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First published online August 4, 2006; 10.1105/tpc.106.041541

The Plant Cell 18:2356-2368 (2006)
© 2006 American Society of Plant Biologists

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Rice NTRC Is a High-Efficiency Redox System for Chloroplast Protection against Oxidative Damage[W]

Juan Manuel Pérez-Ruiza, María Cristina Spínolaa, Kerstin Kirchsteigera, Javier Morenob, Mariam Sahrawyc and Francisco Javier Cejudoa,1

a Instituto de Bioquímica Vegetal y Fotosíntesis, Centro de Investigaciones Científicas Isla de la Cartuja, 41092 Seville, Spain
b Departamento de Biología Celular, Facultad de Biología, Universidad de Sevilla, 41013 Seville, Spain
c Departamento de Bioquímica y Biología Molecular de Plantas, Estación Experimental del Zaidín, 18008 Granada, Spain

1 To whom correspondence should be addressed. E-mail fjcejudo{at}us.es; fax 34-954460-0065.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 METHODS
 REFERENCES
 
One of the mechanisms plants have developed for chloroplast protection against oxidative damage involves a 2-Cys peroxiredoxin, which has been proposed to be reduced by ferredoxin and plastid thioredoxins, Trx x and CDSP32, the FTR/Trx pathway. We show that rice (Oryza sativa) chloroplast NADPH THIOREDOXIN REDUCTASE (NTRC), with a thioredoxin domain, uses NADPH to reduce the chloroplast 2-Cys peroxiredoxin BAS1, which then reduces hydrogen peroxide. The presence of both NTR and Trx-like domains in a single polypeptide is absolutely required for the high catalytic efficiency of NTRC. An Arabidopsis thaliana knockout mutant for NTRC shows irregular mesophyll cell shape, abnormal chloroplast structure, and unbalanced BAS1 redox state, resulting in impaired photosynthesis rate under low light. Constitutive expression of wild-type NTRC in mutant transgenic lines rescued this phenotype. Moreover, prolonged darkness followed by light/dark incubation produced an increase in hydrogen peroxide and lipid peroxidation in leaves and accelerated senescence of NTRC-deficient plants. We propose that NTRC constitutes an alternative system for chloroplast protection against oxidative damage, using NADPH as the source of reducing power. Since no light-driven reduced ferredoxin is produced at night, the NTRC-BAS1 pathway may be a key detoxification system during darkness, with NADPH produced by the oxidative pentose phosphate pathway as the source of reducing power.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 METHODS
 REFERENCES
 
Oxygenic photosynthesis is the source of atmospheric oxygen and provides organic material for virtually all life forms (Nelson and Ben-Shem, 2004Go). However, photosynthesis involves transport of electrons in the presence of oxygen; therefore, it is a process that inevitably produces reactive oxygen species (ROS), which are harmful for the cell (Mittler et al., 2004Go). Consequently, the evolution of photosynthesis has occurred along with the evolution of nonenzymatic and enzymatic ROS-scavenging mechanisms (Ledford and Niyogi, 2005Go). Nonenzymatic antioxidants include glutathione, ascorbate, carotenoids, or tocopherols, whereas enzymatic scavenging mechanisms include ascorbate peroxidase, superoxide dismutase, glutathione peroxidase (GPX), and catalase (Apel and Hirt, 2004Go). Plants contain gene families for superoxide dismutase, ascorbate peroxidase, and GPX. The isoforms encoded by the members of these gene families are targeted to different subcellular locations, cytosol, mitochondria, and chloroplasts, as has been shown for GPX genes in Arabidopsis thaliana (Rodriguez Milla et al., 2003Go). By contrast, catalase is mainly localized in peroxisomes (Apel and Hirt, 2004Go).

The ROS produced by the chloroplast include singlet oxygen, superoxide and hydroxyl radicals, and hydrogen peroxide (Mittler, 2002Go). Of these species, singlet oxygen is highly reactive and probably generates oxidative damage near the site of its production (Ledford and Niyogi, 2005Go). By contrast, superoxide radicals are rapidly converted to hydrogen peroxide in a reaction catalyzed by superoxide dismutase (Asada, 1999Go). Hydrogen peroxide is a relatively stable and permeable molecule with a double effect for the cell. When it accumulates at a high level, it becomes toxic because it can be converted to highly reactive hydroxyl radicals (Apel and Hirt, 2004Go); in addition, hydrogen peroxide is an important signaling molecule (Wood et al., 2003aGo). Indeed, global expression analysis revealed that hydrogen peroxide is involved in the regulation of a large number of genes in tobacco (Nicotiana tabacum) (Vandenabeele et al., 2003Go) and Arabidopsis (Vanderauwera et al., 2005Go). As a signaling molecule, hydrogen peroxide, most of which is produced by the chloroplast in plant cells, is involved in the regulation of several processes, including photosynthesis (Pfannschmidt, 2003Go) and the response of the plant to different environmental stimuli (Laloi et al., 2004Go).

Therefore, the mechanism to control the production of hydrogen peroxide by chloroplasts is important to balance its signaling role and deleterious effects. In this regard, peroxiredoxins, a family of antioxidant enzymes able to reduce hydrogen or organic peroxides, play an important role (Dietz, 2003Go; Wood et al., 2003bGo). Of this family of enzymes, 2-Cys peroxiredoxins (2-Cys Prxs) have been proposed to play a key role as antioxidants and for regulating the level of hydrogen peroxide for signal transduction (Wood et al., 2003aGo). The catalytic mechanism of 2-Cys Prxs involves the formation of a disulfide, which needs to be reduced for a new catalytic cycle. In prokaryotes, this reduction is catalyzed by an NADH-dependent peroxiredoxin oxidoreductase termed AhpF (Poole et al., 2000aGo). In eukaryotes, however, no homologue of AhpF has been described so far, and it is considered that 2-Cys Prxs are reduced by a two-component system formed by thioredoxins (Trxs) and NADPH thioredoxin reductase (NTR).

Previous analysis of the chloroplast detoxification system identified a 2-Cys Prx, termed BAS1, involved in chloroplast protection against oxidative damage (Baier and Dietz, 1997Go). The redox state of BAS1 depends on the NAD(P)H system, and oxidized BAS1 can be reduced by plastid thioredoxins (Konig et al., 2002Go), among which, x-type Trx is the most efficient electron donor (Collin et al., 2003Go). In addition, in vitro and in vivo interaction of the plastid Trx CDSP32 with BAS1 has been shown (Broin et al., 2002Go; Rey et al., 2005Go). Furthermore, BAS1 displayed CDSP32-dependent peroxidase activity, though the rate of peroxide reduction catalyzed by BAS1 in the presence of CDSP32 (Broin et al., 2002Go) was much lower than in the presence of Trx x (Collin et al., 2003Go). Based on these results, it was proposed that CDSP32 and Trx x are physiological electron donors to BAS1, the expected source of reducing power being ferredoxin reduced by the photosynthetic electron chain. In this regard, Arabidopsis mutants lacking the variable subunit of the ferredoxin thioredoxin reductase (FTR) are more sensitive to oxidative damage than the wild type (Keryer et al., 2004Go), suggesting that the FTR-Trx (type-x and CDSP32) system is involved in chloroplast protection against oxidative damage.

Recently, we identified a gene in Arabidopsis and rice (Oryza sativa) encoding a novel type of NADPH thioredoxin reductase, termed NTRC (Serrato et al., 2004Go). The gene was cloned from rice, and the encoded protein, NTRC, was shown to be a bimodular enzyme formed by an NTR-like module at the N terminus and a Trx-like module at the C terminus. Since NTRC is localized to the chloroplast stroma and shows a high affinity for NADPH (Serrato et al., 2004Go), we hypothesized that this enzyme might function as an alternative electron donor for the chloroplast detoxification system, allowing the use of NADPH for such a protective role. To test this possibility, homologs of CDSP32, Trx x, and the 2-Cys Prx BAS1 were cloned from rice and expressed in Escherichia coli. Here, we show that rice NTRC is a high-efficiency system to transfer electrons from NADPH to the plastid 2-Cys Prx (BAS1) from rice. An Arabidopsis knockout mutant for NTRC shows abnormal chloroplast structure, reduced content of photosynthetic pigments, and a lower rate of photosynthesis. The mutant also is hypersensitive to prolonged darkness. Based on these results, we propose that NTRC is an alternative system, which allows the use of NADPH for chloroplast detoxification.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 METHODS
 REFERENCES
 
Cloning and Expression in E. coli of the 2-Cys Prx, BAS1, and the Thioredoxins CDSP32 and Trx x from Rice
We have previously described NTRC as a bifunctional enzyme showing NTR and Trx activity. However, NTRC was unable to catalyze the NADPH-dependent reduction of insulin; that is, NTRC did not function as an NTR/Trx system under these assay conditions (Serrato et al., 2004Go). Given that NTRC is localized to the chloroplast stroma and shows a high affinity for NADPH, we tested its possible role as an alternative electron donor to the chloroplast detoxification system and its interaction with the FTR-Trx (Trx x and CDSP32) system. To that end, the plastid 2-Cys Prx, BAS1, and thioredoxins CDSP32 and Trx x were cloned from rice, expressed in E. coli with a His-tag at the N terminus, and purified by nitrilotriacetic acid (NTA) chromatography (Figure 1A ).


Figure 1
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  		Figure 1. Purification of Recombinant Rice BAS1, CDSP32, and Trx x, and Thioredoxin Activity Assays.

(A) Rice homologs of the 2-Cys Prx (BAS1), the plastid CDSP32, a truncated polypeptide containing the C-terminal Trx fold of CDSP32 (CDSP32-C), and Trx x were expressed in E. coli as N-terminal His-Tagged proteins and purified by NTA chromatography. Purified proteins (2.5 µg) were subjected to SDS-PAGE under reducing conditions. Molecular mass markers (kD) are on the left.

(B) Thioredoxin activity was assayed as reduction of insulin in a reaction mixture containing 100 mM potassium phosphate buffer, pH 7.0, 2 mM EDTA, 0.5 mg mL–1 insulin, and 0.5 mM DTT. Trx x (closed squares), Trx hA (open squares), CDSP32-C (open triangles), and CDSP32 (closed triangles) were added at 5 µM final concentration. A negative control in the absence of thioredoxin was performed (closed circles).

 
The thioredoxin activity of the purified His-tagged Trx x and CDSP32 was determined with the insulin reduction assay in the presence of DTT. Trx x showed a high rate of insulin reduction, even higher than wheat (Triticum aestivum) Trx hA (Serrato et al., 2001Go), which was used as a positive control (Figure 1B). However, CDSP32 displayed a rather low insulin reduction activity compared with Trx x and Trx hA (Figure 1B). It should be noted that though rice CDSP32 showed low activity, it was detectable, in contrast with CDSP32 from Arabidopsis, which did not show any insulin reduction activity (Broin et al., 2002Go). In addition, the C-terminal domain of CDSP32 was produced in E. coli as an N-terminal His-tagged protein (Figure 1A). This truncated polypeptide, containing a Trx active site, showed higher insulin reduction activity than the full-length protein CDSP32 (Figure 1B), in agreement with previous results showing that the C-terminal Trx fold of CDSP32 from Arabidopsis, but not the full-length enzyme, displayed insulin reduction activity (Broin et al., 2002Go).

NTRC Diverts Reducing Power to the Chloroplast Detoxification System
To test the possible function of NTRC as electron donor to the chloroplast detoxification system, purified recombinant NTRC and BAS1 were incubated in the presence of NADPH and hydrogen peroxide. Figure 2A shows that the system formed by NTRC and BAS1 efficiently catalyzed the oxidation of NADPH when hydrogen peroxide was present. No activity was observed in the absence of NTRC, and a residual rate of NADPH oxidation occurred in the absence of BAS1. This residual activity was also detected in the absence of hydrogen peroxide and therefore is most probably due to diaphorase activity, characteristic of flavoproteins, of NTRC. The initial rate of NADPH oxidation by the NTRC-BAS1 system (40.3 µmol NADPH min–1 µmol–1 BAS1) was remarkably higher than the rate obtained with standard two-component assay conditions in the presence of wheat NTR and Trx hA (2.1 µmol NADPH min–1 µmol–1 BAS1). NTRC activity was confirmed by direct determination of hydrogen peroxide in the in vitro assay (Figure 2B). The residual activity of NTRC in the absence of BAS1, determined as NADPH oxidation (Figure 2A), was not detected when activity was assayed as hydrogen peroxide reduction (Figure 2B), lending further support to the proposal that it is due to the diaphorase activity of NTRC.


Figure 2
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  		Figure 2. Hydrogen Peroxide Reduction by NTRC and BAS1.

(A) and (B) NTRC-BAS1 was assayed as NADPH oxidation (A) or hydrogen peroxide reduction (B) in a reaction mixture containing 100 mM phosphate buffer, pH 7.0, 2 mM EDTA (omitted for H2O2 determination), 0.25 mM NADPH, 0.5 mM hydrogen peroxide, and purified enzymes at the following concentrations: 2 µM NTRC plus 6 µM BAS1 (closed squares), 6 µM BAS1 (open squares), or 2 µM NTRC (closed diamonds).

(C) The same conditions were used to determine the activity of the modules of NTRC, which were added at the following concentrations: 2 µM Trx-like module plus 6 µM BAS1 (closed circles), 2 µM NTR module plus 6 µM BAS1 (open squares), 2 µM Trx-like module plus 2 µM NTR module plus 6 µM BAS1 (open triangles), or 2 µM NTRC plus 6 µM BAS1 (closed squares).

 
To characterize the biochemical properties of this new enzyme further, the activity of truncated polypeptides containing either the NTR or the Trx-like domain of NTRC was analyzed. Previously, it was shown that both truncated proteins show NTR and thioredoxin activity (Serrato et al., 2004Go). When assayed in the presence of BAS1, the Trx-like module did not show any activity (Figure 2C). By contrast, the NTR module showed a low rate of NADPH oxidation (Figure 2C). This activity was also detected in the absence of hydrogen peroxide, thus confirming that the diaphorase activity of NTRC is localized in the NTR module of the enzyme, which contains FAD. Activity assays with a mixture of NTR and Trx-like modules showed a higher rate of NADPH oxidation than the NTR module alone, but very low when compared with the activity of the full-length NTRC (Figure 2C). These results show that the native conformation of NTRC with both NTR and Trx domains in a single polypeptide is required for full activity and lends further support to the proposal that NTRC is a high-efficiency system for 2-Cys Prx-mediated hydrogen peroxide reduction.

The Rice 2-Cys Prx, BAS1, Is Efficiently Reduced by NTRC
According to the mechanism of catalysis proposed for 2-Cys Prxs (Dietz, 2003Go; Konig et al., 2003Go; Wood et al., 2003bGo), the results described above suggest that NTRC might be able to reduce the disulfides formed by the Cys residues (Cys-61 and Cys-183) at the active site of BAS1.This possibility was analyzed in two ways. First, a BAS1 mutant was produced, replacing Cys-61 by Ser. In SDS-PAGE analysis under nonreducing conditions, wild-type BAS1 was detected in dimeric form, whereas BAS1(C61S) was detected as a monomer (Figure 3A ). No activity was observed when wild-type NTRC was incubated with BAS1(C61S) mutant, the residual oxidation of NADPH being due to the diaphorase activity of NTRC (Figure 3B). Considering that, as stated before, NTRC acts as an NTR/Trx system, it should be expected that the active site of the NTR and Trx-like modules of the enzyme are involved in the redox interchange with BAS1. To test this possibility, the Cys residues at the active sites of the NTR and Trx-like modules of NTRC were replaced by Ser, producing NTRC mutants C140S and C143S (NTR domain) and C377S and C380S (Trx-like domain). None of these mutants showed activity when incubated with wild-type BAS1 (Figure 3B; data not shown), hence confirming that the active sites of both NTR and Trx-like modules of NTRC are essential for activity.


Figure 3
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  		Figure 3. Effect of Active Site Cys Mutations on NTRC-BAS1 Activity.

(A) Purified BAS1 wild type and mutant C61S (10 µg of protein) were subjected to SDS-PAGE under nonreducing conditions. Molecular mass markers (kD) are on the left.

(B) NADPH-dependent reduction of hydrogen peroxide activity was assayed in a reaction mixture containing 100 mM phosphate buffer, pH 7.0, 2 mM EDTA, 0.25 mM NADPH, and 0.5 mM hydrogen peroxide in the presence of 2 µM NTRC plus 6 µM BAS1 (closed squares), 2 µM NTRC plus 6 µM BAS1(C61S) (closed triangles), or 2 µM NTRC(C377S) plus 6 µM BAS1 (open triangles).

 
The second approach consisted in the determination of the direct reduction of BAS1 by NTRC. Given that active site Cys-61 and Cys-183 are the only Cys residues of BAS1, it was possible to distinguish reduced and oxidized BAS1 by 4-acetamido-4'-maleimidylstilbene (AMS) labeling of sulfhydryl groups. AMS did not label the oxidized, dimeric form of BAS1 (Figure 4A , lane 1). A similar result was obtained when NADPH was added in the absence of NTRC (Figure 4A, lane 2) or when NTRC was added in the absence of NADPH (Figure 4A, lane 3). However, when BAS1 was preincubated with the complete system, NTRC and NADPH, BAS1 was partially reduced as shown by the shift of the BAS1 band (Figure 4A, lane 4). BAS1 was also detected in monomeric form, which corresponds to the fully reduced form of the enzyme, suggesting that NTRC is able to use NADPH to reduce the two disulfides of BAS1. The different electrophoretical mobility of NTRC depending on the presence or absence of NADPH should be noted. In the presence of NADPH, most of the enzyme was detected as a band corresponding to the monomer (Figure 4A, lane 4), which was almost undetectable in the absence of NADPH (Figure 4A, lane 3). To test whether the difference of electrophoretical mobility was a direct effect of NADPH, purified NTRC was preincubated in the presence or absence of NADPH and then subjected to electrophoresis. Figure 4B shows that NTRC was detected in monomeric and dimeric forms in the presence of NADPH; however, these forms were almost undetectable in the absence of NADPH because most of the enzyme was detected as a high-molecular band at the upper part of the gel (Figure 4B). These results suggest that NADPH affects the aggregation state of NTRC, but the mechanism of this effect of NADPH is not yet understood.


Figure 4
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  		Figure 4. Reduction of BAS1 by NTRC.

(A) Reduction level of BAS1 was determined by AMS labeling. Purified BAS1 (3.5 µg) was incubated with 6.25 mM hydrogen peroxide for oxidation, then the protein was diluted 12.5-fold with 100 mM phosphate buffer, pH 7.0, and 2 mM EDTA and incubated in the presence of 0.45 mM NADPH and/or 7.5 µM NTRC as indicated. Molecular mass markers (kD) are on the left.

(B) Purified NTRC (4.5 µg) was preincubated in the presence or absence of 1 mM NADPH for 4 min at room temperature and subjected to SDS-PAGE. Samples were not boiled before loading. d, dimer; m, monomer; o, oligomer.

 
NTRC Is the Most Efficient Reductant of BAS1
Previous reports described two plastidial thioredoxins, CDSP32 and Trx x, as electron donors to the 2-Cys Prx, and it was proposed that the source of reducing power for this system is photosynthetically reduced ferredoxin and FTR (Broin et al., 2002Go; Collin et al., 2003Go). The activity of NTRC as an NTR/Trx system able to use NADPH to reduce the 2-Cys Prx BAS1 suggests that this may be an alternative system to transfer reducing power from NADPH to the chloroplast detoxification system. We therefore established which of the systems is more efficient as a reductant of 2-Cys Prx. Since both Trx x and CDSP32 were assayed in the presence of DTT (Broin et al., 2002Go; Collin et al., 2003Go), we determined the DTT-dependent activity of NTRC, corresponding to the Trx-like module (Serrato et al., 2004Go), to allow a direct comparison. Under these assay conditions, NTRC was the most efficient electron donor to BAS1 (Figure 5 ). In addition, these assays confirm that Trx x was a more efficient reductant of BAS1 than CDSP32, which showed almost undetectable activity. In the presence of NADPH, its physiological electron donor, NTRC proved to be by far the most efficient reductant (Figure 5). The kinetic parameters confirmed the high efficiency of NTRC, which showed higher affinity for BAS1 than Trx x, as deduced of the lower Km value (Table 1 ). Similarly, the catalytic efficiency (Kcat/Km) of NTRC was ~100-fold higher (Table 1). Although Trx x kinetic parameters were determined with a nonphysiological assay using DTT, these results highly suggest that the presence of both NTR and Trx activity in a single polypeptide increases the efficiency of the system.


Figure 5
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  		Figure 5. Comparison of NTRC, Trx x, and CDSP32 Activities.

DTT-dependent reduction of t-BOOH was determined with the Peroxoquant method in reaction mixtures containing 100 mM phosphate buffer, pH 7.0, 75 µM t-BOOH, 0.5 mM DTT, and 15 µM BAS1 without added thioredoxins (closed circles) or with CDSP32 (open triangles), Trx x (closed triangles), or NTRC (open squares) at 7.5 µM final concentration. The NADPH-dependent assay of NTRC (closed squares) contained 100 mM phosphate buffer, pH 7.0, 75 µM t-BOOH, 0.25 mM NADPH, 6 µM BAS1, and 3 µM NTRC.

 

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  		Table 1. Kinetic Parameters of Rice NTRC and Trx x with BAS1

 
An Arabidopsis NTRC Knockout Mutant Shows Damage of the Photosynthetic Apparatus
The in vitro assay results shown above suggest that NTRC is an alternative system able to divert reducing power (NADPH) to the 2-Cys Prx (BAS1) involved in chloroplast detoxification. To test the functional relevance of this system, we characterized the photosynthetic phenotype of an Arabidopsis knockout mutant deficient in NTRC (Serrato et al., 2004Go). The content of photosynthetic pigments, chlorophylls a and b, and carotenoids was significantly lower in the ntrc knockout plants (Table 2 ). The rate of electron transfer, determined in vitro as oxygen production by thylakoid fractions in the presence of ferricyanide, was severely reduced in the knockout mutant (Table 2). In addition, we analyzed the rate of photosynthesis in vivo, as CO2 fixation, at different light intensities. As expected, the ntrc mutant showed a lower rate of CO2 fixation, the difference being highly dependent on light intensity (Figure 6 ). Indeed, at low intensity (100 µmol m–2 s–1), the rate of photosynthesis of the ntrc mutant was negative, meaning that it was lower than the rate of respiration, whereas at higher intensity (500 to 1500 µmol m–2 s–1), the ntrc mutant presented slightly lower CO2 fixation than the wild type. So, NTRC deficiency seems to exert a stronger effect on CO2 fixation at lower light intensity.


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  		Table 2. Effect of NTRC Deficiency on the Content of Photosynthetic Pigments and the Rate of Photosynthesis

 

Figure 6
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  		Figure 6. Light-Dependent CO2 Fixation Response Curves of Wild-Type, Knockout, and Transgenic Plants.

Wild-type, NTRC-deficient Arabidopsis, and transgenic lines expressing NTRC under the control of the 35S promoter as indicated were grown for 30 d. Six determinations were performed per light intensity with leaves from different plants, and the mean values ± SD are represented. PAR, photosynthesis active radiation.

 
To test that the phenotype of the knockout mutant was due to NTRC deficiency, the wild-type NTRC gene was expressed under the control of the cauliflower mosaic virus 35S promoter in ntrc knockout and wild-type Arabidopsis plants. Two transgenic lines showing a high expression of the NTRC gene, as detected by RT-PCR analysis (Figure 7A ), were analyzed in detail. Despite the higher accumulation of NTRC transcripts in both transgenic lines, the amount of NTRC polypeptide was similar in leaves from wild-type and transgenic lines (Figure 7B), which suggests a posttranscriptional regulation of the amount of NTRC enzyme in vivo. The transgenic mutant plants expressing wild-type NTRC, KO:P35SNTRC, showed a growth indistinguishable from the wild-type plants or the wild-type plants expressing the NTRC gene, WT:P35SNTRC (Figure 7C).


Figure 7
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  		Figure 7. Expression of the NTRC Gene in the Arabidopsis Wild Type and ntrc Knockout Mutant.

Full-length NTRC cDNA was expressed in Arabidopsis wild-type and T-DNA insertion lines under the control of the cauliflower mosaic virus 35S promoter.

(A) Gene expression was analyzed by RT-PCR with RNA isolated from leaves of 35-d-old plants.

(B) Aliquots (25 µg of protein) from leaves of the different plant lines after 35 d of growth were subjected to SDS-PAGE, electrotransferred to nitrocellulose sheets, and probed with anti-NTRC or anti-Rubisco antibodies as indicated.

(C) Rosette leaves from wild-type, ntrc mutant, and transgenic plants grown for 27 d.

 
Mesophyll cells of the ntrc knockout mutant show an irregular shape and a variable content of chloroplasts compared with the wild-type cells (Figures 8A and 8B ). Moreover, NTRC deficiency caused abnormal chloroplast structure, which affected the content and organization of thylakoids (Figures 8E and 8F) compared with chloroplasts from wild-type cells (Figure 8D). Accordingly, mesophyll cells (Figure 8C) and chloroplast structure (Figure 8G) of KO:P35SNTRC transgenic plants showed wild-type morphology. Moreover, KO:P35SNTRC and WT:P35SNTRC transgenic plants showed a wild-type level of photosynthetic pigments and rate of oxygen production by isolated thylakoids (Table 2), as well as rate of photosynthesis determined as CO2 fixation (Figure 6). Therefore, these results show that the mutant phenotype was rescued by expression of wild-type NTRC.


Figure 8
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  		Figure 8. Mesophyll Cells and Chloroplast Ultrastructure from Wild-Type, NTRC-Deficient, and KO:P35SNTRC Arabidopsis Plants.

(A) to (C) Arabidopsis plants were grown for 29 d, and semithin sections from the wild type (A), NTRC-deficient mutant (B), and KO:P35SNTRC (C) were stained with Toluidine blue. Bars = 20 µm

(D) to (G) Representative chloroplast structure of wild-type (D), NTRC-deficient mutant ([E] and [F]), and KO:P35SNTRC transgenic plants (G). Bars = 1 µm.

 
Effect of Environmental Stresses on Arabidopsis Wild-Type and ntrc Knockout Plants
To analyze in more detail the function of NTRC, Arabidopsis wild-type and ntrc knockout plants were subjected to different environmental stresses, and the photosystem II (PSII) photochemical efficiency, determined as Fv/Fm, was used to determine the plant response to stress treatments. The PSII photochemical efficiency of the ntrc knockout was slightly lower than the wild-type plants under standard growth conditions (Table 3 ). Previously, we have shown that treatments with salt, drought, or methyl viologen caused growth inhibition on the ntrc mutant (Serrato et al., 2004Go); however, these treatments, as well as cold, produced a low effect on the PSII photochemical efficiency in the mutant (Table 3). Since the rate of CO2 fixation of the ntrc knockout was more affected at lower light intensity (Figure 6), we analyzed the effect of continuous darkness. This treatment produced a clear decrease of the photochemical efficiency both in wild-type and mutant plants, though the ntrc knockout was more sensitive (Table 3).


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  		Table 3. Effect of Abiotic Stress Treatments on PSII Photochemical Efficiency in Arabidopsis Wild-Type and NTRC-Deficient Plants

 
NTRC Deficiency Causes Hypersensitivity to Prolonged Darkness
During darkness, when there is no photosynthetic electron transport, NADPH in chloroplasts is still produced by the initial reactions of the oxidative pentose phosphate pathway (Neuhaus and Emes, 2000Go). So, NTRC, which uses NADPH to reduce the 2-Cys Prx, may become a key reductant system during the night. In this regard, it should be mentioned that the phenotype of the NTRC-deficient plants depends on light intensity and photoperiod showing more severe growth retardation when grown under short-day conditions, with a 16-h-dark/8-h-light photoperiod (see Supplemental Figure 1 online).

To characterize the function of NTRC further, we analyzed in more detail the effect of continuous darkness on wild-type, transgenic, and knockout plants. A treatment of 72 h in continuous darkness produced pronounced symptoms of leaf bleaching in the knockout plants (Figure 9A , D treatment). When plants were incubated under normal growth conditions following dark treatment (16 h light/8 h dark) for 3 d (D+LD), the wild-type and transgenic plants resumed growth, although showing more symptoms of leaf senescence than control (LD+LD) plants (Figure 9A). However, the NTRC-deficient plants presented accelerated leaf senescence (Figure 9A, D+LD treatment). The level of peroxides (Figure 9B) and lipid peroxidation (Figure 9C) in leaves, which were slightly higher in the ntrc mutant, were not significantly affected by a treatment of continuous darkness in any of the plant lines analyzed but were increased in the mutant plants after 3 d under standard growth conditions (Figures 9B and 9C, D+LD treatment). Therefore, stress symptoms in leaves were detected when mutant plants were again illuminated after the prolonged darkness treatment.


Figure 9
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  		Figure 9. Effect of Continuous Darkness and Reillumination on Wild-Type and NTRC-Deficient Arabidopsis Plants.

(A) Arabidopsis wild-type, ntrc knockout, and transgenic plants, as indicated, grown for 28 d in a growth chamber in a 16-h-light/8-h-dark cycle were incubated for 3 d under the same light/dark cycle (LD) or with continuous darkness (D). Both control (LD+LD) and dark-treated (D+LD) were then incubated for an additional 3 d under a 16-h-light/8-h-dark cycle. A representative plant is shown.

(B) and (C) Plants treated as indicated were collected, and the level of hydrogen peroxide (B) or lipid peroxidation as malonyldialdehyde (MDA) (C) in leaves was determined. Values represent means ± SD of at least three determinations per sample. FW, fresh weight.

 
To analyze the higher susceptibility of mutant plants, we studied the effect of darkness treatment and reillumination on the amount and stability of NTRC as well as the redox state of BAS1. The amount of NTRC was not significantly altered by the darkness treatment in the wild-type or transgenic plants, whereas, as expected, no NTRC was detected in the mutant plants (Figure 10A ). When BAS1 was analyzed under reducing conditions, a double band was identified (Figures 10B and 10C), probably corresponding to the expression of two highly similar genes (At5g06290 and At3g11630) encoding 2-Cys Prx in Arabidopsis. The amount of BAS1, detected under reducing conditions, was similar in wild-type, transgenic, and knockout plants and was not significantly affected by the darkness treatment in any of the Arabidopsis lines analyzed (Figures 10B and 10C). Protein gel blot analysis under nonreducing conditions detected most of BAS1 as a double band corresponding to the dimeric forms of the protein, the amount of dimeric form not being significantly affected by the darkness treatment in the wild-type (Figure 10B) and transgenic lines (Figure 10C). A progressive decrease of BAS1 dimers was observed in dark-treated mutant plants, BAS1 dimers being almost undetectable after reillumination (Figure 10B). Under nonreducing conditions, part of BAS1 was detected in monomeric form, which corresponds to fully reduced BAS1, in wild-type and transgenic lines (Figures 10B and 10C). By contrast, in NTRC-deficient plants, the fully reduced, monomeric form of BAS1 was almost undetectable (Figure 10B). These results show that NTRC deficiency causes unbalance of the redox state of BAS1 and lowers its capacity to form dimers in dark-treated plants.


Figure 10
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  		Figure 10. Effect of Continuous Darkness and Reillumination on the Level of NTRC and the Oxidation State of BAS1.

Protein extracts (40 µg of protein) from leaves of wild-type, NTRC-deficient, and transgenic Arabidopsis lines treated for the indicated time (h), as described in the legend to Figure 9, were subjected to SDS-PAGE (12% polyacryalmide), electrotransferred to nitrocellulose sheets, and probed with anti-NTRC antibodies (A). Protein extracts (20 µg of protein) from wild-type and NTRC-deficient plants (B) or from transgenic lines (C) were subjected to SDS-PAGE (15% polyacrylamide) under nonreducing and reducing conditions, as indicated, transferred to nitrocellulose sheets, and probed with anti-BAS1 antibodies. Molecular mass markers (kD) are on the left.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
METHODS
 REFERENCES
 
Oxygenic photosynthesis is the primary source of organic matter in the biosphere but is inevitably accompanied by the production of ROS, such as hydrogen peroxide. Given the dual role of hydrogen peroxide in signaling (Kwak et al., 2003Go; Overmyer et al., 2003Go; Vandenabeele et al., 2003Go; Foreman et al., 2004Go) and as a potentially harmful molecule (Mittler et al., 2004Go), the mechanism to control its production by the chloroplast is of great relevance. Previous studies identified a 2-Cys Prx, termed BAS1, as an important component for chloroplast peroxide detoxification (Konig et al., 2002Go). This Prx is efficiently reduced by Trx x (Collin et al., 2003Go) and interacts in vitro and in vivo with CDSP32, a plastid protein containing two Trx folds (Broin et al., 2002Go; Rey et al., 2005Go). Based on these results, the current model for chloroplast peroxide detoxification is based on the transfer of photosynthetic reducing power to ferredoxin, which in a reaction catalyzed by FTR would transfer reducing power to Trx x and/or CDSP32 and then to the 2-Cys Prx. Though no in vitro reconstitution of the FTR/(Trx x or CDSP32)/BAS1 system has been reported, the hypersensitivity of Arabidopsis FTR-deficient mutants to oxidative stress (Keryer et al., 2004Go) shows that this redox system plays an important role in chloroplast detoxification.

In this report, we show that rice NTRC, a novel type of NTR recently discovered in oxygenic photosynthetic organisms (Serrato et al., 2004Go), is able to transfer reducing power from NADPH to the 2-Cys Prx BAS1. This result suggests that NADPH, which is produced in the photosynthetic electron chain during the light period (Nelson and Ben-Shem, 2004Go), but also from the oxidative pentose phosphate pathway during the night (Neuhaus and Emes, 2000Go), may act as an alternative source of reducing power for chloroplast detoxification.

The catalytic mechanism of typical 2-Cys Prx involves two Cys residues: stated peroxidatic and resolving (Wood et al., 2003bGo). In the case of rice BAS1, the peroxidatic Cys residue Cys-61 is essential for activity and participates in the formation of the double disulfide, Cys-61–Cys-183, linking the oxidized, homodimeric form of BAS1 (Figure 3). AMS-labeling experiments clearly demonstrate that NTRC is able to reduce both disulfides using NADPH as source of reducing power (Figure 4A). Prokaryotes and eukaryotes seem to have evolved different mechanisms for 2-Cys Prx disulfide reduction. In eukaryotes, reduction is catalyzed by thioredoxins, which in turn are reduced by NTR (Schürmann and Jacquot, 2000Go). In prokaryotes, the more deeply characterized 2-Cys Prx, AhpC, is reduced by an NADH:peroxiredoxin reductase, AhpF (Poole et al., 2000aGo), a bimodular flavoenzyme composed of an NTR module and two Trx-like folds, with only one containing a redox-active disulfide (Poole et al., 2000bGo). Like AhpF, NTRC is a bimodular flavoenzyme and catalyzes the same reaction, that is, the transfer of electrons to the 2-Cys Prx BAS1. However, both enzymes show important differences: AhpF contains a double Trx-like fold at the N terminus, whereas NTRC contains a single Trx-like module at the C terminus, and, most importantly, AhpF is NADH dependent (Poole et al., 2000aGo; Wood et al., 2003bGo), whereas NTRC uses NADPH as source of reducing power (Figure 2).

The transfer of reducing power from NADPH to the 2-Cys Prx catalyzed by NTRC means that NTRC is a novel type of enzyme conjugating the activity of a NTR/Trx system in a single polypeptide. The lack of activity of any of the NTRC active sites mutants (C140S, C143S, C377S, and C380S) reveals that both modules, NTR and Trx-like, are essential for NTRC activity. Moreover, it is absolutely required that both activities form part of a single polypeptide as shown by the low activity obtained when mixtures of NTR and Trx-like truncated polypeptides were assayed compared with the full-length protein (Figure 2C).

Prxs are considered peroxidases of low enzymatic efficiency based on standard activity assays using NTR and Trx (Dietz, 2003Go). In agreement with this view, when the 2-Cys Prx BAS1 was assayed with wheat NTR and Trx hA or truncated NTR and Trx-like modules from NTRC (Figure 2C), the enzyme showed low catalytic efficiency. Remarkably, the activity of NTRC as a NTR-Trx system produces a drastic increase of the catalytic efficiency of the 2-Cys Prx, implying that the NTRC-BAS1 system is a high-efficiency redox system for peroxide reduction. The fact that NTRC genes are found only in cyanobacteria and plants (Serrato et al., 2004Go) suggests that this high-efficiency mechanism of detoxification is exclusive of oxygenic photosynthesis, probably because of the large amount of hydrogen peroxide produced by this process. In rice and Arabidopsis, we have shown that NTRC is localized to chloroplasts (Serrato et al., 2004Go). However, the rice gene undergoes several alternative splicing events (J.M. Pérez-Ruiz and F.J. Cejudo, unpublished data), which might produce versions of NTRC localized in other cell compartments not yet identified. So, the possibility of high-efficiency detoxification systems in cell compartments other than the chloroplast cannot be ruled out and deserves future investigation.

Based on the results reported here, we propose the existence of two pathways for transferring reducing power to the 2-Cys Prx involved in chloroplast detoxification (Figure 11 ). One of the pathways is formed by NTRC and uses NADPH as a source of reducing power; the other one is formed by FTR and Trx x and/or CDSP32 and uses reduced ferredoxin as a source of reducing power (Broin et al., 2002Go; Collin et al., 2003Go; Rey et al., 2005Go). Based on kinetic parameters, we conclude that NTRC is more efficient than the FTR-Trx pathway. Moreover, our results with the rice enzymes show that Trx x is much more efficient than CDSP32 to reduce BAS1 (Figure 5), in agreement with previous reports with the Arabidopsis enzymes (Broin et al., 2002Go; Collin et al., 2003Go). Given the low efficiency of CDSP32 as reductant of BAS1, compared with Trx x and NTRC, the function of this protein in peroxide detoxification is not clear. It should be taken into account that CDSP32 is induced during severe drought and oxidative stress (Rey et al., 1998Go; Broin et al., 2000Go). Therefore, CDSP32 interaction with BAS1 may have a function more relevant under stress conditions that is yet to be determined.


Figure 11
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  		Figure 11. Proposed Model for the Function of NTRC in Chloroplast Detoxification.

It is proposed that NTRC is an alternative system for transferring electrons from NADPH to the 2-Cys Prx BAS1 for peroxide detoxification. When light is on, ferredoxin reduced by the photosynthetic electron chain serves as source of reducing power by the FTR/Trx x (and CDSP32) pathway. In addition, NADPH can be used as source of reducing power through the NTRC pathway. During the night, when no ferredoxin is reduced by the photosynthetic chain, NTRC can transfer electrons to BAS1 using NADPH produced by the initial reactions of the oxidative pentose phosphate pathway, catalyzed by glucose-6P dehydrogenase (Glc6PDH) and 6P-gluconate dehydrogenase (6PGDH), respectively.

 
According to the proposed model (Figure 11), it would be expected that NTRC has an important function in plant growth and development as well as in response to environmental conditions. We have addressed the function of NTRC by analyzing an Arabidopsis knockout mutant. A previous description of this mutant showed retarded growth, light-green leaf color, and hypersensitivity to abiotic stress (Serrato et al., 2004Go). Here, we show that NTRC deficiency severely affects leaf photosynthetic cells (Figure 8), most of the effects (altered chloroplast structure, lower content of photosynthetic pigments, and lower rate of photosynthesis) being directly related with the photosynthetic machinery. These results provide an explanation for the retarded growth of the mutant and are in agreement with the effect of decreased levels of 2-Cys Prx, the target of NTRC, in transgenic Arabidopsis plants, which showed impaired photosynthesis and increased oxidative damage (Baier and Dietz, 1999Go; Baier et al., 2000Go).

The low rate of CO2 fixation of the NTRC-deficient plants (Figure 6) is an expected effect of the altered photosynthetic machinery of these plants. However, it is remarkable that this effect was light dependent, being negative at low light intensity and recovering to levels slightly lower than the wild type at higher light intensity. The strong effect of continuous darkness on the NTRC-deficient plants determined by the Fv/Fm ratio (Table 3) and leaf bleaching (Figure 9) and the severe retardation of growth when plants were grown under short-day conditions (see Supplemental Figure 1 online) lends further support to the proposal that NTRC plays an important role during the night.

The proposed model of NTRC function in chloroplast detoxification provides a possible explanation for the severe effect of darkness/reillumination treatment on the mutant plants. Under standard long-day growth conditions during the light period, both sources of reducing power for chloroplast detoxification, NADPH and reduced ferredoxin, are produced. So, NTRC deficiency might be partially compensated for by the light-dependent FTR/Trx pathway. However, during darkness, ferredoxin cannot be reduced by the photosynthetic electron chain, the only source of reduced ferredoxin being the reversal of the reaction catalyzed by ferredoxin NADPH oxidoreductase (Emes and Neuhaus, 1997Go); therefore, the FTR/Trx pathway for BAS1 reduction is probably less operative. The fact that this treatment is not lethal for the wild-type and transgenic plants suggests that NTRC is still able to transfer reducing power from NADPH produced in the oxidative pentose phosphate pathway to the 2-Cys Prx. The analysis of BAS1 under nonreducing conditions shows that most of the protein is detected in dimeric form (Figures 10B and 10C). However, in wild-type and transgenic plants, part of BAS1 appeared in monomeric form, indicating the amount of BAS1 fully reduced in vivo. The scarce detection of monomeric BAS1 in the mutant plants shows that NTRC deficiency causes unbalance of the redox state of BAS1, so that the fully reduced form of the enzyme is severely decreased. It should be noted that no monomeric BAS1 was detected in mutant plants incubated under control conditions (Figure 10B, LD), suggesting that the NTRC pathway is more relevant for BAS1 reduction than the FTR/Trx pathway even during the light period. These results are in agreement with the in vitro data showing that NTRC is able to reduce the two disulfides of BAS1 to produce the fully reduced monomeric form of the enzyme (Figure 4A) and the higher catalytic efficiency of NTRC compared with the Trx x or CDSP32 system (Figure 5). Though the amount of BAS1 was not altered, as shown by the analysis in reducing conditions, prolonged darkness treatment progressively reduced the capacity of BAS1 to form dimers in the NTRC-deficient mutant (Figure 10B). These results show that the redox status of BAS1 is severely altered in the mutant plants and suggest that BAS1 is very sensitive to darkness treatment in the absence of NTRC.

In summary, we describe a novel enzyme, NTRC, conjugating the activities of the NTR/Trx system in a single polypeptide and show that it is a high-efficiency reductant of the 2-Cys Prx BAS1. In vitro kinetic data and the unbalance of the redox state of BAS1 in the NTRC-deficient mutant suggest that the NTRC pathway plays a predominant role for BAS1 reduction in vivo. Whether the severe effect of NTRC deficiency on the photosynthetic machinery described here is due exclusively to the suppression of the NTRC-BAS1 detoxification pathway or to additional processes in which NTRC might be involved is not yet known. To address this point, the identification of additional targets of NTRC in the chloroplast is now underway.


   METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 METHODS
REFERENCES
 
Plant Material
Rice (Oryza sativa ssp japonica cv Nipponbare) grains were sterilized and germinated at 25°C on filter paper soaked with water. Arabidopsis thaliana wild-type (ecotype Columbia) and the T-DNA insertion mutant SALK_012208 were grown in moist vermiculite supplemented with Hoagland medium in culture chambers at 22°C during the light and 20°C during darkness. The light intensity was set at 140 µmol m–2 s–1. Long-day conditions consisted of 16 h light/8 h darkness, whereas short-day conditions consisted of 14 h darkness/10 h light. For production of Arabidopsis transgenic lines, full-length NTRC cDNA (DNA stock U-14278), obtained from the ABRC, was digested with KpnI and SalI and inserted into the binary vector pBIB-A7 (Becker, 1990Go). The construct was integrated into Arabidopsis wild-type and T-DNA insertion mutant SALK_012208 by Agrobacterium tumefaciens (C58pMP90)–mediated transformation using the floral dip method (Clough and Bent, 1998Go). Transformants were selected on Murashige and Skoog plates containing 50 µg mL–1 hygromycin. Transgenic lines with a single insertion, and homozygous for the transgene, were selected for further characterization.

Stress Treatments
Stress treatments were applied to Arabidopsis plants grown for 24 to 28 d. For salt treatment, plants were irrigated during 5 d with nutritive solution supplemented with 200 mM NaCl at days 1, 3, and 5. For drought treatment, plants were not irrigated during 5 d. Methyl viologen was sprayed on rosette leaves at days 1 and 3 of treatment. For cold treatment, plants were incubated during 5 d at 10°C constant temperature under the same day/night cycle, and continuous darkness was applied for periods of up to 3 d in the growth chamber at 22 to 20°C. When indicated, plants were incubated with standard growth conditions following darkness treatment (16 h light, 22°C/8 h darkness, 20°C).

Cloning of BAS1, CDSP32, and Trx x cDNAs from Rice: Expression and Purification of the Recombinant Proteins
The sequences of the rice homologs of BAS1, CDSP32, and Trx x genes were obtained from the Torrey Mesa Research Institute (Goff et al., 2002Go), and their putative gene structure was established using the GeneScan program. Rice BAS1 and Trx x cDNAs were cloned by RT-PCR from 1 µg total RNA of 5-d-old rice seedling shoots, which was reverse transcribed in the presence of oligo(dT) and 200 units of reverse transcriptase (Invitrogen). For rice BAS1 cloning, an aliquot (1 µL) was used as template in a PCR reaction with oligonucleotides 5'-AAGCTCCCAAACCCCTCT-3' and 5'-ATCAGACGAGCACACGG-3'. A 0.96-kb fragment was obtained, cloned in pGEMt vector (Promega), and sequenced in both strands. To express rice BAS1 in Escherichia coli, the coding sequence, excluding the putative signal peptide (53 residues), was amplified from the full-length cDNA with oligonucleotides 5'-GAGAGGATCCGCCAGCAGCTTCGTCGC-3' and 5'-GAGAAAGCTTCCCTACATCCATCACCAG-3', which added BamHI and HindIII sites (underlined) at the 5' and 3' ends, respectively. The cDNA encoding the putative mature form of rice Trx x, excluding 65 residues at the N terminus, was directly produced by PCR using as template 1 µL of the above-described reverse transcriptase reaction and oligonucleotides 5'-TTTTGGATCCGGCGCGGCGGTGAGGTT-3' and 5'-GGGGAAGCTTATTAAGCAACTGTTGAGG-3', which added BamHI and HindIII sites (underlined) at the 5' and 3' ends, respectively. Rice CDSP32 is apparently an intronless gene in rice, so the full-length DNA for CDSP32, except the putative signal peptide (47 residues), was directly amplified from rice genomic DNA using oligonucleotides 5'-GAGAGGATCCGAGCGGGTGGTGCAGGTG-3' and 5'-GAGAAAGCTTATATGTCTAGGTGACCTT-3', adding BamHI and HindIII sites (underlined) at the 5' and 3' ends, respectively. The truncated polypeptide containing exclusively the C-terminal Trx fold of CDSP32 was produced by the same approach using as template full-length CDSP32 cDNA and the pair of oligonucleotides 5'-AAAAGGATCCGGCGACCACCACTCCGC-3' and 5'-GAGAAAGCTTATATGTCTAGGTGACCTT-3', adding BamHI and HindIII sites. All cDNAs were sequenced in both strands. The PCR fragments were digested with BamHI and HindIII, subcloned into the pQE-30 expression vector (Qiagen), and introduced into E. coli XL1-Blue. Overexpressed proteins contained a His-tag at the N terminus and were purified by NTA affinity chromatography in prepacked Hi-Trap affinity columns (Amersham Biosciences).

Mutants of rice BAS1 (C61S) and NTRC (C140S, C143S, C377S, and C380S) were obtained by site-directed mutagenesis on the corresponding wild-type cDNAs cloned in the expression vector pQE30, according to the method previously described (Serrato et al., 2001Go). Pairs of oligonucleotides for each mutant were as follows: BAS1-C61S (5'-CTTCACCTTCGTCTCCCCGACCGAGATTAC-3') and (5'-GTAATCTCGGTCGGGGAGACGAAGGTGAAG-3'), NTRC-C140S (5'-AGGTATCAGTGCATCTGCAATATGTGATGG-3') and (5'-CCATCACATATTGCAGATGCACTGATACCT-3'), NTRC-C143S (5'-TGCATGTGCAATATCTGATGGAGCATCCCC-3') and (5'-GGGGATGCTCCATCAGATATTGCACATGCA-3'), NTRC-C377S (5'-ATACTTCTCCAACATCTGGTCCTTGCAGAA-3') and (5'-TTCTGCAAGGACCAGATGTTGGAGAAGTAT-3'), NTRC-C380S (5'-CAACATGTGGTCCTTCCAGAACTTAAAGC-3') and (5'-GCTTTAAGGTTCTGGAAGGACCACATGTTG-3'). The mutant nucleotide is underlined. Other recombinant proteins used in this work, NTR and Trx hA, were produced in E. coli as N-terminal His-tagged proteins (Serrato et al., 2001Go, 2002Go).

Relative RT-PCR, Production of Anti-BAS1 Polyclonal Antibodies, and Protein Gel Blot Analysis
RT-PCR analysis was performed as previously described (Sánchez and Cejudo, 2003Go) with gene-specific oligonucleotides for NTRC (5'-GAAGGGTATCAGATGGGC-3' and 5'-GGTTGGAGGAGGAGATAC-3'). Anti-BAS1 antibodies were raised by immunizing rabbits with purified His-tagged rice BAS1 at the Service for Animal Production (University of Seville, Spain). Protein gel blot analysis were performed as previously described (Serrato et al., 2002Go) using anti-NTRC as probe (Serrato et al., 2004Go). The anti-Rubisco polyclonal antibodies were purchased from Agrisera. Protein gel blot analysis of BAS1 was performed under reducing and nonreducing conditions. For nonreducing conditions, frozen leaves were ground with extraction buffer (100 mM Tris HCl, pH 7.9, 10% [v/v] glycerol, 1% [v/v] protease inhibitor cocktail [Sigma-Aldrich], 1 mM PMSF, 1 mM EDTA, and 100 mM Mg2Cl), and protein extracts were subjected to SDS-PAGE (15% polyacrylamide) in 16 x 20-cm gels overnight. For reducing conditions, the extraction buffer was supplemented with 10 mM DTT and 1.25% (v/v) ß-mercaptoethanol, and samples were boiled before loading.

Thioredoxin and Peroxiredoxin Activity Assays
Peroxiredoxin activity was determined as oxidation of NADPH following absorbance at 340 nm in a reaction mixture containing 100 mM phosphate buffer, pH 7.0, 2 mM EDTA, 0.25 mM NADPH, 0.5 mM hydrogen peroxide, and purified enzymes at the concentrations indicated in the figure legends. Alternatively, disappearance of hydrogen peroxide or t-BOOH was followed colorimetrically at 560 nm in the same reaction mixture without EDTA with the Peroxoquant reagent (Perbio Science). Thioredoxin activity was determined by the DTT-dependent reduction of insulin as described by Holmgren and Björnstedt (1995)Go.

Determination of Photosynthetic Pigments
Photosynthetic pigments (chlorophyll a and b and carotenoids) were extracted with 80% (v/v) ice-cold acetone from 0.5 g (fresh weight) of leaf fragments. Chlorophyll determinations were performed according to Lichtenthaler and Wellburn (1983)Go. Chloroplasts were isolated with the chloroplast isolation kit (Sigma-Aldrich) according to manufacturer's instructions.

Determination of Lipid Peroxidation
Measurements of lipid peroxidation in leaves were performed with the thiobarbituric acid (TBA) test, which determines MDA as an end product of lipid peroxidation (Heath and Parker, 1968Go). Briefly, 200 mg of leaves were homogenized in 2 mL of 0.1% (w/v) trichloroacetic acid (TCA) solution on ice. The homogenate was centrifuged at 12,000g for 15 min, and 0.4 mL of the supernatant was added to 0.8 mL of 0.5% (w/v) TBA in 20% TCA. The mixture was incubated in boiling water for 30 min, and the reaction was stopped by incubation in ice. Samples were then centrifuged at 10,000g for 5 min, and the absorbance of the supernatant was measured at 532 nm, subtracting the value for nonspecific absorption at 600 nm. The amount of MDA-TBA complex was calculated from the extinction coefficient 155 mm–1 cm–1.

Determination of Hydrogen Peroxide in Leaf Extracts
Hydrogen peroxide was determined according to Velikova et al. (2000)Go. Leaf tissues (200 mg) were homogenized in 2 mL of 0.1% (w/v) TCA solution on ice. The homogenate was centrifuged at 12,000g for 15 min, and 0.4 mL of the supernatant was added to 0.4 mL of 10 mM potassium phosphate buffer, pH 7.0, and 0.8 mL of 1 M KI. The absorbance of the supernatant was measured at 390 nm. The content of H2O2 was calculated by comparison with a standard calibration curve previously made using different concentrations of H2O2.

Determination of the Photosynthesis Rate, and PSII Photochemical Efficiency
In vitro photosynthetic rate of electron transport was determined as oxygen evolution at 25°C in a Hansatech oxygen electrode using osmotic shock-treated chloroplasts and 5 mM potassium ferricyanide as electron acceptor. Response curves of CO2 fixation to different light intensities (photosynthesis active radiation) were made with the portable photosynthesis system (LI-6400; LiCor), which allows environmental conditions inside the cuvette to be precisely controlled. Air temperature in the growth chamber was set at 25°C, and humidity was maintained at 50%. Six response curves were measured, each on a leaf from a different plant.

The change in chlorophyll fluorescence was measured at 22°C with the chlorophyll fluorometer PAM 2000 (Waltz). The maximal quantum yield of PSII (Fv/Fm) was determined from the following equation: Fv/Fm = (Fm – Fo)/Fm, where Fo is the initial minimal fluorescence on dark-adapted leaves for 15 min and Fm is maximal dark-adapted fluorescence.

In Vitro Determination of the Reduced State of BAS1
The redox state of BAS1 was determined by labeling Cys sulfhydryl groups with AMS (Hosoyoga-Matsuda et al., 2005Go). BAS1 incubated under different conditions, as indicated in the figure legends, was precipitated with 10% (v/v) TCA and collected by centrifugation. Precipitated protein was then washed with ice-cold acetone and dissolved in freshly prepared 50 mM Tris-HCl, pH 7.5, 1% (w/v) SDS, and 10 mM AMS. Labeled proteins were subjected to nonreducing SDS-PAGE (10% polyacrylamide).

Optical and Electron Microscopy Studies
For morphological analysis, small fragments of leaves from 29-d-old Arabidopsis wild-type, ntrc knockout mutant, and KO:P35SNTRC transgenic plants were fixed in 4% (v/v) glutaraldehyde prepared in 0.1 M cacodylate buffer, pH 7.2, for 3 h at 4°C and postfixed in 1% OsO4 for 2 h at 4°C. Samples were dehydrated in an acetone series and embedded in Epon (epoxy embedding medium). Toluidine blue–stained semithin sections (0.5-µm thick) were viewed in a Leitz (Aristoplan) light microscope. The images were captured by means of a digital camera (Leica DC-100). For electron microscopy, thin sections (60- to 80-nm thick) were stained with uranyl acetate and lead citrate and examined in a Philips CM-10 microscope. The images were captured by means of a digital camera (Megaview 3).

Accession Numbers
Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers AM039889 (BAS1), AM039890 (CDSP32), and AM183298 (Trx x).

Supplemental Data
The following material is available in the online version of this article.

Supplemental Figure 1. Effect of Short-Day Conditions on ntrc Mutant Growth.


   Acknowledgments
 
This work was supported by Grant BIO2004-02023 from the Ministerio de Educación y Ciencia (Spain) and Grant CVI-182 from Junta de Andalucía (Spain). J.M.P.-R. was supported by a predoctoral fellowship from the Ministerio de Educación y Ciencia, M.C.S. by a postdoctoral fellowship from Fundación Carolina (Spain), and K.K. by a predoctoral fellowship from the Consejo Superior de Investigaciones Científicas (Spain). We thank A. Díaz-Espejo (Instituto de Recursos Naturales y Agrobiología, Seville, Spain) for aid to determine the in vivo CO2 fixation curves and M.C. Díaz-Barradas (Universidad de Sevilla) for aid and facilities to determine the Fv/Fm. Technical assistance of R. Rodríguez is deeply appreciated.


   Footnotes
 
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Francisco Javier Cejudo (fjcejudo{at}us.es).

[W] Online version contains Web-only data. Back

www.plantcell.org/cgi/doi/10.1105/tpc.106.041541

Received January 30, 2006; Revision received June 13, 2006. accepted July 12, 2006.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 METHODS
 REFERENCES
 
Apel, K., and Hirt, H. (2004). Reactive oxygen species: Metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 55, 373–399.[CrossRef][Medline]

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

Baier, M., and Dietz, K.-J. (1997). The plant 2-Cys peroxiredoxin BAS1 is a nuclear-encoded chloroplast protein: Its expressional regulation, phylogenetic origin, and implications for its specific physiological function in plants. Plant J. 12, 179–190.[CrossRef][Web of Science][Medline]

Baier, M., and Dietz, K.J. (1999). Protective function of chloroplast 2-cysteine peroxiredoxin in photosynthesis: Evidence from transgenic Arabidopsis. Plant Physiol. 119, 1407–1414.[Abstract/Free Full Text]

Baier, M., Noctor, G., Foyer, C.H., and Dietz, K.J. (2000). Antisense suppression of 2-cysteine peroxiredoxin in Arabidopsis specifically enhances the activities and expression of enzymes associated with ascorbate metabolism but not glutathione metabolism. Plant Physiol. 124, 823–832.[Abstract/Free Full Text]

Becker, D. (1990). Binary vectors which allow the exchange of plant selectable markers and reporter genes. Nucleic Acids Res. 18, 203.[Free Full Text]

Broin, M., Cuiné, S., Aymery, F., and Rey, P. (2002). The plastidic 2-Cysteine peroxiredoxin is a target for a thioredoxin involved in the protection of the photosynthetic apparatus against oxidative damage. Plant Cell 14, 1417–1432.[Abstract/Free Full Text]

Broin, M., Cuiné, S., Peltier, G., and Rey, P. (2000). Involvement of CDSP32, a drought-induced thioredoxin, in the response to oxidative stress in potato plants. FEBS Lett. 467, 245–248.[CrossRef][Web of Science][Medline]

Clough, S.J., and Bent, A.F. (1998). Floral dip: A simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16, 735–743.[CrossRef][Web of Science][Medline]

Collin, V., Issakidis-Bourguet, E., Marchand, C., Hirasawa, M., Lancelin, J.-M., Knaff, D.B., and Miginiac-Maslow, M. (2003). The Arabidopsis plastidial thioredoxins. New functions and new insights into specificity. J. Biol. Chem. 278, 23747–23752.[Abstract/Free Full Text]

Dietz, K.-J. (2003). Plant peroxiredoxins. Annu. Rev. Plant Biol. 54, 93–107.[CrossRef][Medline]

Emes, M.J., and Neuhaus, H. (1997). Metabolism and transport in non-photosynthetic plastids. J. Exp. Bot. 48, 1996–2005.

Foreman, J., Demidchik, V., Bothwell, J.H.F., Mylona, P., Miedema, H., Torres, M.A., Linstead, P., Costa, S., Brownlee, C., Jones, J.D.G., Davies, J.M., and Dolan, L. (2004). Reactive oxygen species produced by NADPH oxidase regulate plant cell growth. Nature 422, 442–446.

Goff, S.A., et al. (2002). A draft sequence of the rice genome (Oryza sativa L. ssp. japonica). Science 296, 92–100.[Abstract/Free Full Text]

Heath, R.L., and Parker, L. (1968). Photoperoxidation in isolated chloroplasts. I. Kinetics and stoichiometry of fatty acid peroxidation. Arch. Biochem. Biophys. 125, 189–198.[CrossRef][Web of Science][Medline]

Holmgren, A., and Björnstedt, M. (1995). Thioredoxin and thioredoxin reductase. Methods Enzymol. 252, 199–208.[CrossRef][Web of Science][Medline]

Hosoyoga-Matsuda, N., Motohashi, K., Yoshimura, H., Nozaki, A., Inoue, K., Ohmori, M., and Hisabori, T. (2005). Anti-oxidative stress system in cyanobacteria. Significance of the type II peroxiredoxin and the role of 1-Cys peroxiredoxin in Synechocystis sp. strain PCC 6803. J. Biol. Chem. 280, 840–846.[Abstract/Free Full Text]

Keryer, E., Collin, V., Levergne, D., Lemaire, S., and Issakidis-Bourguet, E. (2004). Characterization of Arabidopsis mutants for the variable subunit of ferredoxin:thioredoxin reductase. Photosynth. Res. 79, 265–274.[CrossRef][Web of Science][Medline]

Konig, J., Baier, M., Horling, F., Kahmann, U., Harris, G., Schurmann, P., and Dietz, K.-J. (2002). The plant-specific function of 2-Cys peroxiredoxin-mediated detoxification of peroxides in the redox-hierarchy of photosynthetic electron flux. Proc. Natl. Acad. Sci. USA 99, 5738–5743.[Abstract/Free Full Text]

Konig, J., Lotter, K., Plessow, R., Brockhinke, A., Baier, M., and Dietz, K.-J. (2003). Reaction mechanism of plant 2-Cys peroxiredoxin. J. Biol. Chem. 278, 24409–24420.[Abstract/Free Full Text]

Kwak, J.M., Mori, I.C., Pei, Z.M., Leonhardt, N., Torres, M.A., Dangl, J.L., Bloom, R.E., Bodde, S., Jones, J.D., and Schroeder, J.I. (2003). NADPH oxidase AtrbohD and AtrbohF genes function in ROS-dependent ABA signalling in Arabidopsis. EMBO J. 22, 2623–2633.[CrossRef][Web of Science][Medline]

Laloi, C., Apel, K., and Danon, A. (2004). Reactive oxygen signalling: The latest news. Curr. Opin. Plant Biol. 7, 323–328.[CrossRef][Web of Science][Medline]

Ledford, H.K., and Niyogi, K.K. (2005). Singlet oxygen and photo-oxidative stress management in plants and algae. Plant Cell Environ. 28, 1037–1045.[CrossRef]

Lichtenthaler, H.K., and Wellburn, A.R. (1983). Determination of total carotenoids and chlorophylls a and b of leaf extracts in different solvents. Biochem. Soc. Trans. 603, 591–592.

Mittler, R. (2002). Oxidative stress, antioxidants and stress tolerance. Trends Plant Sci. 7, 405–410.[CrossRef][Web of Science][Medline]

Mittler, R., Vanderauwera, S., Gollery, M., and Van Breusegem, F. (2004). Reactive oxygen gene network of plants. Trends Plant Sci. 9, 490–498.[CrossRef][Web of Science][Medline]

Nelson, N., and Ben-Shem, A. (2004). The complex architecture of oxygenic photosynthesis. Nat. Rev. Mol. Cell Biol. 5, 1–12.[Free Full Text]

Neuhaus, H.E., and Emes, M.J. (2000). Nonphotosynthetic metabolism in plastids. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51, 111–140.[CrossRef][Web of Science][Medline]

Overmyer, K., Brosché, M., and Kangasjärvi, J. (2003). Reactive oxygen species and hormonal control of cell death. Trends Plant Sci. 8, 335–342.[CrossRef][Web of Science][Medline]

Pfannschmidt, T. (2003). Chloroplast redox signals: How photosynthesis controls its own genes. Trends Plant Sci. 8, 33–40.[CrossRef][Web of Science][Medline]

Poole, L.B., Godzik, A., Nayeem, A., and Schmitt, J.D. (2000b). AhpF can be dissected into two functional units: Tandem repeats of two thioredoxin-like folds in the N-terminus mediate electron transfer from the thioredoxin reductase-like C-terminus to AhpC. Biochemistry 39, 6602–6615.[CrossRef][Medline]

Poole, L.B., Reynolds, C.M., Wood, Z.A., Karplus, P.A., Ellis, H.R., and LiCalzi, M. (2000a). AhpF and other NADH:peroxiredoxin oxidoreductases, homologues of low Mr thioredoxin reductase. Eur. J. Biochem. 267, 6126–6133.[Web of Science][Medline]

Rey, P., Cuiné, S., Eymery, F., Garin, J., Court, M., Jacquot, J.-P., Rouhier, N., and Broin, M. (2005). Analysis of the proteins targeted by CDSP32, a plastidic thioredoxin participating in oxidative stress responses. Plant J. 41, 31–42.[CrossRef][Web of Science][Medline]

Rey, P., Pruvot, G., Becuwe, N., Eymery, F., Rumeau, D., and Peltire, G. (1998). A novel thioredoxin-like protein located in the chloroplast is induced by water deficit in Solanum tuberosum L. plants. Plant J. 13, 97–107.[CrossRef][Web of Science][Medline]

Rodriguez Milla, M.A., Maurer, A., Rodríguez Huete, A., and Gustafson, J.P. (2003). Glutathione peroxidase genes in Arabidopsis are ubiquitous and regulated by abiotic stresses through diverse signalling pathways. Plant J. 36, 602–615.[CrossRef][Web of Science][Medline]

Sánchez, R., and Cejudo, F.J. (2003). Identification and expression analysis of a gene encoding a bacterial-type phosphoenolpyruvate carboxylase from Arabidopsis and rice. Plant Physiol. 132, 949–957.[Abstract/Free Full Text]

Schürmann, P., and Jacquot, J.-P. (2000). Plant thioredoxin systems revisited. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51, 371–400.[CrossRef][Web of Science][Medline]

Serrato, A.J., Crespo, J.L., Florencio, F.J., and Cejudo, F.J. (2001). Characterization of two thioredoxins h with predominant localization in the nucleus of aleurone and scutellum cells of germinating wheat seeds. Plant Mol. Biol. 46, 361–371.[CrossRef][Web of Science][Medline]

Serrato, A.J., Pérez-Ruiz, J.M., and Cejudo, F.J. (2002). Cloning of thioredoxin h reductase and characterization of the thioredoxin reductase-thioredoxin h system from wheat. Biochem. J. 217, 392–399.

Serrato, A., Pérez-Ruiz, J.M., Spínola, M.C., and Cejudo, F.J. (2004). A novel NADPH thioredoxin reductase, localized in the chloroplast, which deficiency causes hypersensitivity to abiotic stress in Arabidopsis thaliana. J. Biol. Chem. 279, 43821–43827.[Abstract/Free Full Text]

Vandenabeele, S., Van der Kelen, K., Dat, J., Gadjev, I., Bonefaes, T., Morsa, S., Rttiers, P., Sloten, L., Van Montagu, M., Zabeau, M., Inzé, D., and Van Breusegem, F. (2003). A comprehensive analysis of hydrogen peroxide-induced gene expresión in tobacco. Proc. Natl. Acad. Sci. USA 100, 16113–16118.[Abstract/Free Full Text]

Vanderauwera, S., Zimmermann, P., Rombauts, S., Vandenabeele, S., Langebartels, C., Gruissem, W., Inzeé, D., and Van Breusegem, F. (2005). Genome-wide analysis of hydrogen peroxide-regulated gene expression in Arabidopsis reveals a high light-induced transcriptional cluster involved in anthocyanin biosynthesis. Plant Physiol. 139, 806–821.[Abstract/Free Full Text]

Velikova, V., Yordanov, I., and Edreva, A. (2000). Oxidative stress and some antioxidant system in acid rain treated bean plants: Protective role of exogenous polyammines. Plant Sci. 151, 59–66.

Wood, Z.A., Poole, L.B., and Karplus, P.A. (2003a). Peroxiredoxin evolution and the regulation of hydrogen peroxide signalling. Science 300, 650–653.[Abstract/Free Full Text]

Wood, Z.A., Schröder, E., Harris, J.R., and Poole, L.B. (2003b). Structure, mechanism and regulation of peroxiredoxins. Trends Biochem. Sci. 28, 32–40.[CrossRef][Web of Science][Medline]




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