RESEARCH PAPER |
Roles for redox regulation in leaf senescence of pea plants grown on different sources of nitrogen nutrition
1Crop Performance and Improvement Division, Rothamstead Research, Harpenden, Hertfordshire AL5 2JQ, UK
2Departamento de Bioquímica, Biología Celular y Molecular de Plantas, Estación Experimental del Zaidín, CSIC, Granada, Spain
3Departamento de Agroecología y Protección Vegetal, Estación Experimental del Zaidín, CSIC, Granada, Spain
4Departamento de Biología del Estrés y Patología Vegetal, Centro de Edafología y Biología Aplicada del Segura, CSIC, Murcia, Spain
To whom correspondence should be addressed. E-mail: luisalfonso.delrio{at}eez.csic.es
Received 31 March 2006; Accepted 6 April 2006
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Leaf senescence and associated changes in redox components were monitored in commercial pea (Pisum sativum L. cv. Phoenix) plants grown under different nitrogen regimes for 12 weeks until both nodules and leaves had fully senesced. One group of plants was inoculated with Rhizobium leguminosarum and grown with nutrient solution without nitrogen. A second group was not inoculated and these were grown on complete nutrient solution containing nitrogen. Leaf senescence was evident at 11 weeks in both sets of plants as determined by decreases in leaf chlorophyll and protein. However, a marked decrease in photosynthesis was observed in nodulated plants at 9 weeks. Losses in the leaf ascorbate pool preceded leaf senescence, but leaf glutathione decreased only during the senescence phase. Large decreases in dehydroascorbate reductase and catalase activities were observed after 9 weeks, but the activities of other antioxidant enzymes remained high even at 11 weeks. The extent of lipid peroxidation, the number of protein carbonyl groups and the level of H2O2 in the leaves of both nitrate-fed and nodulated plants were highest at the later stages of senescence. At 12 weeks, the leaves of nodulated plants had more protein carbonyl groups and greater lipid peroxidation than the nitrate-fed controls. These results demonstrate that the leaves of nodulated plants undergo an earlier inhibition of photosynthesis and suffer enhanced oxidation during the senescence phase than those from nitrate-fed plants.
Key words: Redox regulation, redox signalling, root nodules, senescence, symbiotic nitrogen fixation
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Leaf senescence is essentially a reversible process until the very last stages which culminate in programmed cell death (PCD; Thomas et al., 2003). It is characterized by a decline in photosynthesis accompanied by the loss of ribulose-1,5-bisphosphate carboxylase/oxygenase and chlorophyll/protein complexes, together with increases in proteinase activities and in lipid peroxidation and membrane leakiness (Buchanan-Wollaston, 1997; del Río et al., 1998; Thompson et al., 1999; Palma et al., 2002; Navabpour et al., 2003). In some species such as Arabidopsis, leaf age can be used as a predictor of the timing of senescence; the sequential senescence of rosette leaves coinciding with maximum inflorescence development (Hensel et al., 1993). Leaf chlorophyll and protein contents are often used as indicators of leaf senescence. Transcripts encoding genes related to photosynthesis and protein synthesis are decreased during senescence and there is increased expression of senescence-associated genes (SAGs). These are useful molecular markers of the developmental changes in gene expression that underpin the senescence process (Nam, 1997). However, SAGs are also expressed in response to increases in the tissue contents of reactive oxygen species (ROS; Navabpour et al., 2003). Increased cellular oxidation is a key feature of leaf senescence (del Río et al., 1998; Navabpour et al., 2003).
PCD may be triggered by enhanced ROS levels and cellular oxidation (Chen and Dickman, 2004; Laloi et al., 2004; Wagner et al., 2004). Recent genetic evidence suggests that ROS do not trigger PCD or senescence by causing physicochemical damage to the cell but rather these metabolites act as signals that activate genetically programmed pathways of gene expression that lead to regulated cell suicide events (Foyer and Noctor, 2005a, b). Plants use ROS as second messengers in many signal transduction cascades from mitosis to PCD, and thus ROS accumulation is crucial to plant development as well as defence. Similarly, the plant antioxidant defence network that is comprised of low molecular weight antioxidants and antioxidant enzymes, is important in controlling the life-time of the ROS signals as well as in preventing uncontrolled oxidation.
Leguminous plants such as soybean, pea, cowpea, and peanut, are central to environment-friendly agriculture. They are extremely valuable in all sustainable agricultural systems because they provide a biological alternative to the application of chemical nitrogen (N) fertilizers. Legumes are widely used as a natural source of N for crop rotation and intercropping techniques, particularly in developing countries as they add N to the soil. This valuable attribute is due to the symbiotic association between leguminous plants and the N-fixing rhizobia that are housed in specialized organs called nodules (Schultze and Kondorosi, 1998; Gage and Margolin, 2000). The present study was undertaken to explore the influence of nodulation on the oxidative events associated with leaf senescence compared with those occurring in plants supplied with an optimal level of soil nitrate and ammonium. It is shown that the leaves of nodulated plants exhibit symptoms of enhanced oxidation during the senescence process.
|
Materials and methods |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Plant material and growth conditions
Pea seeds (Pisum sativum L. cv. Phoenix) were supplied by Dr Peter Römer from Südwestdeutsche Saatzucht, Rastatt, Germany. Seeds were surface-sterilized and then germinated on vermiculite for 14 d in controlled-environment chambers. One group of plants was inoculated with Rhizobium leguminosarum (Microbio Rhizogen Corporation, UK) and grown with nitrogen-free Hoagland's nutrient solution (Olivares et al., 1980). A second group was not inoculated and was grown on a similar solution containing 5 mM KNO3 (controls). Healthy and vigorous seedlings of the two plant groups were selected and grown in aerated nutrient solutions under the two nitrogen regimes. Plants were grown in growth chambers with a 25/19 °C day/night and with 70–85% humidity. They were grown with a 14 h photoperiod at a photosynthetic photon flux density of 600 µmol m–2 s–1. Flower initiation occurred at 9 weeks, a stage when the plants were designated as mature. Plants were grown up to 12 weeks, a point where both nodules and leaves had fully senesced.
Photosynthesis measurements
Steady-state photosynthetic gas exchange parameters were measured 3 h into the photoperiod on the fourth leaf from each plant using a model LCA-3 portable, integrated infrared CO2 analyser (open system; Analytical Development Co., Hoddesdon, UK). Light (1180 µmol m–2 s–1) was provided by a halogen lamp (General Electric 300 PAR 56/WFL). Irradiance was sufficient to ensure that maximal rates of CO2 assimilation (Amax) were attained at each stage of development (Long and Hällgren, 1985). Water use efficiency was measured as described by Ludlow and Muchow (1990).
Preparation of soluble protein and metabolite extracts
Total soluble protein extracts:
Leaves were washed and homogenized in ice-cold 100 mM HEPES buffer, pH 6.5, containing 10 mM MgCl2, 5 mM EDTA, 5 mM ascorbate, 1 mM PMSF, and 20% (w/v) PVP (1/4; w/v). Homogenates were filtered through two layers of Miracloth and centrifuged at 27 000 g for 20 min and the supernatants were used for the enzymatic determinations.
Metabolite extracts:
Leaf tissue (50–150 mg) was ground in liquid N2 and 1 ml of ice-cold 1 M HClO4 was added and ground into the frozen powder. On thawing, the homogenate was ground again. Aliquots (0.05 ml) were taken for the assay of chlorophyll as pheophytin, and the remainder was centrifuged for 15 min at maximum speed on a desktop centrifuge at 4 °C. An aliquot (0.5 ml) of the supernatant was decanted into to a fresh tube and 0.1 ml 0.12 M NaH2PO4 (pH 5.6) added. Then sufficient K2CO3 was added drop-wise to bring the pH to 5–6. The insoluble KClO4 thus produced was removed by centrifugation and the supernatant was decanted into to a fresh tube and used for metabolite (ascorbate and glutathione) analysis.
Determination of ascorbate and glutathione
Reduced ascorbate was measured via the ascorbate oxidase-dependent decrease in absorbance at 265 nm as described by Foyer et al. (1983). Total ascorbate was measured in the same way but following incubation of the sample with dithiothreitol. Dehydroascorbate was determined as the difference between total and reduced ascorbate. Total and oxidized glutathione were measured as the NADPH-driven glutathione-dependent reduction of DTNB (Griffith, 1980), in the presence of glutathione reductase. Inclusion of glutathione reductase in the assay medium facilitates glutathione cycling, the assay thus measuring both reduced glutathione (GSH) and glutathione disulphide (GSSG) as total glutathione. To distinguish between GSH and GSSG, extracts were treated with 2-vinylpyridine, a reagent that complexes GSH (but not GSSG). Thus, only GSSG is measured in vinylpyridine-treated samples. GSH was determined as the difference between total glutathione and GSSG.
Determination of antioxidative enzyme activities
All measurements were made at 25 °C, except for catalase (20 °C), and were performed four times for each sample. The APX, MDHAR, DHAR, and GR activities were assayed spectrophotometrically, as described by Vanacker et al. (1998). SOD activity was determined by the ferricytochrome c method using xanthine/xanthine oxidase as the source of superoxide radicals (McCord and Fridovich, 1969). Catalase activity was measured at 20 °C following the O2 evolution produced by the enzyme action on H2O2 with a Clark oxygen electrode (del Río et al., 1977).
Semi-quantitative RT-PCR
For the isolation of total RNA, pea leaves were frozen in liquid nitrogen and stored at –80 °C. Total RNA was extracted using the Trizol method (GibcoBRL, Life Technologies, Carlsbad, CA, USA) according to the manufacturer's instructions and was stored at –80 °C. Two micrograms of total RNA from senescent leaves of both nodulated and nitrate-fed plants was used as a template for the reverse transcriptase (RT) reaction and was added to a mixture containing 5 mM MgCl2, 1 mM dNTPs, 0.5 micrograms oligo (dT) primers, 1x RT-buffer, 20 U RNasin ribonuclease inhibitor, 15 U AMV reverse transcriptase (Promega, Madison, WI, USA). The reaction was carried out at 42 °C for 40 min, followed by a 5 min step at 98 °C, and then by cooling at 4 °C. Amplification of actin II cDNA from pea (X68649
[GenBank]
) was chosen as a control. Each specific cDNA and actin II cDNAs were amplified by polymerase chain reaction (PCR) as follows: 1 µl of the cDNA produced (diluted 1/20) was added to 250 µM dNTPs, 1.5 mM MgCl2, 1x PCR buffer, 1 U of Ampli Taq Gold (PE Applied Biosystems), and 0.5 µM of each specific nucleotides (Table 1) in a final volume of 20 µl. Reactions were carried out in a Hybaid thermo-cycler. A first step of 10 min at 94 °C was followed by 28–33 cycles (depending on the target gene) of 30 s at 94 °C, 30 s at 60 °C, and 45 s at 72 °C. Amplified PCR products were detected after electrophoresis in 1% agarose gels stained with ethidium bromide.
|
Quantification of the bands was performed using a Gel Doc system (Bio-Rad Laboratories, Hercules, CA, USA) coupled to a highly sensitive CCD camera. Band intensity was expressed as relative absorbance units. To normalize for initial variations in sample concentration, the ratio between the specific cDNA and actin II amplification was calculated. Mean and standard deviations were calculated after normalization to actin II.
Determination of H2O2
Assay in leaf extracts:
The H2O2 concentration of crude extracts from pea leaves was determined by spectrofluorometry according to the method of Creissen et al. (1999), as modified by Romero-Puertas et al. (2004). All operations were performed at 0–4 °C. Leaves (0.4 g) were homogenized in 1.2 ml of 25 mM HCl, and the crude extracts were filtered through two nylon layers, and the pigments were removed by mixing with 15 mg of charcoal (Sigma). The pigment-containing charcoal was separated by centrifugation at 5000 g for 5 min, and the supernatants were clarified by filtration through a 0.22 µm filter unit. The pH of leaf extracts was adjusted to 7.0 with NaOH and these extracts were used to measure the H2O2 concentration. The reaction mixtures (3 ml) contained 50 mM HEPES buffer, pH 7.6, 5 mM homovanillic acid, and 10–100 µl of sample. The reaction was started by adding horseradish peroxidase to a final concentration of 40 µM, and the fluorescence produced was measured in a spectrofluorophotometer (Shimadzu RF-540) at excitation and emission wavelengths of 315 and 425 nm, respectively. The H2O2 concentration was determined from a calibration curve of H2O2 (Merck) in the range 0.5–80 µM.
H2O2 cytochemistry:
The method used for the localization of H2O2 was based on the generation of precipitates of cerium perhydroxides (Bestwick et al., 1997), slightly modified by Romero-Puertas et al. (2004). Leaf pieces (c. 1 mm2) were incubated in freshly prepared 5 mM CeCl3 in 50 mM MOPS buffer, pH 7.2, for 1 h. As controls, samples were incubated separately with 25 µg µl–1 bovine liver catalase, and without CeCl3 (negative control). Leaf pieces were then fixed in 1.25% (v/v) glutaraldehyde and 1.25% (p/v) paraformaldehyde in 50 mM Na-cacodylate buffer, pH 7.2, for 1 h. Tissues were washed with the same buffer without fixatives, three times for 10 min, post-fixed for 1 h in 1% (v/v), OsO4, dehydrated in a graded ethanol series (30–100% v/v) and then embedded in Spurr resin. Ultra-thin sections were stained with uranyl acetate and lead citrate and examined in a Zeiss EM 10C transmission electron microscope.
Assay of protein-bound carbonyls
The spectrophotometric dinitrophenyl hydrazine (DNPH) method of Levine et al. (1994) was used for the determination of carbonyl groups. For each determination, two replicates and their respective blanks were measured. Samples containing at least 0.5 mg protein were incubated with 0.3% (v/v) Triton X-100 and 1% (w/v) streptomycin sulphate for 20 min to remove the nucleic acids, and centrifuged at 2000 g. Supernatants (200 µl) were mixed with 300 µl of 10 mM DNPH in 2 M HCl. The blank was incubated in 2 M HCl. After 1 h incubation at room temperature, proteins were precipitated with 10% (w/v) trichloroacetic acid and the pellets washed three times with 500 µl of ethanol:ethylacetate (1:1). The pellets were finally dissolved in 6 M guanidine hydrochloride in 20 mM potassium phosphate at pH 2.3, and the absorption at 370 nm was measured. Protein recovery was estimated for each sample by measuring the A280. Carbonyl content was calculated using a molar absorption coefficient for aliphatic hydrazones of 22 000 M–1 cm–1 (Levine et al., 1994).
Other assays
Protein concentrations were determined with the Bio-Rad microassay (Bio-Rad, Richmond, CA, USA), and chlorophyll and phaeophytin were estimated spectrophotometrically via phaeophytin, according to Vernon (1960). Lipid peroxidation was determined by measuring the concentration of thiobarbituric acid-reacting substances (TBARS) as described previously (Buege and Aust, 1972).
Statistical analysis
The significance of differences between mean values obtained from four samples produced in three to five independent experiments was determined by one-way analysis of variance. When the main effect was significant (P <0.05) differences between means were evaluated for significance by Duncan's multiple range test.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
The nodulated and nitrate-fed peas flowered at 9 weeks after sowing. After this stage leaf total soluble protein content and chlorophyll started to decline (Fig. 1). Leaf chlorophyll had fallen to very low levels in both sets of plants by 12 weeks, but leaf protein showed a much slower rate of decline (Fig. 1). The pattern of leaf protein loss showed a similar time-course to that of nodule proteins that has been reported previously (Groten et al., 2005). By 12 weeks the pods were at an advanced stage of development on both nitrate-fed and nodulated peas and the leaves had entered the late senescence stage, as illustrated by the decreased levels of chlorophyll at this time point (Fig. 1).
|
Nitrate-fed plants had a much greater shoot biomass than nodulated plants at both 9 and 12 weeks (Table 2). At the 9 week stage, the leaves on the nodulated peas had much lower photosynthetic rates than those supplied with nitrate, suggesting that senescence had already begun (Table 2). By contrast, the leaves of the nitrate-fed plants still retained relatively high rates of CO2 assimilation even at 12 weeks. At this point, photosynthesis was almost fully inhibited in the leaves of the nodulated peas. Transpiration rates remained high throughout this last stage of development (Table 2).
|
Ascorbate accumulated in the leaves of both sets of plants up to the 7 week stage, after which it began to decline (Fig. 1). The ascorbate pool was about 80% reduced up to week 7 in the nodulated plants and up to week 9 in the nitrate-fed peas, after which the ascorbate pool became rapidly oxidized in both sets of plants (Fig. 1). The decrease in leaf ascorbate and the increased oxidation of the pool occurred before changes in leaf chlorophyll or protein in both sets of plants, but this was most marked in the nodulated peas (Fig. 1). By contrast, the leaf glutathione pool increased up to 9 weeks in both sets of plants (Fig. 1). The glutathione pool was about 80–95% reduced up to week 11 in both sets of plants, after which point substantial oxidation of the pool occurred. After 9 weeks the total amount of glutathione in the leaves decreased (Fig. 1).
The activities of catalase and ascorbate–glutathione cycle enzymes were high until the later stages of development in both nitrate-fed and nodulated plants (Fig. 2). APX, GR, and MDHAR activities decreased at week 12 in both sets of plants (Fig. 2). By contrast, DHAR and catalase activities had decreased to very low levels at week 12. Some differences in the abundance of transcripts encoding antioxidant enzymes were observed at 12 weeks in the leaves from nitrate-fed and nodulated peas (Fig. 3). For example, APX, MDHAR, GR1, and Fe-SOD transcripts were higher in nitrate-fed plants than nodulated plants, whereas the levels of mRNAs encoding CuZn-SOD I and CuZn-SOD II were lower in nitrate-fed plants than in nodulated plants (Fig. 3). Other transcripts such as catalase, Mn-SOD, and GR2 were similar in abundance in both sets of plants (Fig. 3).
|
|
Leaf H2O2 contents were similar in nitrate-fed and nodulated peas at 9 weeks (Fig. 4). Values were much higher at 12 weeks than at 9 weeks in both sets of plants (Fig. 4). CeCl3-staining and transmission electron microscopy were used to determine the intracellular localization of H2O2 (Fig. 5). The precipitates of electron-dense cerium perhydroxide, indicating the presence of H2O2, were not uniformly distributed in any of the intracellular compartments, but showed a very patchy distribution. At 12 weeks, the leaves showed small localized pools of cerium perhydroxide in the chloroplasts and cell wall, and to a lower extent in mitochondria and peroxisomes (Fig. 5).
|
|
The number of protein carbonyl groups was similar in leaves of nitrate-fed and nodulated plants at 9 weeks (Fig. 6). Protein carbonyl groups had increased only slightly at 12 weeks in the leaves of plants supplied with nitrate (Fig. 6). By marked contrast, a very large increase in the number of protein carbonyl groups was found in the leaves of nodulated plants at 12 weeks (Fig. 6).
|
The extent of lipid peroxidation was similar in leaves of nitrate-fed and nodulated plants at 9 weeks (Fig. 6). Values had increased only slightly at 12 weeks in the leaves of plants supplied with nitrate (Fig. 6). However, a large increase in the content of lipid peroxidation products was found in the leaves of nodulated plants at 12 weeks (Fig. 6). Despite the marked increases in number of protein carbonyl groups and the content of lipid peroxidation products observed in the leaves of nodulated plants at 12 weeks, superoxide dismutase activities were similar in the leaves of nodulated plants and nitrate-fed plants at 9 weeks and had hardly changed at 12 weeks (Fig. 7).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Senescence is the final step in leaf development but it can be triggered prematurely by exposure to environmental stress or nutrient deprivation (Quirino et al., 2000; Lim et al., 2003). Mitochondria, chloroplasts and peroxisomes have roles in the senescence process. For example, when expression of the plastid ndhF gene was repressed in transgenic tobacco plants, leaf senescence was delayed relative to the wild type, leading to the hypothesis that chloroplasts have a major role in the regulation of leaf senescence (Zapata et al., 2005). Previous studies have shown that senescence in pea leaves is accompanied by enhanced cellular oxidation (Pastori and del Río, 1997; del Río et al., 1998; Jiménez et al., 1998). In the present study, the impact of the symbiotic union in the nodule in providing pea plants with reduced N compounds relative to the pathway of primary N assimilation were compared, together with effects on the processes contributing to the enhanced cellular oxidation that is characteristic of leaf senescence.
The nitrate-fed peas produced up to 50% more shoot biomass than the nodulated peas. The lower shoot biomass observed in the nodulated plants may result from the high cost of producing and maintaining the nodule. Changes in leaf protein and chlorophyll were used as markers of the senescence process. The leaves of both sets of plants contained similar amounts of protein and chlorophyll up to the 9 week stage after which both parameters rapidly declined. These results suggest that the timing of plant development and senescence is not greatly altered by the source of reduced N. However, the loss of protein and chlorophyll was more rapid in the nodulated plants compared with those supplied with nitrate at 11 weeks, suggesting that the progression of leaf senescence was much faster at this point in plant development. Despite having comparable amounts of protein and chlorophyll, the leaves of the nitrate-fed plants had much higher photosynthesis rates at week 9 than nodulated plants. These results illustrate that by week 9, photosynthetic carbon assimilation was not operating as efficiently in the nodulated plants as in those supplied with nitrate suggesting that, by week 9, the leaves of the nodulated plants have entered senescence. There is evidence to suggest that nodulated plants suffer from N deficiency at the pod-filling stage (Merbach and Schilling, 1980) and this might induce early leaf senescence. The peas studied here were flowering at 9 weeks and entering the pod-filling stage. The application of N to field-grown legumes often leads to greater seed yields with higher seed protein contents (Merbach and Schilling, 1980; Wesley et al., 1998). This observation suggests that the legume–Rhizobium symbiosis breaks down too soon in the plant development cycle and the plants become deprived of N, requiring faster remobilization from the leaves.
Although the leaves of nodulated plants retained less protein than the leaves of nitrate-fed plants during senescence, they still had substantial amounts of protein in their leaves at 12 weeks (Fig. 1). The leaf proteins accumulated during the seed-filling stage of development might be more important in defence against pathogens and abiotic stress protection than in processes associated with photosynthesis and assimilate production. The expression of pathogenesis-related proteins and pathogen resistance is associated with loss of leaf ascorbate (Barth et al., 2004; Kiddle et al., 2003; Pastori et al., 2003; Pavet et al., 2005), an important leaf metabolite that has been linked to leaf senescence (Kingston-Smith et al., 1997; Foyer, 2004). Ascorbate and glutathione are key antioxidants in plants acting in part through the control of ROS, as well as independent redox-signalling molecules (Noctor and Foyer, 1998; Foyer and Noctor, 2005a, b). As shown in Fig. 1 the changes in leaf protein during senescence were less marked than the changes in chlorophyll. The glutathione and ascorbate data were therefore expressed on a protein basis. Leaf ascorbate decreased significantly prior to leaf senescence in both nitrate-fed and nodulated plants. While glutathione was decreased in senescent leaves compared with young leaves, the leaf glutathione pool was nevertheless still significant, even in the oldest nodules. The endogenous pools of glutathione and other thiols are important in the regulation of senescence-induced proteinase activities in pea nodules (Groten et al., 2006). While the induction of proteinases is a key feature of leaf senescence, little information is available on the regulation of the activities of these proteins by cellular oxidants and reductants such as ascorbate and glutathione. The ubiquitin-dependent protein degradation pathway, which selectively targets a diverse range of substrates for degradation by the 26S proteosome, is important in the degradation of leaf proteins during senescence (Stone et al., 2005). The activity of this pathway is influenced by cellular reductants such as glutathione (Theriault et al., 2000). High GSH/GSSG ratios can prevent the binding of target proteins and regulation of the reduction state of protein thiols might participate in the regulation of this reaction cascade (El Yahyaoui et al., 2004). The oxidation observed in senescent leaves might, therefore, be predicted to favour activation of this protein degradation pathway.
The build-up of H2O2 in the leaf tissues during senescence is consistent with previously published results (del Río et al., 1998). The increase in leaf H2O2 levels occurs in parallel with increases in lipid peroxidation and protein oxidation, which were most marked in the leaves of nodulated plants. This result suggest that the extent of cellular oxidation is greater in the leaves of nodulated plants relative to those fed on nitrate. Contributing factors, other than those measured here, such as the activation of lipoxygenases, must account for the enhanced oxidation observed in the leaves of the nodulated peas.
Catalase activity decreased in senescent leaves of both nitrate-fed and nodulated plants after 9 weeks, and tissue H2O2 contents increased. The localization studies reported here show that H2O2 accumulated largely in the chloroplasts and in the cell wall. Little change in peroxisomal H2O2 was observed despite the measured decrease in catalase activity indicating that other peroxisomal peroxidases and/or non-enzymatic antioxidants probably take over the role of catalase. Indeed, there is a well-characterized peroxisomal APX, and other peroxisomal peroxidases, which have important metabolic functions in leaf senescence, as well as the glutathione pool of peroxisomes which is strongly increased by senescence (Jiménez et al., 1997, 1998; del Río et al., 1998, 2003). While catalase and DHAR activities decreased in senescent leaves, little change in the corresponding transcript abundance was observed. Similarly, the relative abundance of APX, MDHAR, cytosolic GR (GR1), and Fe-SOD transcripts was highest in nitrate-fed plants, but the activities of these enzymes were similar in both sets of plants. This suggests the possibility of post-translational modification of some antioxidant enzymes in senescent leaves, particularly those from nitrate-fed plants. By contrast with chloroplastic Fe-SOD, transcripts encoding cytosolic and chloroplastic CuZn-SODs were lower in senescent leaves from nitrate-fed plants.
Taken together, these results emphasize the role of oxidation in the orchestration of leaf senescence. This provides a parallel to the ageing process in other organisms, where antioxidants, antioxidant response element linked gene expression, and mitochondrial function are key players. Current evidence in plants and animals suggests that inappropriate production of oxidants and carbonyls, are intricately connected to ageing. It is probable that systemic signals, such as a deficit in amino acids or other metabolites associated with N metabolism, arising from the breakdown legume–Rhizobium symbiosis leads to early senescence and a more oxidized state of the leaf cells in the nodulated peas during senescence.
|
Acknowledgements |
|---|
This work was supported by a RTN grant of the European Commission (HPRN-CT-2000-00094) and the Junta de Andalucía (research group CVI 0192). HV and ML acknowledge contracts from the European Commission. The authors are grateful to Dr Peter Römer (Südwestdeutsche Saatzucht, Rastatt, Germany) for the supply of pea seeds cv. Phoenix. The electron microscopy assays were carried out at the Centre of Scientific Instrumentation of the University of Granada. Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council of the UK.
|
Footnotes |
|---|
* Present address: Institut de Biotechnologie des Plantes, Université Paris XI, Bâtiment 630, F-91405 Orsay cedex, France.
|
Abbreviations |
|---|
ASC, ascorbate; APX, ascorbate peroxidase; CAT, catalase; DHA, dehydroascorbate; DHAR, dehydroascorbate reductase; DNPH, 2,4-dinitrophenylhydrazine; GOX, glycolate oxidase; GR, glutathione reductase; GSH, reduced glutathione; GSSG, glutathione disulphide; MDHAR, monodehydroascorbate reductase; ROS, reactive oxygen species; SOD, superoxide dismutase, SNF symbiotic nitrogen fixation.
|
References |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Barth C, Moeder W, Klessig DF, Conklin PL. (2004) The timing of senescence and response to pathogens is altered in ascorbate-deficient mutant vitamin C-1. Plant Physiology 134:178–192.
Bestwick CS, Brown IR, Bennett MHR, Mansfield JW. (1997) Localization of hydrogen peroxide accumulation during the hypersensitive reaction of lettuce cells to Pseudomonas syringae pv. phaseolicola. The Plant Cell 9:209–221.[Abstract]
Buchanan-Wollaston V. (1997) The molecular biology of leaf senescence. Journal of Experimental Botany 48:181–199.
Buege JA and Aust SD. (1972) Microsomal lipid peroxidation. Methods in Enzymology 52:302–310.
Chen C and Dickman MB. (2004) Bcl-2 family members localize to tobacco chloroplasts and inhibit programmed cell death induced by chloroplast-targeted herbicides. Journal of Experimental Botany 55:2617–2623.
Creissen G, Firmin J, Fryer M, et al. (1999) Elevated glutathione biosynthetic capacity in the chloroplast of transgenic tobacco plants paradoxically causes increased oxidative stress. The Plant Cell 11:1277–1291.
del Río LA, Corpas FJ, Sandalio LM, Palma JM, Barroso JB. (2003) Plant peroxisomes, reactive oxygen metabolism and nitric oxide. IUBMB Life 55:71–81.[Web of Science][Medline]
del Río LA, Gómez-Ortega M, Leal-López A, López-Gorgé J. (1977) A more sensitive modification of the catalase assay with the Clark oxygen electrode. Analytical Biochemistry 80:409–415.[CrossRef][Web of Science][Medline]
del Río LA, Pastori GM, Palma JM, Sandalio LM, Sevilla F, Corpas FJ, Jiménez A, López-Huertas E, Hernández JA. (1998) The activated oxygen role of peroxisomes in senescence. Plant Physiology 116:1195–1200.
El Yahyaoui F, Küster H, Ben Amor B, et al. (2004) Expression profiling in Medicago truncatula identifies more than 750 genes differentially expressed during nodulation, including many potential regulators of the symbiotic program. Plant Physiology 136:3159–3176.
Foyer CH. (2004) The role of ascorbic acid in defence networks and signalling in plants. In Asard H, May JM, Smirnoff N (Eds.). Vitamin C: functions and biochemistry in animals and plants (Bios Scientific Publishers, Oxford, UK) pp. 65–82.
Foyer CH and Noctor G. (2005a) Oxidant and antioxidant signalling in plants: a re-evaluation of the concept of oxidative stress in a physiological context. Plant, Cell and Environment 28:1056–1071.[CrossRef]
Foyer CH and Noctor G. (2005b) Redox homeostasis and antioxidant signalling: a metabolic interface between stress perception and physiological responses. The Plant Cell 17:1866–1875.
Foyer CH, Rowell J, Walker D. (1983) Measurements of the ascorbate content of spinach leaf protoplasts and chloroplasts during illumination. Planta 157:239–244.
Gage DJ and Margolin W. (2000) Hanging by a thread: invasion of legume plants by rhizobia. Current Opinion in Microbiology 3:613–617.[CrossRef][Web of Science][Medline]
Griffith OW. (1980) Determination of glutathione and glutathione disulfide using glutathione reductase and 2-vinylpyridine. Analytical Biochemistry 106:207–212.[CrossRef][Web of Science][Medline]
Groten K, Dutilleul C, van Heerden PDR, Vanacker H, Bernard S, Finkemeier I, Dietz K-J, Foyer CH. (2006) Redox regulation of peroxiredoxin and proteinases by ascorbate and thiols during pea root nodule senescence. FEBS Letters 580:1269–1276.[CrossRef][Web of Science][Medline]
Groten K, Vanacker H, Dutilleul C, Bastian F, Bernard S, Carzaniga R, Foyer CH. (2005) The roles of redox processes in pea nodule development and senescence. Plant, Cell and Environment 28:1293–1304.[CrossRef]
Hensel LL, Grbic V, Baumgarten DA, Bleecker AB. (1993) Developmental and age-related processes that influence the longevity and senescence of photosynthetic tissues in Arabidoposis. The Plant Cell 5:553–564.[Abstract]
Jiménez A, Hernández JA, del Río LA, Sevilla F. (1997) Evidence for the presence of the ascorbate glutathione cycle in mitochondria and peroxisomes of pea leaves. Plant Physiology 114:275–284.[Abstract]
Jiménez A, Hernández JA, Pastori GM, del Río LA, Sevilla F. (1998) Role of the ascorbate–glutathione cycle of mitochondria and peroxisomes in the senescence of pea leaves. Plant Physiology 118:1327–1335.
Kiddle G, Pastori GM, Bernard B, Pignocchi C, Antoniw J, Verrier PJ, Foyer CH. (2003) Effects of leaf ascorbate content on defense and photosynthesis gene expression in Arabidopsis thaliana. Antioxidants and Redox Signalling 5:23–32.[CrossRef][Web of Science][Medline]
Kingston-Smith AH, Thomas H, Foyer CH. (1997) Chlorophyll a fluorescence, enzyme and antioxidant analyses provide evidence for the operation of alternative electron sinks during leaf senescence in a ‘stay-green’ mutant of Festuca pratensis. Plant, Cell and Environment 20:1323–1337.[CrossRef]
Laloi C, Apel K, Danon A. (2004) Reactive oxygen signalling: the latest news. Current Opinion in Plant Biology 7:323–328.[CrossRef][Web of Science][Medline]
Levine RL, Williams JA, Stadtman ER, Shacter E. (1994) Carbonyl assays for determination of oxidatively modified proteins. Methods in Enzymology 233:346–363.[Web of Science][Medline]
Lim PO, Woo HR, Nam HG. (2003) Molecular genetics of leaf senescence in Arabidopsis. Trends in Plant Science 8:272–278.[CrossRef][Web of Science][Medline]
Long SP and Hällgren JE. (1985) Measurement of CO2 assimilation by plants in the field and the laboratory. In Coombs J, Hall DO, Long SP, Scurlock JMO (Eds.). Techniques in bioproductivity and photosynthesis (Pergamon Press, Oxford, UK) pp. 62–94.
Ludlow MM and Muchow RC. (1990) A critical evaluation of trails for improving crops yield in water-limited environments. Advances in Agronomy 43:107–153.
McCord JM and Fridovich I. (1969) Superoxide dismutase: an enzymic function for erythrocuprein (hemocuprein). Journal of Biological Chemistry 244:6049–6055.
Merbach W and Schilling G. (1980) Effectiveness of symbiotic N2-fixation in leguminous plants, as affected by inoculation with rhizobia, by substrate, N-fertilizing, and 14C-sucrose application. Zentralblatt Bakteriologie Naturwissenschaften 135:99–118.
Nam HG. (1997) Molecular genetic analysis of leaf senescence. Current Opinion in Biotechnology 8:200–207.[CrossRef][Web of Science][Medline]
Navabpour S, Morris K, Allen R, Harrison E, A-H-Mackerness S, Buchanan-Wollaston V. (2003) Expression of senescence-enhanced genes in response to oxidative stress. Journal of Experimental Botany 54:2285–2292.
Noctor G and Foyer CH. (1998) Ascorbate and glutathione: keeping active oxygen under control. Annual Review of Plant Physiology and Plant Molecular Biology 49:249–279.[CrossRef][Web of Science]
Olivares J, Casadesus J, Bedmar EJ. (1980) Method for testing degree of infectivity of Rhizobium meliloti strains. Applied and Environmental Microbiology 56:389–393.
Palma JM, Sandalio LM, Corpas FJ, Romero-Puertas MC, McCarthy I, del Río LA. (2002) Plant proteases, protein degradation, and oxidative stress: role of peroxisomes. Plant Physiology and Biochemistry 40:521–530.[CrossRef]
Pastori GM and del Río A. (1997) Natural senescence of pea leaves: an activated oxygen-mediated function for peroxisomes. Plant Physiology 113:411–418.[Abstract]
Pastori GM, Kiddle G, Antoniw J, Bernard S, Veljovic-Jovanovic S, Verrier PJ, Noctor G, Foyer CH. (2003) Leaf vitamin C contents modulate plant defense transcripts and regulate genes that control development through hormone signaling. The Plant Cell 15:939–951.
Pavet V, Olmos E, Kiddle G, Mowla S, Kumar S, Antoniw J, Alvarez ME, Foyer CH. (2005) Ascorbic acid deficiency activates cell death and disease resistance responses in Arabidopsis thaliana. Plant Physiology 139:1291–1303.
Quirino BF, Noh YS, Himelblau E, Amasino RM. (2000) Molecular aspects of leaf senescence. Trends in Plant Science 5:278–282.[CrossRef][Web of Science][Medline]
Romero-Puertas MC, Rodríguez-Serrano M, Corpas FJ, Gómez M, del Río LA, Sandalio LM. (2004) Cadmium-induced subcellular accumulation of 
and H2O2 in pea leaves. Plant, Cell and Environment 27:1122–1134.[CrossRef]
Schultze M and Kondorosi A. (1998) Regulation of symbiotic root nodule development. Annual Review of Genetics 32:33–57.[CrossRef][Web of Science][Medline]
Stone SL, Hauksdottir H, Troy A, Herschleb J, Kraft E, Cullis J. (2005) Functional analysis of the RING-type ubiquitin ligase family of Arabidopsis. Plant Physiology 137:317–327.
Theriault A, Wang Q, Van Iderstine SC, Chen B, Franke AA, Adeli K. (2000) Modulation of hepatic lipoprotein synthesis and secretion by taxifolin, a plant flavonoid. Journal of Lipid Research 41:1969–1979.
Thomas H, Ougham HJ, Wagstaff C, Stead AD. (2003) Defining senescence and death. Journal of Experimental Botany 54:1127–1132.
Thompson JE, Froese CD, Madey E, Smith MD, Hong Y. (1999) Lipid metabolism during plant senescence. Progress in Lipid Research 37:119–141.
Vanacker H, Carver TLW, Foyer CH. (1998) Pathogen-induced changes in the antioxidant status of the apoplast in barley leaves. Plant Physiology 117:1103–1114.
Vernon LP. (1960) Spectrophotometer determination of chlorophylls and pheophytins in plant extracts. Analytical Chemistry 32:1144–1150.[CrossRef]
Wagner D, Przybyla D, Op den Camp R, et al. (2004) The genetic basis of singlet oxygen-induced stress responses of Arabidopsis thaliana. Science 306:1183–1185.
Wesley TL, Lamond RE, Martin VL, Duncan SR. (1998) Effects of late-season nitrogen fertilization on irrigated soybean yield and composition. Journal of Production Agriculture 11:331–336.
Zapata JM, Guéra A, Esteban-Carrasco A, Martín M, Sabater B. (2005) Chloroplasts regulate leaf senescence: delayed senescence in transgenic ndhF-defective tobacco. Cell Death and Differentiation 12:1277–1284.[CrossRef][Web of Science][Medline]
CiteULike 
Connotea 
Del.icio.us What's this?
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||