI inhibited wound induction of APX2LUC expression by >80% (Figure 7a), but this was accompanied by an inhibition of PET (Figure 7b,c). There is considerable evidence for the requirement of PET for expression of APX2 (Figure 2; Fryer et al., 2003; Karpinski et al., 1997, 1999). Therefore, we favour the inhibition of PET as the explanation for the action of DPI on APX2 expression rather than implying a direct role for an NADPH oxidase.

Hydraulic signals, such as changes in turgor, have been demonstrated to appear very rapidly upon wounding in many plant species (Malone, 1992; Malone and Alarcon, 1995). In high light-stressed leaves, a transient increase in the rate of water loss prior to stomatal closure is also another physiological cue that is required for identification of APX2 and it has been proposed that ABA is required for maximum APX2 expression in high light-stressed leaves (Fryer et al., 2003). From these considerations, it was possible that same parameters could be required in the wound induction of APX2. Furthermore, ABA has been implicated in the regulation of some wound-inducible genes such as PROTEINASE INHIBITOR 1 from tomato and potato (Peña-Cortés et al., 1995).

In wounded Arabidopsis leaves, no transient increase in transpiration rate occurred that might implicate a decline in leaf water status (Figure 5a). Instead, the leaves immediately reduced transpiration, gas exchange and decreased stomatal conductance (Figure 5a). Despite the influence of ABA on the expression of APX2 (Fryer et al., 2003), abi mutants showed no inhibition of wound-inducible APX2 expression (Figure 6). Therefore, it was concluded that the wound-mediated induction of APX2 does not require ABA. The reduced levels of APX2 induction in wounded leaves compared with that in high light-stressed leaves (Figure 1) may be a consequence of only a partial signal derived from photosynthesis and H2O2 alone, with no augmentation of expression by ABA or a water-related signal.

The lack of impact of the abi mutations on wound-induced APX2 expression also indicated that stomatal closure plays no discernable role in this process, despite the observed closure of stomata in response to wounding (Figure 5a). Lowered rates of gas exchange as a consequence of closed or restricted stomata and the resulting depletion in internal leaf CO2 concentration are often invoked to explain an increase in production of ROS by photorespiration or the Mehler reaction (Long et al., 1994; Ort and Baker, 2002). Clearly, while this is an important physiological response of the leaf to wounding, it cannot be a critical factor influencing the wounding induction of APX2 expression. In support of this conclusion, in the abi2-1 mutant, H2O2 accumulation in high light-stressed-leaves is not inhibited, implying that closure of stomata is not critical for the accumulation of ROS in bundle sheath cells (Fryer et al., 2003).

The signal transduction pathway, which includes H2O2 from the chloroplast and achieves regulation of APX2 expression in bundle sheath cells of both high light-stressed and wounded Arabidopsis leaves, remains to be determined (Mullineaux and Karpinski, 2002). However, it is clear that the signalling role of ROS derived from changes in PET may be a common feature in controlling the expression of genes in response to environmental stimuli, such as in wounded leaves described here. Consequently, it may not be necessary to invoke a complex network of stimulus-specific or novel signal transduction pathways in all cases.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Plant material

Arabidopsis thaliana L. plants were germinated and grown under controlled environmental conditions (PPFD of 250 µmol m−2 sec−1 during a 10-h photoperiod at 22°C in a relative humidity of 70%) for 4–5 weeks. In all experiments, unless otherwise stated, the Col-0 ecotype of A. thaliana was used. The APX2LUC transformant (Col-0) has been described previously by Karpinski et al. (1999). The mutants etr1-3, ein2-1, jar1-1, abi1-1 and abi2-1 were obtained from the Nottingham Arabidopsis Stock Centre (NASC, Nottingham, UK). The coi1-1 mutant was a kind gift from Prof. John Turner (University of East Anglia, UK). The abi and coi1-1 mutations were verified using a PCR-based test and a fertility test, respectively (Ellis and Turner, 2002; Feys et al., 1994; Murata et al., 2001).

Treatments

Leaves were wounded using blunt-end forceps. Pressure was applied so that the leaves were damaged but not to the extent that the lamina was pierced through. Unless otherwise stated, each leaf was wounded three times at the base, apex and middle across the entire width of the leaf (see Figure 1a). Plants were subjected to excess light, fivefold above growth PPFD, as described previously by Karpinski et al. (1997, 1999).

To determine the effect of JA, chitosan, DCMU or DPI on APX2LUC expression and PET, whole rosettes (n = 3 for each treatment time point unless stated otherwise) were sprayed with JA (50 µm; Sigma, St. Louis, USA) or chitosan (250 µg ml−1) until the leaves dripped wet. JA and chitosan (Sigma) were prepared as described by Rojo et al. (1999) and Hadwiger and Beckman (1980), respectively. Control plants were sprayed with water and compared with wounded plants. In the JA and chitosan treatments, the treated rosettes were kept under their growth conditions for 2 h prior to determination of in vitro luciferase activity. To determine the effects of JA and chitosan on photosynthesis, chlorophyll a fluorescence measurements were made on fully expanded leaves at 0 and 2 h post-treatment.

DPI chloride was dissolved in water to 50 µm and was injected under the epidermis of the mid-vein of leaves still attached to their rosettes. The leaves were left for 1 h prior to wounding. It should be noted that this treatment and the control injection of water had a delaying effect on the wound induction of APX2LUC expression compared with wounded untreated leaves.

DCMU was made as a 10 mm stock in 50% (v/v) ethanol and diluted to 10 µm in water. This solution was vacuum infiltrated into leaves for 3 min and then floated on the same DCMU concentration for 3 h prior to wounding or excess light treatments as described by Karpinski et al. (1999).

Imaging of luciferase activity and H2O2 accumulation

The expression of APX2LUC in leaves was imaged after spraying with 1 mm d (–)-luciferin (Promega, USA). The imaging of luciferase activity (Figure 1a) using a Berthold Luminograph (Berthold, Germany) LB 980 charge-coupled device (CCD) camera was performed as described by Karpinski et al. (1999), while the sharper images of luciferase activity (Figure 5b) were collected with a Peltier-cooled CCD camera as described by Fryer et al. (2003). The visual detection of H2O2 accumulation in detached leaves loaded with DAB (Sigma) (Thordal-Christiansen et al., 1997) was carried out as described by Fryer et al. (2002).

Measurement of luciferase activity in vitro

This was carried out as described previously using an in vitro luciferase assay kit (Promega, WI, USA; Karpinski et al., 1999), except that a Biolumat LB950 luminometer (Berthold, Germany) was used, which will account for values for APX2LUC-derived luciferase activity higher than those previously described by Karpinski et al. (1999).

Fluorescence measurements

Chlorophyll a fluorescence measurements from leaves were made using a modulating fluorimeter FMS1 (Hansatech, Kings Lynn, UK). The operating efficiency of PSII photochemistry was determined from (Fm′ − F′)/Fm′, where Fm′ is the maximal fluorescence level in the light-adapted state and F′ is the fluorescence level immediately prior to measuring Fm′ (Baker et al., 2001). Images of chlorophyll fluorescence parameters were obtained using a FluorImager chlorophyll fluorescence imaging system (Technologica Ltd., Colchester, UK), exactly as described by Barbagallo et al. (2003).

Gas exchange measurements and stomatal conductance

Detached leaves were wounded only once with forceps (see Figure 5), and the top half of each leaf, including the wounded section, was immediately enclosed in the chamber of an infrared gas analysis system (CIRAS; PP Systems, Hitchin, UK). In the chamber, the temperature was maintained at 25°C, the water vapour pressure at c. 0.5 kPa and airflow at 350 cm3 min−1. The bottom half of the leaf was kept at room temperature (23°C) in the dark. The top half of each leaf in the chamber was illuminated using a 75 W xenon lamp at growth PPFD (Leica UK Ltd., Milton Keynes, UK). Measurements of stomatal conductance, transpiration rate and CO2 assimilation of leaves were made using the CIRAS.

Photosynthetic O2 evolution

Oxygen exchange rates were measured in gas phase using a Clark-type oxygen electrode (LD2/3 oxygen electrode chamber) connected to an Oxylab control unit (Hansatech Instruments Ltd., Norfolk, UK) and recorded on-line with a computer. The leaves were illuminated with consecutive light pulses of increasing light intensity provided by an liquid electronic display (LED) light source (Hansatech LH36-2), and photosynthesis versus irradiance curves were calculated from the linear increase/decrease of oxygen during light/dark pulses using MS Excel.

Preparation and analysis of RNA

Total RNA was isolated from leaves using a Qiagen Plant RNeasy Kit (Qiagen, Crawley, UK) and quantified as described in the manufacturer's instructions. A minimum of three RNA preparations each from a separate plant was made per time point. Each experiment was repeated at least three times, except where indicated differently in the figure legends. The figures show typical data from a single experiment.

The analysis of APX2 transcript levels using RNA gel blots and an APX2-specific probe has been described previously by Karpinski et al. (1997), except that radioactive bands on the nylon membranes were detected using a phosphorimager (BAS 1000; Fuji Photo Film Co. Ltd., Japan). The pea ACTIN probe used to check RNA loadings was recovered by PCR (see below) from a full-length cDNA inserted in the plasmid pBluescript SK+ (European Molecular Biology Laboratory (EMBL)/GenBank No. X67666) using the T3 and T7 primers, 5′-AATTAACCCTCACTAAAGGG-3′ and 5′-GTAATACGACTCACTATAGGGC-3′, respectively.

PCR procedures

Gene-specific probes were made by amplifying either inserts in plasmids (100–500 pg) or cDNA (from the equivalent of 500 ng of total RNA prepared from wounded plants) according to standard protocols for both PCR and RT-PCR conditions (Innis et al., 1990).

The cDNA for RT-PCR or 3′ rapid amplification of cDNA ends (RACE; Innis et al., 1990) PCR was synthesized using 3 µg of total RNA and Superscript™· reverse transcriptase according to the manufacturer's instructions (Invitrogen, Paisley, UK) and as described previously by Karpinski et al. (1997). All cDNAs to be used as probes were verified as correct by cloning into plasmid pCR2.1 (http://www.invitrogen.com) and sequencing of the insert. All primers used for PCR in this study were specific for the coding region of the target gene. For the 3′ RACE PCR procedure to amplify cDNA of transcripts from wounded tissue, the same forward sense strand primer was used in conjunction with the non-specific 3′ AMP primer (Innis et al., 1990). For some studies as indicated, a quantitative PCR procedure kit was used according to the manufacturer's instructions (Ambion Ltd., Huntingdon, UK). The APX2- and APX3-specific primers have been described previously by Karpinski et al. (1997). The sequences of the CHIB- and VSP1-specific primers were obtained from Ellis and Turner (2002) and Rojo et al. (1999), respectively. 3′ RACE PCR products were separated on agarose gels by electrophoresis, blotted on nitrocellulose and hybridized with gene-specific probes as previously described by Karpinski et al. (1997), except that a phosphorimager was used to visualize the radioactive bands as described for RNA gel blots.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The authors are grateful to Mrs Helen Reynolds for skilled technical assistance. Grants from the Biotechnology and Biological Sciences Research Council (BBSRC) to N.R.B. and P.M.M., the BBSRC Core Strategic Grant to the John Innes Centre (P.M.M.), the Swedish Council for International Co-operation in Research and Higher Education (S.K. and P.M.M.) and the Swedish Science Foundation (S.K.) supported this work. C.C.-C.C. and L.B. gratefully acknowledge research studentships from Stockholm University and the BBSRC, respectively.

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  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
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