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Journal of Experimental Botany, Vol. 54, No. 392, pp. 2529-2540, November 1, 2003
© 2003 Oxford University Press

Early production and scavenging of hydrogen peroxide in the apoplast of sunflower plants exposed to ozone

Received 11 February 2003; Accepted 17 July 2003

A. Ranieri*,1, A. Castagna1, J. Pacini1, B. Baldan2, A. Mensuali Sodi3 and G. F. Soldatini1

1 Department of Agricultural Chemistry and Biotechnology, University of Pisa, Via del Borghetto 80, I-56124 Pisa, Italy
2 Department of Biology, University of Padova, Padova, I-35131, Italy
3 Sant’Anna School of University Studies and Doctoral Research, Pisa, I-56124, Italy

* To whom correspondence should be addressed. Fax: +39 50 598614. E-mail: aranieri{at}agr.unipi.it
Abbreviations: APX, ascorbate peroxidase; CAB, Na-cacodylate buffer; CB, cell wall covalently-bound fraction; DPI, diphenylene iodonium; DTT, dithiothreitol; HEPES, N-2-hydroxyethylpiperazine-N'-2-ethanesulphonic acid; IB, cell wall ionically-bound fraction; IWF, intercellular washing fluid; MOPS, 3-(N-morpholino)propanesulphonic acid; NAD(P)H PODs, H2O2 producing NAD(P)H-oxidizing peroxidases; PM, plasma membrane; PODs, peroxidases; pCMB, p-chloromercuribenzoate; ROS, reactive oxygen species; Syr-PODs, H2O2 scavenging syringaldazine-reducing peroxidases; TBARS, thiobarbituric acid reactive substances.


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 References
 
The present work set out to define the processes involved in the early O3-induced H2O2 accumulation in sunflower plants exposed to a single pulse of 150 ppb of O3 for 4 h. Hydrogen peroxide accumulation only occurred in the apoplast and this temporally coincided with the fumigation period. The inhibitor experiments suggested that both the plasma membrane-bound NAD(P)H oxidase complex and cell-wall NAD(P)H PODs contributed to H2O2 generation. To investigate the mechanisms responsible for O3-induced H2O2 accumulation further, both production and scavenging of H2O2 were investigated in the extracellular matrix after subcellular fractionation. The results indicated that H2O2 accumulation is a complex and highly regulated event requiring the time-dependent stimulation and down-regulation of differently located enzymes, some of which are involved in H2O2 generation and degradation, not only during the fumigation period but also in the subsequent recovery period in non-polluted air. Owing to the possible interplay between H2O2 and ethylene, the time-course of ethylene emission was analysed too. Ethylene was rapidly emitted following O3 exposure, but it declined to control values as early as after 4 h of exposure. The early contemporaneous detection of increased ethylene and H2O2 levels after 30 min of exposure does not allow a clear temporal relationship between these two signalling molecules to be established.

Key words: Ascorbate peroxidase, ethylene, hydrogen peroxide, NAD(P)H oxidase, ozone, peroxidases, reactive oxygen species, signal transduction, sunflower (Helianthus annuus L.).


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 References
 
Among the major gaseous pollutants studied to date, tropospheric ozone (O3) causes more damage to both natural and cultivated plants in industrialized nations than any other pollutant (Heagle, 1989; Kley et al., 1999; Krupa and Kickert, 1989).

O3 toxicity was previously believed to be mainly due to the loss of membrane integrity and the formation of toxic and oxidative products by O3 degradation (Heath, 1994). Recently, however, the similarities between O3-induced responses and the hypersensitive responses found in incompatible plant–pathogen interaction have led to the suggestion that O3 could act as an abiotic elicitor of phytopathological and antioxidant responses (Sandermann et al., 1998; Schraudner et al., 1998; Sharma and Davis, 1997). The similarity between O3 and pathogen attacks is most probably related to the occurrence of reactive oxygen species (ROS) such as the superoxide anion radical O.2– and hydrogen peroxide (H2O2) in the apoplast, the so-called ‘oxidative burst’. In fact, once having penetrated the leaf apoplast, O3 is rapidly converted to ROS. In addition to this rapid spontaneous ROS generation from O3 decomposition and/or from reaction with components of the plasma membrane and cell wall (Laisk et al., 1989; Salter and Hewitt, 1992), an active endogenous ROS production by cell enzymatic systems, such as the PM-bound neutrophil-like NAD(P)H oxidase complex, may occur (Bolwell and Wojtaszek, 1997). Peroxidases (PODs), which are important components of plant responses to different kinds of stresses, may also regulate the level of ROS. In fact, besides their role in the H2O2-dependent polymerization of lignin precursors and the cross-linking between cell-wall proteins and polysaccharides (Christensen et al., 1998; Otter and Polle, 1997; Penel et al., 2000; Polle et al., 1994), PODs are also capable of H2O2 generation through the oxidation of various mole cules, including NAD(P)H (Bolwell and Wojtaszek, 1997).

Among the different ROS, directly or indirectly generated following O3 exposure, H2O2 is receiving increasing attention because of its dual role as a signal molecule and as a toxic-compound mediator of oxidative damage. In fact, due to its relatively long life and capability to cross the membranes, H2O2 can diffuse into the cell and/or move to the neighbouring cells, thus acting as a short-distance signal or it can react with cell-wall and membrane components, thus generating other compounds which, in turn, can act as extra- or intracellular messengers (Rao et al., 2000). However, since H2O2 is rapidly removed by antioxidant enzymes and metabolites, it is unlikely that increased production of H2O2 alone is sufficient to orchestrate the complex response to O3 at the whole plant level. Additional signal molecules, such as ethylene, salicylic and jasmonic acid, are believed to be required to act in concert with H2O2 to trigger the plant defence responses (Rao et al., 2000).

On the other hand, although it is scarcely toxic by itself unless at very high concentrations, H2O2 can damage membranes particularly following the reduction to the extremely reactive hydroxyl radicals by transition metals (Fenton reaction). To avoid the oxidative damage, H2O2 levels are kept under control by both enzymatic and non enzymatic scavenging mechanisms present in the apoplast, among which ascorbate peroxidase (APX) is very important (Foyer et al., 1994; Ranieri et al., 1996).

Although extensive studies have been performed aimed at defining the processes involved in O3-induced ROS accumulation (Pellinen et al., 1999; Schraudner et al., 1998; Wohlgemuth et al., 2002), they mainly focused on H2O2 sources and relied on histochemical detection and inhibitor methods. In addition, most of these studies relied on plants showing visible signs of O3-induced injuries. In the present paper, both the generation and degradation of H2O2 in the extracellular matrix of O3-treated sunflower leaves are taken into account by a dual approach which incorporates enzyme activities and subcellular fractionation in addition to staining and inhibitor studies. For this purpose, sunflower (Helianthus annus L., cv. Hor) plants were exposed to a single pulse of 150 ppb of O3 for 4 h. The complex interaction among the different extracellular enzymes involved in H2O2 production and scavenging was studied at different time points during fumigation as well as during the recovery period in non-polluted air. The time-course of ethylene emission is also reported because of the possible interplay between H2O2 and ethylene.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 References
 
Plant material
Sterilized sunflower seeds (Heliantus annuus L., cv. Hor) were germinated, in the dark, in Petri dishes for 3 d and the seedlings were grown in perlite for a week. The seedlings were then transplanted and grown for 4 weeks in a greenhouse at 17/25 °C, night/day, RH 60–80%, a 14 h photoperiod, and a photosynthetic photon flux density of 530 µmol m–2 s–1 (photosynthetic active radiation: 400–700 nm)

Only uniform plants with eight fully-expanded leaves were selected (about 35 d after sowing). All biochemical analyses were carried out on fully-expanded middle-aged leaves from both the control and treated plants.

Fumigation treatment
O3 fumigation was performed in air-conditioned chambers (0.48 m3). The temperature was maintained at 20±1 °C and RH at 85±5%. A photon flux density at plant height of 530 µmol m–2 s–1 was provided by incandescent lamps. O3 was generated by electric disharge passing pure oxygen through a Fisher ozone generator 500 (Fisher Labor und Verfahrenstechnik, Meckenheim, Germany). Ozone concentration in the fumigation chambers was continuously monitored with a Monitor Labs Analyzer mod. 8810 (Monitor Labs, San Diego, CA) operating on the principle of UV absorption and interfaced with a personal computer. Plants were pre-adapted to the chamber conditions for 48 h and then exposed to an acute fumigation with 150 ppb O3 for 4 h. Leaves were collected before (0 h), during (0.5, 1, 2, and 4 h) and after (5, 7, 24, and 48 h) the exposure to the pollutant. Times of measurement refer to hours after the onset of fumigation. Untreated plants were kept in charcoal-filtered air chambers under the same conditions and used as controls.

Ion leakage
Cell damage was assessed by measuring ion leakage from leaf discs. Ten discs of known area (1.13 cm2) were incubated by shaking in 5 ml of distilled water for 3 h at room temperature. The conductivity of the incubation medium was recorded by a JENWAY 4010 conductivity meter (JENWAY, Dunmow, Essex, England). The results are expressed as a percentage of the total conductivity, measured after overnight incubation of the discs frozen in liquid N2.

Lipid peroxidation
The extent of lipid peroxidation was evaluated by the thiobarbituric acid reaction. Leaf tissue was homogenized in 0.1% trichloroacetic acid (1:10, w:v) and centrifuged at 10 000 g for 5 min. 1.0 ml of the supernatant was incubated with 4 ml of 0.5% thiobarbituric acid in 20% trichloroacetic acid at 95 °C for 30 min and then cooled in an ice bath. After centrifugation at 10 000 g for 10 min, the absorbance of the supernatant was read at 532 nm and corrected for the non-specific absorbance recorded at 600 nm. The concentration of thiobarbituric acid reactive substances (TBARS) was calculated as malondialdehyde equivalents using the extinction coefficient of 155 mM–1 cm–1 for malondialdehyde (Ranieri et al., 1996).

Preparation of the apoplastic fluid
Freshly harvested intact leaves (10 g) were rinsed with distilled water and vacuum infiltrated (–65 kPa, three cycles of 30 s each) in 50 ml of 66 mM K-phosphate buffer (pH 7) and 100 mM KCl. Having been wiped, the leaves were centrifuged at 1500 g for 10 min at 4 °C to obtain the intercellular washing fluid (IWF) which, after dialysis against diluted infiltration buffer, was immediately used for biochemical analyses (Ranieri et al., 1996). For APX assay, Na-ascorbate (5 mM) was added to the infiltration and the dialysis buffers. To check the purity of the IWF fraction, the cytoplasmatic and chloroplastic enzyme markers, glucose-6-P dehydrogenase (G6PDH) and glyceraldehyde-3-P dehydrogenase (GAPDH), were measured in both the IWF and the residual cell material (RCM) as previously reported (Ranieri et al., 2000).

Cell-wall fractions
The cell wall ionically- (IB) and covalently-bound (CB) fractions were separated as reported in Ranieri et al. (2001). Freshly-harvested leaves were homogenized at 4 °C with 66 mM Na-phosphate buffer (pH 6.1) and centrifuged at 800 g for 5 min. The pellet was washed twice with phosphate buffer, twice with water and, after 1 h incubation in 2% Triton X-100 at 4 °C with continuous shaking, it was again rinsed 5x with water. The pellet was then incubated with 1 M CaCl2 for 1 h and centrifuged at 800 g for 10 min at 4 °C. The resulting supernatant was the IB fraction. The pellet was washed several times with water and then incubated for 16 h at room temperature with 0.3% cellulase, 0.3% macerase and 0.3% cellulolysin in 50 mM Na-acetate buffer, pH 5.5, to obtain the CB fraction, after centrifugation at 800 g for 10 min. The residual pellet was dried at 80 °C and weighed.

Plasma membrane extraction
Plasma membrane (PM) vescicles were extracted according to Larsson et al. (1987) with some modifications. Freshly-harvested leaves were homogenized at 4 °C with 50 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulphonic acid (HEPES)-KOH (pH 7.5), 330 mM sucrose, 5 mM EDTA, 5 mM dithiothreitol (DTT), 1 mg ml–1 BSA, 2 µg ml–1 PMSF, filtered through four layers of gauze and centrifuged at 13 000 g for 15 min. The recovered supernatant was centrifuged at 80 000 g for a further 30 min and an aliquot of the resulting membrane pellet (microsomal fraction) was resuspended in 5 mM K-phosphate buffer (pH 7.8), 330 mM sucrose, 5 mM KCl, 1 mM DTT, and 0.1 mM EDTA. Plasma membrane vesicles were prepared using a 9 g aqueous two-phase partitioning system. Resuspended microsomal fractions were mixed with 6.5% (w/w) Polyethylene Glycol 3350 (PEG), 6.5% (w/w) Dextran T500, 5 mM K-phosphate buffer (pH 7.8), 330 mM sucrose, 5 mM KCl, 1 mM DTT, and 0.1 mM EDTA. After mixing, the phases were separated by centrifuging at 4000 g for 5 min. The upper phase, enriched in PM vesicles, was repartitioned twice with fresh lower phase without PEG and, after dilution with 50 mM HEPES-KOH (pH 7.5), 330 mM sucrose, the PM vesicles were collected by centrifuging at 40 000 g for 45 min.

The sensitivity of ATPase activity to vanadate (0.1 mM), NaN3 (1 mM) and KNO3 (120 mM) was used as marker of PM, mitochondria and tonoplast activities, respectively (Hodges and Leonard, 1974).

Enzyme activity assay
APX activity was determined following the decrease in absorbance at 290 nm due to the oxidation of ascorbic acid in the first 30 s from the start of the reaction, using the extinction coefficient of 2.8 mM–1 cm–1 for ascorbate. The reaction medium contained 0.5 mM Na-ascorbate, 0.1 mM H2O2, 1 mM EDTA, and 0.1 M HEPES-KOH buffer (pH 7.8) (Ranieri et al., 1996). One enzymatic unit is equivalent to 1 µmol of ascorbic acid oxidized min–1 cm–1. To discriminate between APX and POD activities, 50 mM p-chloromercuribenzoate (pCMB), known to inactivate APX, was added to the enzymatic reaction mixture (Miyake and Asada, 1992).

The activity of POD involved in the lignification process was tested using syringaldazine, a synthetic substrate analogue to the syringilic residue of lignin, as the reducing substrate. The activity of Syr-POD was determined by measuring the increase in absorbance at 530 nm of the reaction mixture containing 100 mM Na-K phosphate buffer pH 6.0, 2.5 mM H2O2, 2 mM syringaldazine, and the protein extract (Pandolfini et al., 1992).

The rate of NAD(P)H oxidation was measured by following the decrease in absorbance at 340 nm of a reaction medium containing 40 mM Na-acetate (pH 5.5), 250 mM sucrose, 1 mM MnCl2, 100 µM salicylhydroxamic acid, 100 µM NADH, and the extract aliquot (Vianello et al., 1997). The activity was calculated using the extinction coefficient of 6.22 mM–1 cm–1 for NADH. To discriminate between the activity of NAD(P)H oxidase complex and NAD(P)H PODs, 15 µM diphenylene iodonium (DPI) or 50 µM KCN, specific inhibitors of NAD(P)H oxidase and POD, respectively, were added to the reaction medium (Bolwell and Wojtaszek, 1997).

The protein content of the extracts was measured spectrophotometrically at 595 nm according to Bradford (1976), using BSA as standard.

Ethylene determination
Fifteen minutes after excision, leaves were incubated within sealed containers at room temperature and, after 1 h, 2 ml samples were withdrawn with a hypodermic syringe. Ethylene evolution was measured by injecting samples into a gas chromatograph equipped with a dual flame ionization detector and a metal column (150x0.4 cm id) packed with alumina (70–230 mesh). The column and detector temperatures were 70 °C and 350 °C, respectively. N2 was used as a carrier at a flow rate of 40 ml min–1 (Mensuali Sodi et al., 1992).

In situ localization of H2O2 accumulation
Hydrogen peroxide production was assessed cytochemically via determination of cerium perhydroxide formation after the reaction of CeCl3 with endogenous H2O2 (Bestwick et al., 1997). Freshly harvested leaves were cut into slices (1–2 mm2) which were incubated for 1 h in 5 mM CeCl3 in 50 mM 3-(N-morpholino)propanesulphonic acid (MOPS) pH 7.2, fixed in 1.25% glutaraldehyde, 1.25% paraformaldehyde in 50 mM Na-cacodylate buffer (CAB) pH 7.2 for 1 h, and washed twice in CAB buffer for 10 min (Bestwick et al., 1997). After post-fixation for 2 h in 1% osmium tetroxide in 50 mM Na-cacodylate buffer, pH 7.2, samples were washed twice in the same buffer, dehydrated in a graded ethanol series (25, 50, 75, 90, and 100%), transferred into propylene oxide, and gradually embedded in Epon-Araldite. Thin sections of embedded tissues were obtained on a Reichert-Ultracut microtome, mounted on uncoated copper grids and observed using a transmission electron microscope (Hitachi 300, Tokyo, Japan) at 75kV.

For the treatment with inhibitors, the leaf slices were pre-incubated for 30 min in 50 mM MOPS pH 7.2 containing either 3 mM KCN (to inhibit POD), 8 µM DPI (to inhibit NAD(P)H oxidase complex) or 25 µg ml–1 bovine liver catalase (to decompose H2O2) (Bestwick et al., 1997). The leaf slices were then incubated for 1 h in CeCl3 solutions supplemented with inhibitors at the concentrations reported above and treated as described above.

Statistical analysis
A minimum of 12 plants per treatment were used in all the experiments. Values shown in the figures are the means of six determinations ±standard error, except for ethylene (n=3). Comparison between means was evaluated by t-test and the P=0.05 level of error.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 References
 
With the aim of studying the mechanisms involved in the O3-induced oxidative burst, sunflower plants were exposed to a single pulse of 150 ppb of O3 for 4 h and left to recover in pollutant-free air up to 48 h. Measurements were performed before (0 h), during (0.5, 1, 2, and 4 h) and after (5, 7, 24, and 48 h) the exposure to the pollutant. Times of measurement refer to hours after the beginning of the O3 exposure.

At the end of the O3 fumigation period, sunflower plants did not exhibit any visible sign of injury on the leaf surface. O3-induced lesions were not evident even 48 h after the onset of the exposure. The integrity of cell membranes, assessed by ion leakage from leaf discs, was not affected by O3 treatment at any time of exposure, as shown in Fig. 1A. The degree of lipid peroxidation, evaluated by measuring the content of TBA-reactive substances, was also found to be unaffected by O3 during the exposure to the pollutant as well as during the recovery period in filtered air (Fig. 1B).



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  		Fig. 1. Evaluation of the integrity of cell plasma membranes of sunflower plants exposed to filtered air or to 150 ppb of O3. Black bars indicate the duration of exposure. (A) Ion leakage (±SE) from leaf discs expressed as a percentage of total leakage from discs frozen in liquid N2. (B) Thiobarbituric acid reactive substances content (±SE) expressed as nmol malondialdehyde equivalents g–1 fresh weight. Statistically significant differences between O3-treated samples (filled circles) and the respective controls (open circles) at each time point are indicated with an asterisk; P=0.05, n=6. The boxes show ion leakage and TBARS content, respectively, measured during the 4 h exposure to O3.

 
In situ localization of H2O2 accumulation
The histochemical assay based on the reaction of H2O2 with CeCl3 produced electron-dense insoluble precipitates of cerium perhydroxides at sites where H2O2 was accumulated. In the control sunflower leaves (Fig. 2A), electron dense precipitates of cerium perhydroxides were not detectable in the cell wall of palisade parenchyma or spongy mesophyll cells. Starting from 30 min of O3 exposure, a progressive increase in the number and size of electron-dense precipitates was clearly visible in the cell walls of treated leaves. After 2 h, most cell walls presented a faint or medium intensity of CeCl3 staining (Fig. 2B, C). Four hours after the beginning of the O3 exposure, H2O2 accumulation reached its maximum level, with a particularly strong staining at the sites of connection between adjacent cell walls, close to the intercellular spaces (Fig. 2D). At any time considered, H2O2 accumulation was most evident in the cell walls of spongy mesophyll cells while the CeCl3 precipitates in palisade parenchima cells were faint and dispersed. Hydrogen peroxide never accumulated in the cells; no precipitates were, in fact, detectable in the mithocondria, chloroplasts or in the cytosol (Fig. 2B, C). Hydrogen peroxide accumulation was detected neither in the cell walls nor inside the cells at any time during the post-fumigation period (data not shown).



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  		Fig. 2. Cytochemical localization of O3-induced H2O2 accumulation in spongy mesophyll cells. (A) Cells exposed to filtered air show no CeCl3 staining. (B, C) Cell walls accumulate H2O2 2 h after the beginning of the O3 exposure: the intensity of CeCl3 precipitation appears faint (B) or medium (C). (D) Strong H2O2 accumulation on cell walls between adjacent cells close to the intercellular spaces after 4 h of O3 exposure. Abbreviations: cw, cell wall; c, chloroplast; m, mitochondrion; s, starch grain. Bar, 1 µm.

 
In attempts to determine the possible sources of H2O2 accumulation, leaf slices were pre-incubated with inhibitors of H2O2-producing enzymes before CeCl3 treatment, according to Bestwick et al. (1997). The effects from the spongy mesophyll cells were analysed. Table 1 presents data from the 4 h O3-treated sunflower sample. The specificity of CeCl3 staining for H2O2 was demonstrated by the almost complete removal of electron dense precipitates in the catalase-treated sample. DPI, an inhibitor of the PM NADPH oxidase complex, and KCN, which inhibits PODs, both reduced the number of H2O2 producing cells; moreover, in these treated cells, only faint or medium staining was observed, suggesting that both NADPH oxidase complex and NAD(P)H PODs are involved in the apoplast H2O2 accumulation.


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  		Table 1. Effects of treatment with inhibitors on the cell wall H2O2 accumulation in 4 h O3 exposed sunflower leaves Leaves were incubated with 25 µg ml–1 catalase, 8 µM DPI, and 3 mM KCN. The intensity of CeCl3 staining (% of scored cells), evaluated on 50 spongy mesophyll cells, is classified as faint (as in Fig. 2B), medium (as in Fig. 2C) and strong (as in Fig. 2D).
 
Enzyme activity
The contamination of IWF by intracellular molecules was always lower than 0.1%, as indicated by the activity of the marker enzymes, glucose-6-P dehydrogenase, and glyceraldehyde-3-P dehydrogenase (data not shown).

Syringaldazine is commonly used as a specific substrate to measure POD activities involved in polymerization of lignin precursors. Thirty minutes after the onset of the fumigation treatment, Syr-POD activity of IWF was found to be significantly decreased by O3 exposure (–54%, Fig. 3A). This situation was quickly recovered after 1 h, when no difference between the treated and control samples was detected. Meanwhile, starting from 2 h of exposure, Syr-POD activity increased significantly above that of the controls (+59% and +38% after 2 h and 4 h, compared with the respective controls, Fig. 3A). After 1 h of recovery in filtered air, Syr-POD activity of O3-treated plants remained significantly higher than in the controls (+25%), but it declined against the control values at later time points (Fig. 3A).



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  		Fig. 3. Syringaldazine peroxidase activity (±SE), measured in the intercellular washing fluid (A), in the cell wall ionically (B) and covalently-bound (C) fractions of sunflower plants exposed to filtered air or to 150 ppb of O3. Black bars indicate the duration of exposure. The activity is expressed as {Delta}A530 min–1 mg–1 proteins, except for the covalently-bound fraction, whose activity is reported as {Delta}Abs530 min–1 g–1 dry weight of residual cell wall material. Statistically significant differences between O3-treated samples (filled circles) and the respective controls (open circles) at each time point are indicated with an asterisk; P=0.05, n=6. The boxes show syringaldazine POD activity measured in the different extracellular fractions during the 4 h exposure to O3.

 
A different behaviour was shown by the cell-wall bound Syr-POD. In fact, while IB activity was not affected by O3 fumigation at any time of exposure nor during the post-fumigation period (Fig. 3B), a significant increase in Syr-POD activity was detected in the CB fraction starting from 2 h of treatment (+22% and +65% after 2 h and 4 h, compared with the respective controls, Fig. 3C). The Syr-POD activity of the CB fraction also remained significantly higher than in the controls during the whole recovery period in filtered air except at 48 h (+32%, +13%, 14%, after 5, 7, and 24 h, respectively, Fig. 3C). Similar results were also obtained when POD activity was tested using the natural lignin monomer, ferulic acid, as the reducing substrate (data not shown).

Unlike lignifying POD activities, IWF APX activity was significantly enhanced by O3 as early as 30 min after the onset of exposure (+27%, Fig. 4). The extent of the increase remained almost constant throughout the whole fumigation period (+24%, +28%, +21%, respectively, after 1, 2, and 4 h, Fig. 4) and, with the exception of a more pronounced increase recorded at 5h (+52%), even during the recovery in pollutant-free air (+28%, +20%, and +19%, at 7, 24, and 48 h, respectively, Fig. 4). The addition of p-CMB to the assay mixture, as well as the removal of ascorbic acid from the infiltration washing solution, resulted in the loss of enzymatic activity, confirming the presence of APX in the apoplast (data not shown).



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  		Fig. 4. Ascorbate peroxidase activity (±SE), reported as µmol ascorbic acid oxidized min–1 mg–1 proteins, in the intercellular washing fluid of sunflower plants exposed to filtered air or to 150 ppb of O3. Black bars indicate the duration of exposure. Statistically significant differences between O3-treated samples (filled circles) and the respective controls (open circles) at each time point are indicated with an asterisk; P=0.05, n=6. The boxes show APX activity measured during the 4 h exposure to O3.

 
To evaluate whether an endogenous H2O2 generation occurred following O3 exposure, NAD(P)H oxidation was measured in the free apoplastic and cell-wall fractions as well as in the PM one. Based on marker enzyme activities, vesicle preparations were highly enriched in the PM (data not shown).

DPI and KCN, known to inhibit the NAD(P)H oxidase complex and POD activity, respectively, were added to the reaction medium to discriminate between H2O2 production by NAD(P)H oxidase complex and by NAD(P)H PODs. The use of these two inhibitors revealed that H2O2 generation by PM fraction was sustained by the NAD(P)H oxidase complex, because no NAD(P)H oxidation was recorded when DPI was added to the reaction mixture (data not shown). Conversely, in both the IWF and cell-wall fractions, the production of H2O2 at the expense of NAD(P)H oxidation was catalysed by PODs. In fact, while an almost complete inhibition of enzyme activity was achieved by adding KCN to the reaction medium, the addition of DPI resulted in being ineffective (data not shown).

The activity of the PM NAD(P)H oxidase complex was unaffected by O3 exposure up to 2 h and 4 h of exposure, when a significant increase was detected (+31% and +80%, compared with the respective controls, Fig. 5A). NADP(H) oxidase activity in PM vesicles from O3-treated plants was significantly higher than the control one also at 5 h (+29%). However, 7 h after the onset of the fumigation, the activity    returned to almost the same levels as at the beginning of the exposure (Fig. 5A).



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  		Fig. 5. NAD(P)H oxidation activity (±SE) measured in the plasma membrane fraction (A), in the intercellular washing fluid (B), in the cell wall ionically-bound (C) and covalently-bound (D) fractions of sunflower plants exposed to filtered air or to 150 ppb of O3. Black bars indicate the duration of exposure. The activity is expressed as µmol NAD(P)H oxidized min–1 mg–1 proteins, except for the covalently-bound fraction, whose activity is reported as {Delta}Abs530 min–1 g–1 dry weight of residual cell wall material. Statistically significant differences between O3-treated samples (filled circles) and the respective controls (open circles) at each time point are indicated with an asterisk; P=0.05, n=6. The boxes show NAD(P)H oxidation activity measured in the different extracellular fractions during the 4 h exposure to O3.

 
A similar trend was also observed with regards to the activity measured in the IWF fraction, where no differences were detected between NAD(P)H oxidation rate of treated and control samples, both 30 min and 1 h after the beginning of exposure, whilst a significant stimulation was found after 2, 4, and 5 h (+65%, +90%, and +22%, respectively, Fig. 5B). Again, at later time points of the post-fumigation period, NAD(P)H oxidation activity was found to decline back to the control values (Fig. 5B).

NAD(P)H oxidation by IB fraction was quickly stimulated by O3 treatment (+69%, after 30 min of fumigation) and remained significantly higher than the control activity up to 2 h of exposure (+34% and +46% after 1 h and 2 h, compared with the respective controls), while it returned to the control values 4 h after the onset of the fumigation (Fig. 5C). No increase in the enzyme activity was detected during the recovery period in pollutant-free air (Fig. 5C).

No significant effect of O3 on the rate of NAD(P)H oxidation by CB fraction was observed at any time point during O3 exposure (Fig. 5D). Conversely, a significant stimulation of NAD(P)H oxidation activity was detected starting from the first hour of the post-fumigation period (+36%, +82%, and +30% after 5, 7, and 24 h, respectively, Fig. 5D). However, 48 h after the onset of the O3 exposure, similar activities were measured in the control and O3-treated plants concerned (Fig. 5D).

Ethylene evolution
The time-course of ethylene evolution showed that its emission was a rapid reaction to O3 exposure, being stimulated as early as after 30 min of exposure (+28%). The ethylene evolution reached the maximum value after 2 h of fumigation (+80%) and, at the end of the experiment (4 h), it declined to the control levels (Fig. 6). No further increase in ethylene emission was observed during the post-fumigation period (Fig. 6).



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  		Fig. 6. Leaf ethylene emission (±SE) expressed as nl ethylene h–1 g–1 fresh weight of sunflower plants exposed to filtered air or to 150 ppb of O3. Black bars indicate the duration of exposure. Statistically significant differences between O3-treated samples (filled circles) and the respective controls (open circles) at each time point are indicated with an asterisk; P=0.05, n=3. The boxes show ethylene emission measured during the 4 h exposure to O3.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
References
 
During the last decade, increasing attention has been paid to the dual role of H2O2 as a toxic molecule and cellular messenger triggering the elicitation of the diversified effects which constitute the plant response to O3 (Kangasjärvi et al., 1994; Rao et al., 2000; Schraudner et al., 1998). In sunflower leaves, H2O2 accumulation, observed after only 30 min of exposure to 150 ppb of O3, was one of the earliest detectable metabolic responses to the imposed stress. In accordance with these results, O3 has recently been reported to activate an oxidative burst, also resulting in H2O2 production in both sensitive and resistant tobacco (Nicotiana tabacum L.) cultivars Bel W3 and Bel B during a 5 h exposure period (Schraudner et al., 1998), as well as in some arabidopsis (Arabidopsis thaliana) genotypes (Rao and Davis, 1999), in birch (Betula pendula) leaves (Pellinen et al., 1999), and in tomato (Lycopersicon esculentum) plants (Wohlgemuth et al., 2002). The cytochemical localization of cerium perhydroxide precipitates of O3-treated sunflower leaves demonstrated that H2O2 accumulation only occurred in the apoplast and it coincided temporally with the fumigation period. The sites of H2O2 accumulation have been correlated with the formation of HR-type lesions observed in plants exposed to O3 (Pellinen et al., 1999; Wohlgemuth et al., 2002). This was not so in the case of the sunflower, where, in spite of H2O2 accumulation in O3-treated samples, no visible sign of injury was evident on the leaf surfaces, a result confirmed by the absence of alteration at plasma membrane level, as indicated by both unchanged ion leakage and TBARS content. Such a result is probably to be attributed to the absence of a second oxidative burst and intracellular H2O2 production during the post-fumigation period, as opposed to reports on plants developing leaf injuries (Pellinen et al., 1999; Schraudner et al., 1998).

Hydrogen peroxide production following biotic and abiotic stresses has been mainly ascribed to a plasma membrane NAD(P)H oxidase activity (Amicucci et al., 1999; Keller et al., 1998; Rao and Davis, 1999; Torres et al., 2002) and/or to pH-dependent cell-wall POD activity (Bolwell, 1999; Bolwell and Wojtaszek, 1997), even if the involvement of other H2O2-producing enzymes, such as oxalate oxidase and amine oxidase (Bolwell and Wojtaszek, 1997), which have not been tested in the present experiment, cannot be excluded as well. The inhibitor experiments suggested that in sunflower leaves both PM NAD(P)H oxidase complex and cell-wall NAD(P)H PODs could be the primary site of H2O2 generation. However, attention should be paid to the use of inhibitors, since they are not fully specific. In fact, even if at higher concentrations than those used in the present experiment, DPI shows a peroxidase-inhibitory activity (Barceló, 1998), and both DPI and KCN, in addition to their inhibitory activity, have the ability to scavenge H2O2 (Baker et al., 1998). The results from cytochemical H2O2 location and inhibitor use were thus integrated with measurements of enzyme activities involved in H2O2 turnover at subcellular level.

The origin of H2O2 during and after O3 exposure in sunflower leaves is intriguing and seems to be a complex and highly regulated event, requiring the time-dependent co-ordinated stimulation and down-regulation of different sets of differently located enzymes. In fact, at the beginning of the O3 exposure, besides the possible H2O2 generation by direct O3 degradation in the apoplast or by reaction of O3 with cell wall and PM compounds (Rao et al., 2000), H2O2 production was sustained by NAD(P)H POD activity of the IB fraction, while no contribution by both IWF and CB NAD(P)H PODs or PM NAD(P)H oxidase complex was made. A decreased H2O2 utilization, as a consequence of the reduced activity of IWF Syr-POD recorded 30 min after the onset of fumigation, could also be involved in such an early H2O2 accumulation. Since extracellular PODs, which catalyse the lignification process, are known to require an acidic environment, with a pH optimum ranging from 4.5 to 6.0 (Otter and Polle, 1997), this early and transient decrease in Syr-POD activity could be due to the O3-induced apoplast alkalization. At the same time, such a pH shift towards basic values could be of major importance in triggering H2O2 production by cell-wall NAD(P)H PODs. In fact, as reported by Bolwell and Wojtaszek (1997), POD ability to generate H2O2 is dependent upon extracellular alkalization, the maximum production occurring at neutral to basic pH, depending upon the isoform studied.

Starting from 2 h of exposure to the pollutant, the mechanisms responsible for H2O2 production differed from those observed at the beginning of fumigation. In fact, IWF NAD(P) POD and by PM NAD(P)H oxidase complex seemed to be the primary sites of H2O2 generation during the last time points of O3 treatment (2 h and 4 h) as well as after 1 h of recovery in non-polluted air. Finally, another different scenario was evident during the post-fumigation period (5, 7 and 24 h), when H2O2 production was sustained by the later activation of NAD(P)H PODs present in the CB fraction.

As far as the H2O2-consuming Syr-PODs are concerned, and contrary to what was observed at the beginning of the fumigation, their activity was significantly enhanced by O3 starting from 2 h of O3 exposure in IWF and CB fractions. This finding is in accordance with previously reported results on stimulation of POD activity following O3 stress (Castillo and Greppin, 1986; Peters et al., 1988; Ranieri et al., 1996, 1998). As a consequence, the peroxidative cross linking of lignin precursors, cell wall proteins and polysaccharides led to cell-wall stiffening and, by increasing the tortuosity of the diffusion pathway of O3, contributed to slowing down O3 penetration. In the CB fraction, Syr-POD remained significantly higher than the control up to the end of the recovery period in filtered air, thus contributing to scavenge H2O2. During the post-fumigation period no CeCl3 staining was, in fact, observed despite an active H2O2 production by CB NAD(P)H PODs.

A different behaviour was shown by IB POD, with the results unaffected by O3 exposure at any time point. It thus appears that the reaction of the different POD isoforms to O3 stress was dependent on their subcellular location. A similar different response by extracellular sunflower PODs was previously observed following iron-deficiency, which was found to affect Syr- POD activity in the CB fraction, but not in the IB one (Ranieri et al., 2001). However, until now, in spite of intense investigation, no clear correlation has been established between the role of different POD isozymes and their subcellular location in the cell wall (Penel et al., 2000), although many different reactions catalysed by PODs in the cell wall are supposed to require a fine control of microlocalization.

Although H2O2 is acknowledged to be an important signal molecule, it should be remembered that its levels need to be carefully tuned to avoid uncontrolled oxidative damage. Since lower levels of H2O2 seem to be sufficient to activate defensive genes rather than those required to trigger cell death (Levine et al., 1994), the capability of achieving optimal H2O2 levels sufficiently high to ensure the signal transduction, but not high enough to induce oxidative damage, is of major importance in determining the fate of the plant cell. In this context the antioxidant enzymes and metabolites play a key role. APX is recognized as one of the most efficient ROS scavenging system (Foyer et al., 1994) because of its high affinity for H2O2 and its presence in different subcellular compartments. The stimulation of APX by O3 exposure, frequently observed in many plant species (Castillo and Greppin, 1986; Peters et al., 1988; Ranieri et al., 1996, 1998, 2000), underlines the key role of this enzyme in H2O2 detoxification. In sunflower plants, the prompt stimulation of extracellular APX activity, which was detected as early as 30 min after the beginning of O3 exposure and remained significantly higher than the control even in the post-fumigation period, may be important in avoiding the build-up of toxic H2O2 concentrations. To confirm the importance of the extracellular matrix as a first line of defence against O3-induced stress, no change in intracellular APX activity was observed under the present fumigation conditions (data not shown). The rapid enhancement of APX activity was probably due to an increased rate of ascorbate peroxidation by pre-existing enzymes, although de novo enzyme synthesis could not be excluded either. In fact, an increase in both intra- and extracellular APX protein content was previously detected in sunflower plants after 4 d (4 h d–1) of exposure to 150 ppb of O3 (Ranieri et al., 2000). Similarly, analysis of APX mRNA levels in arabidopsis treated with 100–150 ppb of O3 revealed enhanced cytosolic mRNA levels within 3–4 h (Kubo et al., 1995), while exposure to 300 ppb of O3 lead to increased transcript levels of a cytosolic APX starting from 30 min of treatment (Rao and Davis, 1999).

Increased ethylene production was one of the earliest events observed in both herbaceous and tree species in response to O3 treatment (Langebartels et al., 1991; Overmeyer et al., 2000; Sandermann, 1996; Wellburn and Wellburn, 1996). An interplay between ethylene and ROS was recently suggested by Moeder and co-workers (2002), who proposed that ethylene synthesis and perception were required for active H2O2 production in O3-exposed tomato. On the other hand, the finding that transgenic tobacco lines retaining a very low catalase activity (CAT1AS) showed a dramatic, transient increase in ethylene production 2–3 h after exposure to high light, which followed H2O2 accumulation, suggesting that H2O2 could act as a signal upstream of ethylene (Chamnongpol et al., 1998). Based on the result of the present experiment, a clear temporal correlation could not be established between these two signalling molecules. In fact, in sunflower leaves, both H2O2 and ethylene levels increased at a very early stage of the treatment, i.e. 30 min after the onset of the fumigation, although, in accordance with the finding of Moeder et al. (2002), the peak of ethylene emission, detected after 2 h of O3-exposure, preceded the maximum H2O2 accumulation, observed at the end of the fumigation. Treatments with inhibitors of ethylene biosynthesis or perception are in progress to clarify the possible role of ethylene in inducing H2O2-mediated oxidative burst.

Ethylene emission by O3-exposed plants was correlated to O3 sensitivity and the appearance of leaf injuries (Kangasjärvi et al., 1994; Sandermann et al., 1998). Cell damage has been initially ascribed to a chemical reaction between ethylene and O3, yielding a radical generation that, in turn, would lead to lipid peroxidation and tissue injury (Elstner et al., 1985; Mehlhorn and Wellburn, 1987). More recently, however, some authors (Moeder et al., 2002; Overmeyer et al., 2000; Sandermann et al., 1998; Tuomainen et al., 1997) have reported evidence of ethylene playing an active role in lesion development as a component of the signal transduction pathway leading to programmed cell death, so that both a highly regulated increase in ethylene emission and functional ethylene perception and signalling are required. However, by con trast with the commonly reported evidence, ethylene evolution by O3-treated sunflower leaves was accompanied by neither increased lipid peroxidation nor leaf damage, suggesting the existence of threshold levels below which ethylene is ineffective and/or the involvement of factors other than ethylene in inducing lesion formation. It should be remembered that the final response to O3 depends on the cross-talking between the different signalling routes, involving not only ROS and ethylene, but also jasmonic (JA) and salicylic acid (SA), which interact with each other in a synergetic or antagonistic manner (Overmeyer et al., 2000; Rao et al., 2000). In this context SA is known to induce H2O2 accumulation by inhibiting catalase activity through specific binding to the enzyme (Chen et al., 1993) or by inducing H2O2 formation by peroxidases (Kawano and Muto, 2000), while JA is believed to desensitize the O3-induced oxidative burst and the SA-mediated amplification loop which results in the production of excess ROS (Overmeyer et al., 2000; Rao et al., 2000).

In conclusion, the early H2O2 accumulation in sunflower plants seems to be the result of a highly regulated time-dependent stimulation and down-regulation of differently located enzymes which produce or scavenge H2O2. A summary of the mechanisms involved is illustrated in Table 2. Further experiments aimed at investigating the behaviour of other extracellular enzymes and metabolites involved in H2O2 turn over, as well as studies directed to unravel the cross-talking among H2O2, ethylene and other signalling molecules, are required.


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  		Table 2. Schematic representation of the interplay between H2O2 producing and scavenging mechanisms in sunflower plants exposed to 150 ppb of O3 for 4 h Times of measurement refer to h after the onset of fumigation during both the exposure to the pollutant (0.5, 1, 2, and 4 h) and the recovery in non-polluted air (5, 7, 24, and 48 h). The symbols {uparrow} and {downarrow} preceding the name of enzymes refer to the detected enhanced or diminished activities, respectively.
 

   Acknowledgements
 
This research was supported by a grant from MURST (National Project) Rome, Italy and by funds from the University of Pisa.


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