1 Laboratory of Biophysics, School of Biological Sciences, Seoul National University, Seoul 151-742, Republic of Korea
2 Institute of Microbiology, Seoul National University, Seoul 151-742, Republic of Korea
3 Center for Cell Signalling Research, Division of Molecular Life Sciences, Ewha Women's University, Seoul 120-750, Korea
Correspondence
Sa-Ouk Kang
kangsaou{at}snu.ac.kr
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ABSTRACT |
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INTRODUCTION |
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A number of transcriptional regulators of photosystem gene expression have been identified (Gregor & Klug, 1999). Among them, PpsR was identified as a repressor of photopigment synthesis in R. sphaeroides (Penfold & Pemberton, 1994
; Gomelsky & Kaplan, 1995a
). It contains a helixturnhelix motif at the carboxy terminus and two PerArntSim (PAS) domains in its central region, which serve to bind redox cofactors or are involved in oligomerization (Penfold & Pemberton, 1994
; Gomelsky et al., 2000
; Taylor & Zhulin, 1999
). The PpsR homologue from Rhodobacter capsulatus, CrtJ (75 % homology), was reported to bind to the bchC promoter region in a redox-sensitive manner (Ponnampalam & Bauer, 1997
). Recently, it was reported that CrtJ could form an intramolecular disulphide bond in aerobically grown cells but not in anaerobically grown cells, and the binding activity to the bchC promoter increased fivefold in the presence of a disulphide bond (Masuda et al., 2002
). Subsequently, Masuda & Bauer (2002)
reported that PpsR also had the same redox property as CrtJ, except that the binding activity to the puc promoter increased 2·2-fold in the presence of a disulphide bond.
AppA was identified as a critical component required for the activation of photosystem gene expression in R. sphaeroides (Gomelsky & Kaplan, 1995b). A molecular-genetic analysis suggested that it would interact with PpsR to regulate photosystem gene expression (Gomelsky & Kaplan, 1997
). Recently, it was verified that the FAD cofactor of AppA was essential for the blue-light-dependent sensory transduction of photosystem gene expression (Braatsch et al., 2002
). AppA was reported to be able to break the disulphide bond in the oxidized PpsR as well as to form a stable AppAPpsR2 complex, but blue light inhibited formation of the AppAPpsR2 antirepressor complex (Masuda & Bauer, 2002
).
In the present study, we observed that the binding activity of PpsR to the puc promoter region was increased by the reduction of the disulphide bond in PpsR, and the mobility of the oxidized (disulphide-bonded) and the reduced PpsR could be distinguished on SDS-PAGE. Furthermore, thiol-specific chemical modification verified that the two cysteine residues in PpsR remain in their reduced form in vivo in spite of the presence of oxygen, which is at variance with the report that the disulphide bond of the oxidized PpsR can be formed in the presence of oxygen (Masuda & Bauer, 2002). The fact that the protein expression level and the binding activity of PpsR in APP11 decrease can be explained by the combination of some recent reports.
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METHODS |
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Purification of PpsR from E. coli.
Forward and reverse primers, 5'-AGAAGGAAGACATATGCTGGCCGG-3' and 5'-GTCAAAGCGACCATTGGATCCGCG-3', containing NdeI and BamHI sites (underlined), respectively, were used to amplify the ppsR coding sequence. The PCR product was cloned into pGEM-T easy vector and subsequently introduced into pET15b between the NdeI and BamHI sites to generate pET15b : : ppsR. The plasmid was transformed into E. coli strain BL21(DE3)pLysE. The His-tagged PpsR protein was overexpressed in LB by induction with 1 mM IPTG at 37 °C for 2 h. Cells were harvested, and the His-tagged PpsR was purified using a Ni2+-nitrilotriacetic acid (NTA) Sepharose column according to the guide proposed by the manufacturer (Novagen). The His-tag of PpsR was removed by human thrombin and the detached protein was repurified using a Ni2+-NTA Sepharose column and stored at -70 °C for further experiments. The N-terminal amino acid sequence of the purified protein was confirmed with a Procise Protein Sequencing System (Applied Biosystems). During the purification, proteins were analysed by SDS-PAGE according to Laemmli (1970).
Subcloning of ppsR and appA genes into pRK415.
Forward and reverse primers, 5'-CAGGCGAAGCTTCGCGACATCGCCACCT-3' (HindIII site underlined) and 5'-CGCAGGCCTGCAGCGCCGCCTCAAT-3', were used to amplify the putative promoter region of ppsR from genomic DNA of R. sphaeroides. The amplified DNA was inserted into the pGEM-T easy vector, and the product was designated pGPSPR. The insert fragment digested with AatII and PstI was ligated with the vector fragment of pPPS35 (1·6 kb fragment of ppsR) digested with AatII and PstI, and the product was designated pPSPR35. The last step was the ligation of the insert of pPSPR35 digested with HindIII and NsiI and the vector of pRK415 digested with HindIII and PstI. This plasmid was designated as pRK415 : : ppsR. p484Nco50 containing a
2·7 kb NcoI fragment of appA was digested with HindIII and SacI, and the insert fragment was ligated with pRK415 also digested with HindIII and SacI. This plasmid was designated as pRK415 : : appA.
Antibody preparation.
Purified PpsR was subjected to SDS-PAGE, and the protein band was cut from the gel and homogenized. This preparation was used to immunize mice. Mice were initially immunized subcutaneously with 5 µg of the protein. Two booster injections were given at 2-week intervals and bleeding was done 35 days after last injection. The blood was agglutinated for 1 h at 37 °C and centrifuged at 14 000 g for 15 min. The supernatant, mouse antiserum to PpsR, was obtained and stored at -70 °C.
Analyses of protein from R. sphaeroides.
Cells were harvested by centrifugation at 14 000 g for 10 min and washed with 20 mM Tris/HCl (pH 7·5), 0·1 mM EDTA, 0·1 mM DTT and 10 % (v/v) glycerol. After resuspension in the same buffer, cells were disrupted by sonication with a Microson XL-2000 (Heat System Ultrasonics). The supernatant obtained after centrifugation at 14 000 g for 20 min was used as crude extract. The protein concentration was determined by the Lowry method, with bovine serum albumin as a standard. The expression level of PpsR in crude extract was analysed by Western blotting, using mouse antiserum against PpsR as a primary antibody and alkaline-phosphatase-linked sheep anti-mouse IgG as a secondary antibody. For the colorimetric detection of alkaline phosphatase, p-nitro blue tetrazolium chloride and 5-bromo-4-chloro-indol-3-ol dihydrogen phosphate (ester) mono-p-toluidinium salt were used as substrates.
Gel mobility shift assay (GMSA).
To prepare the puc promoter region, PCR was carried out using the forward primer 5'-TTTTTGCAGCAGCGAGAGGCTG-3' and the reverse primer 5'-AAATCGACGGTTTGCGTGTAGG-3'. The amplified DNA fragment was cloned into pGEM-T easy vector and the construct was designated pGPUC. This plasmid was digested with EcoRI and the insert fragment was gel-purified. About 200 ng of purified fragment was end-labelled with Klenow fragment using digoxigenin (DIG)-dUTP (Roche) and dATP as substrates for 30 min at 37 °C. The labelled probes were clarified by ethanol precipitation and stored at -20 °C for further experiments.
GMSA was performed under the following conditions: 440 fmol probe DNA was incubated with either purified PpsR (0·2 ng2 µg) or crude extract (210 µg) at 30 °C in 15 µl binding buffer containing 50 mM Tris/HCl (pH 7·0), 1 mM EDTA, 150 mM NaCl, 15 µg calf thymus DNA or 1 µg poly(dI-dC) nonspecific competitor DNA, and 10 % (v/v) glycerol. After 30 min incubation, the reaction mixture was loaded onto a 4·5 % (w/v) non-denaturing Tris/acetate/EDTA-buffered polyacrylamide gel and electrophoresed at room temperature.
Measurement of in vivo redox states of PpsR.
The redox states of the two cysteines in PpsR in living cells were estimated by the method of thiol-specific chemical modification following acidic precipitation of protein. The cultured cells were treated with 10 % trichloroacetic acid to avoid thiol oxidation and then sonicated. The precipitate was washed several times with acetone. The pellet was dissolved in 100 mM Tris/HCl (pH 8·0), 1 mM EDTA and 0·1 % (w/v) SDS, and divided into three aliquots. The first one was treated with a final concentration of 70 mM iodoacetamide. The second one was first reduced by a final concentration of 10 mM DTT and then alkylated with iodoacetamide. The third one, with no modification, was used as a reference. These samples were loaded onto the SDS-PAGE gel without -mercaptoethanol in the sample buffer and analysed by Western blotting. In some experiments, N-ethylmaleimide (NEM) was replaced by iodoacetamide at pH 6·8.
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RESULTS |
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To observe the effect of redox reagents on purified PpsR, samples incubated under various redox conditions were subjected to SDS-PAGE using non-reducing SDS-sample buffer. As shown in Fig. 2(a), denatured PpsR existed in several oligomeric forms built of disulphide bonds: tetramer with intermolecular disulphide bonds, dimer with intermolecular disulphide bond, and monomer with or without intramolecular disulphide bond (lane 2). DTT treatment resulted in an increase of reduced monomer and disappearance of tetramer as shown in lane 3. However, the oligomeric state of PpsR was not changed by the reaction of ferricyanide, as shown in lane 4. Due to the diffusion of
-mercaptoethanol contained in the SDS-sample buffer, the additional reduced monomer of PpsR appeared at the edges of lanes 2 and 4 in Fig. 2(a)
. As shown in lanes 14 of Fig. 2(b)
, the reduced monomer of PpsR also increased with increasing DTT concentration. There are only two cysteines in PpsR, so these two amino acid residues must be involved in the intermolecular or intramolecular disulphide bond formation although they are separated by 173 amino acid residues in the primary amino acid sequence. These two residues were reported to be well conserved and to form a disulphide bond in CrtJ in the presence of oxygen(Masuda et al., 2002
). From these results, we conclude that the two cysteines in the purified PpsR are a mixture of thiol forms and residues involved in intramolecular or intermolecular disulphide bonds, and can bind to the puc promoter region when each thiol group in its two cysteines is in the reduced state.
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DISCUSSION |
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While this paper was being prepared, a report describing the redox property and DNA-binding activity of PpsR was published (Masuda & Bauer, 2002). That report presented two different results from ours: first, all of PpsR is oxidized to form an intramolecular disulphide bond under aerobic conditions; second, only PpsR containing intramolecular disulphide bond (oxidized monomer) was shown on SDS-PAGE. Unlike the trend of increasing DNA-binding activity by the reduction of the disulphide bond in our data, Masuda & Bauer (2002)
found that DNA-binding activity to the puc promoter region was decreased 2·2-fold by the reduction of the intramolecular disulphide bond in PpsR. However, we could not obtain a PpsR pool containing only intramolecular disulphide bond and its own binding activity to the target promoter could not be determined, so the comparison in DNA-binding activity might not be meaningful.
In E. coli, cytosolic redox potential was reported to lie between approximately -260 and -280 mV under aerobic and anaerobic conditions (Gilbert, 1990). So even if cells grow under aerobic conditions, the cytoplasm maintains a reduced state, and the cysteines in most proteins exist in their reduced state. When cysteines are oxidized to form a disulphide bond or their thiol groups are converted to sulfenic acid by oxidative stress or certain stimuli, these are reconverted into thiol groups by cellular reductants like reduced glutathione or thioredoxin or the glutaredoxin system (Åslund et al., 1999
; Claiborne et al., 1999
; Lee et al., 2002
). In the case of extracellular proteins containing several cysteine residues, disulphide bonds are made by the periplasmic Dsb system in prokaryotes, or the oxidative protein folding system residing in the endoplasmic reticulum in eukaryotes (Kadokura et al., 2003
; Fassio & Sitia, 2002
). Therefore, since PpsR is not an extracellular protein, it seems unlikely that all of the oxidized PpsR can withstand the tendency to reduction due to the presence of so many reductants under reduced cellular conditions, even though the cells are under aerobic conditions.
A protein containing an intramolecular disulphide bond usually migrates more quickly during SDS-PAGE than when it is fully reduced, because of a decrease in chain flexibility and hydrodynamic volume (Loferer et al., 1995; Kang et al., 1999
; Lee et al., 2002
). Furthermore, since the cysteines in PpsR are separated by 173 amino acid residues in the primary amino acid sequence, the structural difference between oxidized and reduced monomers may be increased. Therefore, as expected, the two forms were easily discriminated without any modifying reagent when subjected to SDS-PAGE.
It was known that the repressor activity of PpsR was active even under anaerobic conditions (Gomelsky & Kaplan, 1997). In apparent agreement with that result, we have shown that binding activity of PpsR to target promoters was maintained even under anaerobic conditions. Even in the results of Masuda & Bauer (2002)
, not all of the repressor activity of PpsR disappeared under anaerobic conditions, since the DNA-binding activity of the reduced PpsR decreased about 2·2-fold. However, in R. capsulatus, a reporter gene assay showed that transcriptional activity of photosystem genes (puc, crt and bch) was higher in DB469 (a CrtJ-null mutant) than in the wild-type under aerobic conditions but those in the two strains were almost the same under anaerobic-light conditions (Ponnampalam et al., 1995
). The DNA-binding activity of the reduced form is about 4·5-fold lower than that of the oxidized one. Unlike PpsR, CrtJ does not seem to work well under anaerobic conditions, which is consistent with the data of Masuda et al. (2002)
.
It would not make sense to maintain repressor activity under anaerobic conditions, since it has long been known that photosystem genes are expressed in large quantities under anaerobic conditions (Cohen-Bazire et al., 1957; Aagaard & Sistrom, 1972
). Since PpsR-binding sites overlap the -10 and -35 promoter regions of puc, crt and bch with RNA polymerase binding sites, it is difficult for these phenomena to happen simultaneously. On the other hand, under anaerobic conditions, several transcriptional activators are activated by the reduction of oxygen: e.g. PrrA, phosphorylation by PrrB; FnrL, reassembly of cofactor (Eraso & Kaplan, 1995
; Zeilstra-Ryalls & Kaplan, 1998
). Also, there was a report that RegA, a PrrA homologue in R. capsulatus, competed with CrtJ to bind to the bchC promoter region (Bowman et al., 1999
). Therefore, it is proposed that an anaerobic activator regulates photosystem gene expression under these conditions.
As mentioned above, photosystem gene expression was severely impaired in the AppA-null mutant APP11; this might be due to PpsR, since AppA was proposed to function as an antirepressor of PpsR (Gomelsky & Kaplan, 1995b; Gomelsky & Kaplan, 1997
). And recently anti-repressor (PpsR) action of AppA was verified by in vitro complex formation of the two proteins (Masuda & Bauer, 2002
). We also investigated the DNA-binding activity to target the promoter, and the protein level of PpsR, in an AppA null background to validate the relationship between the two proteins. Although the DNA-binding activity to the target promoter and the protein level of PpsR were expected to increase in an AppA-null background, both of them in fact decreased when compared with those in wild-type and complemented cells (Fig. 5a, b)
. It is difficult to understand this result from the viewpoint of the antirepressor activity of AppA. Nevertheless, we present a plausible model to rationalize our data and the others (Fig. 6
). APP11, impaired in the photosystem, contains fewer PpsR than wild-type and the difference was more obvious under anaerobic-dark conditions than under aerobic conditions. Anti-repressor (PpsR) action of AppA was verified by in vitro complex formation of the two proteins under anaerobic-dark conditions (Masuda & Bauer, 2002
). Recently PpaA was reported to activate photosystem gene expression and PpsR mediates the repression of ppaA gene expression (Gomelsky et al., 2003
). The ppaA and ppsR genes are located close to each other and have the same transcriptional direction. Therefore, if these two genes are an operon, AppA could indirectly activate transcription of the genes by binding PpsR under anaerobic-dark conditions (Fig. 6a
), but in the AppA-null mutant, the two genes are downregulated by PpsR and photosystem genes are also downregulated by decreased PpaA and derepressed PpsR even though the protein level of PpsR is low (Fig. 6b
). Under aerobic conditions, a similar explanation is possible and the decrease of the differences between wild-type and AppA-null mutant may be ascribed to the weak interaction of oxidized AppA and PpsR. In our experiment, the DNA-binding activity of PpsR increased in proportion to the protein level in the wild-type compared with the AppA-null mutant. It might be due to the dissociation of the complex (AppA and PpsR) by light or oxidized AppA in the experiment according to the model of Masuda & Bauer (2002)
. PpsR had a redox-sensitive property but remained in the reduced state in the cell, and its amount was reduced by disruption of AppA.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Åslund, F., Zheng, M., Beckwith, J. & Storz, G. (1999). Regulation of the OxyR transcription factor by hydrogen peroxide and the cellular thiol-disulfide status. Proc Natl Acad Sci U S A 96, 61616165.
Bowman, W. C., Du, S., Bauer, C. E. & Kranz, R. G. (1999). In vitro activation and repression of photosynthesis gene transcription in Rhodobacter capsulatus. Mol Microbiol 33, 429437.[CrossRef][Medline]
Braatsch, S., Gomelsky, M., Kuphal, S. & Klug, G. (2002). A single flavoprotein, AppA, integrates both redox and light signals in Rhodobacter sphaeroides. Mol Microbiol 45, 827836.[CrossRef][Medline]
Claiborne, A., Yeh, J. I., Mallett, T. C., Luba, J., Crane, E. J. 3rd, Charrier, V. & Parsonage, D. (1999). Protein-sulfenic acids: diverse roles for an unlikely player in enzyme catalysis and redox regulation. Biochemistry 38, 1540715416.[CrossRef][Medline]
Cohen-Bazire, G., Sistrom, W. R. & Stanier, R. Y. (1957). Kinetic studies of pigment synthesis by non-sulfur purple bacteria. J Cell Comp Physiol 49, 2568.
Davis, J., Donohue, T. J. & Kaplan, S. (1988). Construction, characterization, and complementation of a Puf- mutant of Rhodobacter sphaeroides. J Bacteriol 170, 320329.[Medline]
Eraso, J. M. & Kaplan, S. (1995). Oxygen-insensitive synthesis of the photosynthetic membranes of Rhodobacter sphaeroides: a mutant histidine kinase. J Bacteriol 177, 26952706.[Abstract]
Fassio, A. & Sitia, R. (2002). Formation, isomerisation and reduction of disulphide bonds during protein quality control in the endoplasmic reticulum. Histochem Cell Biol 117, 151157.[CrossRef][Medline]
Gilbert, H. F. (1990). Molecular and cellular aspects of thiol-disulfide exchange. Adv Enzymol Relat Areas Mol Biol 63, 69172.[Medline]
Gomelsky, M. & Kaplan, S. (1995a). Genetic evidence that PpsR from Rhodobacter sphaeroides 2.4.1 functions as a repressor of puc and bchF expression. J Bacteriol 177, 16341637.[Abstract]
Gomelsky, M. & Kaplan, S. (1995b). appA, a novel gene encoding a trans-acting factor involved in the regulation of photosynthesis gene expression in Rhodobacter sphaeroides 2.4.1. J Bacteriol 177, 46094618.[Abstract]
Gomelsky, M. & Kaplan, S. (1997). Molecular genetic analysis suggesting interaction between AppA and PpsR in regulation of photosynthesis gene expression in Rhodobacter sphaeroides 2.4.1. J Bacteriol 179, 128134.[Abstract]
Gomelsky, M., Horne, I. M., Lee, H.-J., Pemberton, J. M., McEwan, A. G. & Kaplan, S. (2000). Domain structure, oligomeric state, and mutational analysis of PpsR, the Rhodobacter sphaeroides repressor of photosystem gene expression. J Bacteriol 182, 22532261.
Gomelsky, L., Sram, J., Moskvin, O. V., Horne, I. M., Dodd, H. N., Pemberton, J. M., McEwan, A. G., Kaplan, S. & Gomelsky, M. (2003). Identification and in vivo characterization of PpaA, a regulator of photosystem formation in Rhodobacter sphaeroides. Microbiology 149, 377388.
Gregor, J. & Klug, G. (1999). Regulation of bacterial photosynthesis genes by oxygen and light. FEMS Microbiol Lett 179, 19.[CrossRef][Medline]
Hanahan, D. (1983). Studies on transformation of Escherichia coli with plasmids. J Mol Biol 166, 557580.[Medline]
Kadokura, H., Katzen, F. & Beckwith, J. (2003). Protein disulfide bond formation in prokaryotes. Annu Rev Biochem 72, 111135.[CrossRef]
Kang, J. G., Paget, M. S. B., Seok, Y. J., Hahn, M. Y., Bae, J. B., Hahn, J. S., Kleanthous, C., Buttner, M. J. & Roe, J. H. (1999). RsrA, an anti-sigma factor regulated by redox change. EMBO J 18, 42924298.
Keen, N. T., Tamaki, S., Kobaysahi, D. & Trollinger, D. (1988). Improved broad-host-range plasmids for DNA cloning in Gram-negative bacteria. Gene 70, 191197.[CrossRef][Medline]
Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680685.[Medline]
Lee, S. R., Yang, K. S., Kwon, J., Lee, C., Jeong, W. & Rhee, S. G. (2002). Reversible inactivation of the tumor suppressor PTEN by H2O2. J Biol Chem 277, 2033620342.
Loferer, H., Wunderlich, M., Hennecke, H. & Glockshuber, R. (1995). A bacterial thioredoxin-like protein that is exposed to the periplasm has redox properties comparable with those of cytoplasmic thioredoxins. J Biol Chem 270, 2617826183.
Masuda, S. & Bauer, C. E. (2002). AppA is a blue light photoreceptor that antirepresses photosynthesis gene expression in Rhodobacter sphaeroides. Cell 110, 613623.[Medline]
Masuda, S., Dong, C., Swem, D., Setterdahl, A. T., Knaff, D. B. & Bauer, C. E. (2002). Repression of photosynthesis gene expression by formation of a disulfide bond in CrtJ. Proc Natl Acad Sci U S A 99, 70787083.
Penfold, R. J. & Pemberton, J. M. (1994). Sequencing, chromosomal inactivation, and functional expression in Escherichia coli of ppsR, a gene which represses carotenoid and bacteriochlorophyll synthesis in Rhodobacter sphaeroides. J Bacteriol 176, 28692876.[Abstract]
Ponnampalam, S. N. & Bauer, C. E. (1997). DNA binding characteristics of CrtJ. J Biol Chem 272, 1839118396.
Ponnampalam, S. N., Buggy, J. J. & Bauer, C. E. (1995). Characterization of an aerobic repressor that coordinately regulates bacteriochlorophyll, carotenoid, and light harvesting-II expression in Rhodobacter capsulatus. J Bacteriol 177, 29902997.[Abstract]
Sambrook, J., Fritsch, E. F. & Maniatis, T. (1989). Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.
Simon, R., Priefer, U. & Puhler, A. (1983). A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram negative bacteria. Bio/Technology 1, 784791.
Taylor, B. L. & Zhulin, I. B. (1999). PAS domains: internal sensors of oxygen, redox potential, and light. Microbiol Mol Biol Rev 63, 479506.
Zeilstra-Ryalls, J. H. & Kaplan, S. (1998). Role of the fnrL gene in photosystem gene expression and photosynthetic growth of Rhodobacter sphaeroides 2.4.1. J Bacteriol 180, 14961503.
Received 19 September 2003;
revised 21 November 2003;
accepted 24 November 2003.
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