![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
Vol. 18, Issue 6, 2288-2295, June 2007
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Departament de Ciències Experimentals i de la Salut, Universitat Pompeu Fabra, E-08003 Barcelona, Spain
Submitted November 27, 2006;
Revised March 16, 2007;
Accepted March 26, 2007
Monitoring Editor: Thomas Fox
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
), with six mammalian isoforms distributed in different organelles (Rhee et al., 2005b). Three types of structurally different Prxs have been described: 1-cysteine (Cys), 2-Cys, and atypical 2-Cys (for review, see Wood et al., 2003b). All three types of Prxs are dimers in solution, and they all share the same catalytic mechanism, in which the peroxidatic Cys, located close to the N-terminal domain, is oxidized to a sulfenic acid by either H2O2 or alkyl hydroperoxides. In both types of 2-Cys Prxs, the sulfenic acid then reacts with the C-terminal (or resolving) Cys of the other subunit to form an intermolecular disulfide (classical 2-Cys Prxs), or with the C-terminal Cys of the same monomer to form an intramolecular disulfide (atypical 2-Cys Prxs). In both cases, the disulfides are specifically reduced by the thioredoxin and thioredoxin reductase system, with the exception of some prokaryotic Prxs, such as the Escherichia coli AhpC, which uses another protein, AhpF, for the regeneration of the reduced Prx. In the 1-Cys Prxs, the sulfenic acid is directly reduced to thiol, because there is no nearby Cys available to form a disulfide bond; the source of the reducing equivalents for regenerating this thiol is not known, although glutathione (GSH) has been proposed to serve as the electron donor in this reaction (Kang et al., 1998b).
Prx activity can be regulated by phosphorylation (Chang et al., 2002), and possibly by changes in oligomerization states (Wood et al., 2002, 2003b). Furthermore, some Prxs suffer an oxidation of one of their Cys residues, the peroxidatic Cys, to sulfinic acid upon exposure to high H2O2 doses, which temporarily inactivate these enzymes (Yang et al., 2002; Woo et al., 2003b; Wood et al., 2003a). This hyperoxidation of 2-Cys Prx enzymes is reversible in cells (Woo et al., 2003a,b). The enzymes named sulfiredoxin, found in Saccharomyces cerevisiae (Biteau et al., 2003), Schizosaccharomyces pombe (Bozonet et al., 2005; Vivancos et al., 2005), and mammalian cells (Chang et al., 2004), were isolated as the ATP-dependent reductases for Cys-sulfinic acid in Prxs. Another family of enzymes named sestrins has been shown to have a similar role in mammalian cells (Budanov et al., 2004).
The main role of Prxs has classically been considered to be related to their peroxidase activity. Their affinity for H2O2, with published Km values 
20 µM, is higher than that of other peroxide scavengers such as catalase or GSH peroxidase (Chae et al., 1999; Rhee et al., 2001), which suggests that Prxs could be efficient at removing low concentrations of H2O2. However, their temporary redox inactivation seems to suggest that Prx enzymes might have a role in signal transduction, with their inactivation shunt causing a temporary or local increase in H2O2 concentration that could then trigger antioxidant cascades (Rhee et al., 2005a).
The first evidence for the participation of a Prx in signal transduction pathways came from studies of fission yeast antistress responses. In S. pombe, the Pap1 and Sty1 pathways constitute the key protective responses to oxidative stress. It has been recently reported that the only 2-Cys Prx in fission yeast, Tpx1, is the H2O2 sensor of the Pap1 pathway, its presence being essential for activation of the transcription factor and its specific gene response. The Pap1 pathway is more sensitive to H2O2 than the mitogen-activated protein kinase Sty1 pathway: maximal activation of Sty1 requires a H2O2 concentration at least fivefold higher than that described to fully activate Pap1 (Quinn et al., 2002; Vivancos et al., 2004). As described above, 2-Cys Prxs such as Tpx1 undergo substrate-mediated inactivation, so high concentrations of H2O2 will temporarily inactivate Tpx1, postponing Pap1 activation; meanwhile, the Sty1 pathway will be fully functional under these conditions. Activated Sty1 is then required for reactivation of Tpx1, because synthesis of the Tpx1-reductase Srx1 during severe H2O2 stress is dependent on Sty1 (Bozonet et al., 2005; Vivancos et al., 2005).
Thus, Prxs have aroused a great deal of interest during recent years, and their emerging roles other than H2O2 scavenging—as redox signal transducers and molecular chaperones (Jang et al., 2004; Chuang et al., 2006)—is igniting a very interesting field of study. Indeed, Tpx1 was initially identified on the basis of its role in signal transduction to Pap1 in fission yeast. However, evidence from deletion mutants points to additional actions by this Prx. A strain lacking the tpx1 gene is not viable (Vivancos et al., 2005), whereas deletion of either pap1 or sty1 genes does not compromise cell viability, and cells lacking any of the oxidative stress signaling components only display phenotypes in the presence of extracellular stress (for review, see Ikner and Shiozaki, 2005). In this work, we demonstrate that Tpx1 is an essential H2O2 scavenger in S. pombe, with an extremely high affinity for its substrate and that this peroxidase activity is required only for aerobic growth. A mutant Tpx1 protein lacking the resolving Cys169 (Tpx1.C169S) is also able to support aerobic growth of cells expressing only this isoform, and small thiols such as dithiothreitol and GSH can provide the reducing equivalents to recycle oxidized Tpx1.C169S in vitro. This is the first eukaryotic 2-Cys Prx reported to scavenge H2O2 in vivo in the absence of its resolving Cys, in a manner similar to 1-Cys Prx enzymes.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
), and EA38 (h– leu1 srx1::kanMX6) (Vivancos et al., 2005). To construct S. pombe strains with specific loci deleted, we transformed some of the above-mentioned strains with linear fragments containing ORF::kanMX6, obtained by polymerase chain reaction (PCR) amplification with open reading frame (ORF)-specific primers and plasmid pFA6a-kanMX6 as a template. Following that strategy, we obtained strains AV25 (h– pap1::kanMX6), AV18 (h– sty1::kanMX6) and AM42 (h– leu1 his2 ura4 pap1::ura4 sty1::kanMX6). To isolate cells deleted in sod1, we transformed the wild-type strain PN513 with a PCR-amplified sod1::lacZ:kanMX6 fragment, obtained by PCR amplification with ORF-specific primers and plasmid AY017 as a template (plasmid AY017 is a pREP3x [Maundrell, 1993] derivative containing a lacZ:kanMX6 insert). Transformed cells were grown under anaerobic conditions and selected by their ability to grow in Kanamycin-containing plates. The strain generated was named MS1 (h– leu1 ura4 sod1::lacZ:kanMX6). To isolate haploid cells deleted in tpx1, we transformed the diploid strain AV29 with a PCR-amplified tpx1::kanMX6 fragment, yielding strain AV36dip (h+/h– leu1/leu1 ura4/ura4 ade6-M210/ade6-M276 tpx1::kanMX6/tpx1). Haploid cells were isolated by standard genetic techniques (Moreno et al., 1991), spread in Kanamycin-containing plates and grown anaerobically (see below). This yielded strain AV42 (h+ leu1 ura4 ade6-M210 tpx1::kanMX6). Plasmids p145 (ptpx1’::tpx1), p145.C48S (ptpx1’::tpx1.C48S), and p145.C169S (ptpx1’::tpx1.C169S), containing the promoter of tpx1 fused to the wild-type or mutant ORF of tpx1 (Vivancos et al., 2005), were linearized and introduced at the leu1 locus of strain AV42, yielding strains AV49, AV49.C48S, and AV49.C169S. To construct the strain AV49.C169S disrupted in cys1a, we crossed it with strain cys1a
(h– leu1-32 ura4-C190T cys1a::ura4) (Fujita and Takegawa, 2004). An equal volume of each strain was mixed under starvation conditions for nitrogen source. After 2 d at 25°C, the mating mixture was suspended in 0.7 ml of sterile distilled water with 0.3% 
-glucuronidase (Sigma-Aldrich, St. Louis, MO) and incubated at 25°C overnight. The resulting spores were washed and resuspended in water, spotted on YE5S plates supplemented with Cys and G-418 (Geneticin; Invitrogen, Paisley, United Kingdom) and streaked for single colonies. Kanamycin-resistant cells with auxotrophy for Cys were isolated and confirmed by PCR and immunodetection of trichloroacetic acid (TCA)-based protein extracts. The resulting strain was named MJ1.
Anaerobic Growth Conditions and H2O2 Sensitivity Assay
Anaerobic liquid cultures were grown in flasks filled to the top with medium. Cells were grown at 30°C without shaking. To grow cells in solid media in an anaerobic environment, we incubated the plates at 30°C in a tightly sealed plastic bag containing a water-activated Anaerocult A (Merck, Darmstadt, Germany) sachet. To analyze sensitivity to H2O2 on plates, S. pombe strains were grown anaerobically in liquid rich media to an OD600 of 0.5. Cells were then diluted in water, and the indicated number of cells in 4 µl was spotted onto rich media. The spots were allowed to dry, and the plates were incubated at 30°C in aerobic and anaerobic conditions, in the presence or absence of H2O2, for 3–4 d.
Plasmids
To express S. pombe proteins in Escherichia coli, the ORFs coding for Tpx1, Tpx1.C48S, and Tpx1.C169S were digested from plasmids p123.41x, p123.41x.C48S, and p123.41x.C169S with BamHI and SmaI, and they were subcloned into a modified glutathione S-transferase (GST)-tagging fusion vector pGEX-2T-TEV that encoded a TEV protease cleavage site between the tag and the cloned ORF, digested with the same restriction enzymes. The resulting plasmids were called p210, p210.C48S, and p210.C169S. The trr1 ORF was amplified from an S. pombe cDNA library by using specific primers containing BamHI and EcoRI restriction sites, and then it was cloned into the pGEX-2T-TEV expression vector digested with the same restriction enzymes, yielding plasmid p205. The trx1 ORF was amplified from genomic DNA using specific primers for the gene and containing BamHI and SmaI restriction sites. The fragment was cloned into pGEX-2T-TEV cleaved with BamHI and SmaI, yielding plasmid p206. Once PCR amplified, all ORFs in these expression vectors were sequenced.
Preparation of S. pombe Extracts to Measure Protein Carbonylation
Protein carbonylation was detected after in vitro derivatization of oxidized proteins with 2,4-dinitrophenylhydrazine (DNPH) (Levine et al., 1994; Cabiscol and Levine, 1995). Cells (50 ml) were grown aerobically over 12 h in rich media, harvested at an OD600 of 0.5, and resuspended in carbonylation buffer (50 mM Tris-HCl, pH 7.5, 2 mM EDTA, 0.05% NP-40, 2 mM phenylmethylsulfonyl fluoride [PMSF], 5 mM benzamidine, 0.16 mg/ml aprotinin) and lysed by vortexing after the addition of glass beads. The protein extracts were then centrifuged to eliminate cell debris. Protein concentration was determined using the Bradford protein assay (Bio-Rad, Hercules, CA). Protein concentration was adjusted to a 4–10 mg/ml range with carbonylation buffer, and a concentrated 12% solution of SDS was added to 10 µg of total protein, to reach a final SDS concentration of 6%, and incubated for 3 min at 100°C. One volume of 10 mM DNPH in 10% trifluoroacetic acid was added to 1 volume of the sample at 25°C. The reaction was run for 10 min and stopped by the addition of 1 volume of 2 M Trizma base, 10% glycerol, and 15% 2-mercaptoethanol. Samples were separated electrophoretically in 10% SDS-polyacrylamide gel electrophoresis (PAGE) gels and carbonylated proteins were immunodetected using a polyclonal anti-2,4-dinitrophenyl (DNP) antibody (Sigma-Aldrich). As a loading control, Sty1 was detected using polyclonal anti-Sty1 antiserum raised against an E. coli fusion protein of GST-Sty1, following standard rabbit immunization procedures. The anti-DNP and anti-Sty1 immunoblots were scanned, and quantification of carbonylated proteins was performed using the ImageQuant 5.2 program (GE healthcare, Little Chalfont, Buckinghamshire, United Kingdom).
Purification of Recombinant Tpx1, Thioredoxin (Trx1), and Thioredoxin Reductase (Trr1) Proteins for In Vitro Assays
Bacteria strain FB810 (Benson et al., 1994) transformed with the pGEX-2T-TEV derivatives were inoculated into Luria broth with 100 µg/ml ampicillin and incubated at 37°C for 16 h with vigorous shaking. The overnight cultures were diluted 25-fold into 400 ml of fresh medium and incubated at 37°C until the culture reached an optical density of 0.8 at 600 nm. Isopropyl--thio-D-galactoside (IPTG) was then added to a final concentration of 0.5 mM and shaking continued at 25°C for 4 h. The cells were then harvested, and pellets were resuspended in 10 ml of STET extraction buffer (50 mM Tris-HCl, pH 8.0, 150 mM sodium chloride, 1 mM EDTA, pH 8.0, 1% Triton X-100, and 2 mM PMSF) and broken by sonication. Debris and unbroken cells were removed by centrifugation. Supernatants containing our GST-tagged fusion proteins were then incubated with GSH-Sepharose 4B beads (GE Healthcare) for 1 h at 4°C. The beads were then washed three times with NET-N (20 mM Tris-HCl, pH 8.0, 1 mM EDTA, 100 mM sodium chloride, 0.5% NP-40, and 2 mM PMSF) and once with TEV cleavage buffer (10 mM Tris-HCl, pH 8.0, 150 mM sodium chloride, 0.1% NP-40, and 0.5 mM EDTA). We could then release the GST-tagged protein with GSH-containing elution buffer (100 mM Tris HCl, pH 8.0, 120 mM NaCl, and 20 mM GSH). Alternatively, we also released the S. pombe proteins without the GST tag from the Sepharose beads by incubating a 200-µl bed volume of beads with 10 µg of TEV protease (Invitrogen, Carlsbad, CA) overnight at 4°C. Size differences among GST-containing or untagged protein samples Tpx1, Trr1, and Trx1 were observed by electrophoretic separation on 15% denaturing polyacrylamide gels and Coomassie staining.
Peroxidase Activity Assays
NADPH oxidation was monitored as a decrease in optical density at 340 nm by using a Ultrospec 3100 pro UV/Visible spectrophotometer (GE Healthcare) in a 500-µl reaction mixture containing 50 mM HEPES-NaOH, pH 7.0, 0.25 mM NADPH, 6 µg of Trr1, 20 µg of Trx1, 1 µg of Tpx1, and H2O2. Reactions were started by the addition of the indicated concentrations of H2O2. As peroxidase activity of Tpx1 undergoes substrate-mediated inactivation, an initial linear portion of absorbance change (10 s) was used for the calculation of peroxidase activity, as described previously (Koo et al., 2002).
To determine the H2O2 detoxification driven by Tpx1, Tpx1.C48S, and Tpx1.C169S in the presence of small thiols, we incubated 5–20 µg of enzyme with the Trx system (consisting of the same reaction mixture as described in the previous paragraph), 10 or 100 mM dithiothreitol (DTT) (in 50 mM HEPES-NaOH, pH 7.0), or 50 µM GSH (in 100 mM Tris-HCl, pH 8.8, to avoid acidification of the assay mixture). We started the reaction by the addition of 50 µM H2O2, and 150-µl aliquots were taken at different time points and stopped by the addition of 20 µl of 100% TCA. Samples were then centrifuged to eliminate proteins, and 100 µl of the supernatant were used to determine the remaining H2O2 concentration. This was achieved by addition of 27 µl of 10 mM ferrous ammonium sulfate and 13.5 µl of 2.5 M potassium thiocyanate. The red-colored ferrithiocyanate complex, which occurs as a result of the oxidation of Fe(II) by H2O2, was quantified by measuring the OD at 480 nm and was compared with H2O2 standards.
Preparation of S. pombe TCA Extracts and Immunoblot Analysis
For in vivo redox state analysis of Tpx1, S. pombe cultures (5 ml) at an OD600 of 0.5 were pelleted just after the addition of 100% TCA, to a final concentration of 10%, and washed in 20% TCA. The pellets were lysed by vortexing, after the addition of glass beads and 12.5% TCA. Cell lysates were pelleted, washed in acetone, and dried. Alkylation of free thiols was performed by resuspension of the pellets in 50 µl of a solution containing 75 mM iodoacetamide, 1% SDS, 100 mM Tris-HCl, pH 8.0, and 1 mM EDTA, and incubation at 25°C for 15 min. Alkylated samples were diluted fivefold and electrophoretically separated by reducing SDS-PAGE, as indicated, and proteins were immunodetected using polyclonal anti-Tpx1 antiserum raised against an E. coli fusion protein of GST-Tpx1, following standard rabbit immunization procedures. For the detection of sulfinylated Tpx1, the undiluted alkylated samples were separated by nonreducing SDS-PAGE and immunodetected with anti-peroxiredoxin-SO3 antibody (LabFrontier, Seoul, South Korea).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). To test whether the essential role of Tpx1 in fission yeast was due to its peroxidase activity, we attempted to isolate haploid cells carrying the tpx1 deletion from diploid cells, in which one of the tpx1 loci is substituted by the kanamycin cassette, by growing them under anaerobic conditions inside sealed Anaerocult sachets. The chemical mixture inside the sachets contains components that chemically bind oxygen rapidly and effectively, creating an oxygen-free atmosphere. Positive clones were isolated, grown anaerobically in liquid, and doubly tested for lack of Tpx1 and their inability to oxidize Pap1 in response to H2O2 (data not shown). Deletion of tpx1 increased the sensitivity of cells to aerobic growth, but cells grew as wild type in the absence of oxygen (Figure 1A). On the contrary, cells lacking Pap1, Sty1, or both grew as efficiently under aerobic or anaerobic conditions (Figure 1A).
|
tpx1 cells. To test whether these cells were suffering from high levels of intrinsic oxidative stress when grown in the presence of oxygen, we measured the levels of protein carbonylation. Redox-cycling cations such as reduced iron or copper can bind to proteins, and, with the aid of further H2O2 attack, they can transform side-chain amino groups of several basic amino acids, such as lysine, arginine, proline, or histidine, into carbonyls. These carbonyl groups can be derived in vitro with DNPH, a modification that can be immunodetected with anti-DNP antibodies (see Materials and Methods). Under aerobic growth conditions, protein carbonylation in tpx1 cells was significantly higher than in wild-type cells (Figure 1B). Indeed, extracts of cells lacking Tpx1 contained as many protein carbonyls as those of a strain lacking the sod1 gene, which encodes the cytosolic superoxide dismutase; this strain is also only viable when grown under anaerobic conditions (data not shown).
Tpx1 Is a Peroxidase with Very High Affinity for H2O2
In most cell types, Prxs are not essential for aerobic growth. However, the lack of Tpx1 seems to drive cells grown in the presence of oxygen toward a basal oxidative stress, resulting in growth arrest. We decided to test whether Tpx1 scavenges the majority of endogenous H2O2 in S. pombe by assaying its capacity to decompose low concentrations of peroxides in vitro. The enzymatic activity of Prxs can be measured by coupling them to Trx1 and Trr1 in the presence of NADPH, following the reaction illustrated in Figure 2A. Traditionally, to assay the activity of eukaryotic Prxs, the three protein components of this reaction were purified from cell extracts by a series of multiple purification steps. We have developed an alternative strategy, in which Tpx1, Trx1, and Trr1 of S. pombe are purified from E. coli cells. We constructed expression vectors harboring full-length tpx1, trx1, and trr1 genes fused to the GST-coding gene. The fusion proteins (GST-Tpx1, GST-Trx1, and GST-Trr1) were purified almost to homogeneity from E. coli extracts by affinity binding to GSH-beads. The GST tag interfered with the activity of some of the purified proteins, so we cleaved it with TEV protease, yielding highly purified recombinant Tpx1, Trx1, and Trr1. All the assays described hereafter were performed using these purified E. coli-expressed proteins, even though identical activities were detected in extracts of S. pombe cells overexpressing the wild-type or mutant Tpx1 (data not shown).
|
2–3 µM. The H2O2-dependent inactivation of Tpx1 occurred at peroxide concentrations >50 µM. Thioredoxin has been suggested to act as a H2O2 peroxidase by itself (Hirota et al., 2002); so, to measure the H2O2-scavenging activity of Trx1 alone, we monitored NADPH oxidation by Trx1-Trr1 in the absence of Tpx1. As shown in Figure 2F, Trx1 was able to reduce H2O2, but with a Km value of at least 10 mM. This low affinity suggests that Trx1 is unlikely to contribute to H2O2 scavenging in vivo. Subtraction of the activity values for Trx1 (Figure 2F) from those of Tpx1 in the presence of the Trx1 system (Figure 2E) allowed us to calculate values for Tpx1 activity, which are shown in Figure 2G.
Overoxidation of Cys-48 of Tpx1 to a Cys-Sulfinic Acid Occurs at a Covalent Tpx1 Dimer Interface
Once established that Tpx1 suffers H2O2-mediated inactivation in vitro (Figure 2G), we decided to study whether such inactivation was concomitant with the oxidation of Cys-48 to the sulfinic acid state. To detect hyperoxidized Tpx1, we used commercial polyclonal antibodies against a sulfonylated peptide encompassing the active site of mammalian Prx, which is identical in sequence to that of S. pombe Tpx1. These antibodies recognize sulfinic (and sulfonic) forms of Prxs and permit the detection of sulfinylated Prx enzymes in extracts from H2O2-treated cells with high sensitivity and specificity (Woo et al., 2003b). The presence of oxygen during the protein purification procedure was able to induce the formation of a disulfide bond in Tpx1 (data not shown). Nevertheless, as described for other Prxs, H2O2 is not sufficient to cause hyperoxidation of the peroxidatic Cys-SH of Tpx1 to Cys-SO2H (Figure 3A). Instead, all catalytic components (Trx1, Trr1, and NADPH) are required. To our surprise, we detected two bands on immunoblots of in vitro hyperoxidized Tpx1 separated by nonreducing SDS-PAGE: a faster migrating band corresponding in size to a Tpx1–SO2H monomer, and another band migrating as a disulfide-linked Tpx1–SO2H dimer (Figure 3A). This dimeric form is DTT sensitive (data not shown), and it is not an artifact of purification procedures: immunoblot analysis of S. pombe extracts showed that the relative proportion of the monomeric and dimeric sulfinic acid forms depends on the H2O2 concentration (Figure 3B); in these blots, a nonspecific, H2O2-independent band was present only occasionally, and it is indicated with an asterisk. As shown in Figure 3B, a small amount of Tpx1–SO2H dimer is formed 5 min after exposure of cells to 0.2 mM H2O2, a concentration known to yield catalytically active Tpx1 dimers (Bozonet et al., 2005; Vivancos et al., 2005). On exposure to higher H2O2 concentrations, Tpx1 accumulated in the form of a hyperoxidised monomer (Figure 3B). To ascertain whether the formation of a Tpx1–SO2H dimer always precedes the accumulation of an inactive Tpx1–SO2H monomer, we performed an in vivo time course at 1 mM H2O2, which is the minimum concentration of peroxide known to fully inactivate Tpx1 (Vivancos et al., 2005). As seen in Figure 3C, a Tpx1–SO2H dimer can be detected only 5 s after exposure of the cell culture to 1 mM H2O2, which is then gradually transformed into the monomeric form.
|
; Vivancos et al., 2005). Tpx1 belongs to the typical 2-Cys Prx family. Members of this family are H2O2 scavengers, and the peroxidase reaction of these enzymes also requires both conserved Cys residues, because the oxidized enzyme intermediate generated during the catalytic cycle is a dimer in which the subunits are linked by one or two intermolecular disulfide bonds between the peroxidatic and the resolving Cys of the other subunit (for review, see Wood et al., 2003b). We analyzed what effect mutations of each one of the two Cys residues would have on cell viability. Surprisingly, a protein lacking the resolving Cys, Tpx1.C169S, was able to support aerobic growth of S. pombe cells (Figure 4A, middle), whereas the absence of the peroxidatic residue yielded a protein unable to sustain growth in the presence of oxygen. This result indicates that Tpx1.C169S is able to efficiently scavenge the H2O2 generated during aerobic growth. We subsequently measured protein carbonylation as an index of intrinsic oxidative stress. Under aerobic growth conditions the levels of protein carbonylation in extracts from cells expressing Tpx1.C169S were comparable with those of wild-type cells, whereas extracts from cells expressing Tpx1.C48S or lacking Prx enzymes showed significantly higher protein carbonylation (Figure 4B). Nevertheless, the ability of Tpx1.C169S to decompose high concentrations of peroxides is limited, because expression of Tpx1.C169S decreased cell tolerance to extracellular H2O2 to a greater extent than that of pap1 or srx1 cells (Figure 4A, right).
|
). Indeed, it has been demonstrated that DTT can support full activity in vitro of a Prx lacking its resolving Cys, whereas the Trx system cannot (Chae et al., 1994a). According to the model proposed by Rhee and colleagues, a molecule of DTT could replace the resolving Cys in the formation of a disulfide with the peroxidatic Cys-SOH, and subsequently another molecule of DTT could reduce the mixed disulfide. We assayed the activity of wild-type and mutated Tpx1 by measuring the removal of H2O2 from the reaction mixture by using the ferrithiocyanate method (see Materials and Methods). Wild-type Tpx1 was able to scavenge peroxides in the presence either of the Trx1 system or, weakly, of DTT (Figure 5B). The Tpx1–C48S mutant enzyme was completely inactive under all conditions assayed. The C169S mutant is able to scavenge H2O2 in the presence of either the Trx1 system or DTT. Moderate concentrations of GSH were also able to support Tpx1 and Tpx1.C169S activity. In an attempt to modulate the intracellular concentration of small reductants in vivo and regulate the activity of the Tpx1.C169S protein in aerobically growing cultures, we isolated S. pombe cells expressing the mutant protein in a cys1a background. The cys1a gene product, Cys synthase, is essential for the synthesis of L-Cys, and therefore cys1a cells are auxotrophic for the amino acid (Fujita and Takegawa, 2004). Therefore, decreasing the extracellular supply of L-Cys diminishes the intracellular availability of the amino acid in cys1a cells, thereby limiting the pool of intracellular GSH. We compared the sensitivity of cells lacking or possessing the cys1a gene and expressing wild-type or mutant Tpx1 under Cys-limiting conditions (0.08 mg/ml; Figure 5C). As a loading control, we incubated one plate under anaerobic conditions (Figure 5C, left). The double mutant cys1a tpx1.C169S displayed decreased viability in comparison with the single mutants cys1a or tpx1.C169S. Both the cys1a and the double mutant had a lower intracellular GSH concentration than cells possessing the cys1a gene (5-fold decrease; data not shown). The lower GSH concentration in the cys1a tpx1.C169S strains is probably not sufficient to support the enzymatic activity of the mutant Tpx1 protein, and H2O2 steady-state concentration rises under those conditions, generating oxidative stress and leading to cell death.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
), components of signal transduction pathways (Veal et al., 2004; Bozonet et al., 2005; Vivancos et al., 2005) and chaperones after heat stress (Jang et al., 2004; Chuang et al., 2006). The only 2-Cys Prx of S. pombe, Tpx1, has been previously reported to participate in signal transduction in response to mild extracellular oxidative stress, but we have demonstrated in the present study that it also plays an essential role as a scavenger of H2O2 generated during oxidative metabolism. tpx1 cells shifted from anaerobic to aerobic conditions suffered growth arrest; this was probably a consequence of intracellular oxidative stress, as indicated by their elevated levels of protein carbonylation. Steady-state levels of H2O2 in aerobically growing cells are reported to be 10–7 M (Seaver and Imlay, 2001). As shown for other Prxs, the Vmax of Tpx1 indicates that this is a slow detoxifying enzyme compared with catalases or GSH peroxidases; but its high-affinity for H2O2 (with a Km 2 µM) and its abundance (0.1–1% of total soluble proteins in S. pombe; data not shown) indicate that Tpx1 is very efficient at removing low concentrations of H2O2.
Other model organisms also have been shown to depend on Prxs for survival under aerobic conditions, such as the oxygen-sensitive microaerophile Helicobacter pylori (Baker et al., 2001). In the yeast S. cerevisiae, although cells lacking all five Prxs are still able to grow aerobically, they are hypersensitive to oxidative stress and are genomically unstable (Wong et al., 2004). The fission yeast S. pombe has a limited set of detoxifying enzymes compared with budding yeast. The reason for such a limited number of antioxidants is unknown, but this may explain why deletion of only one gene has such dramatic consequences for the fission yeast but not for S. cerevisiae.
Previous studies have demonstrated that both Cys residues of Tpx1 are required for Pap1 activation in response to low H2O2 stress (Bozonet et al., 2005; Vivancos et al., 2005). It also has been reported that the chaperone function of S. cerevisiae Tsa1 requires the peroxidatic but not the resolving Cys of this Prx (Jang et al., 2004). We can discount the participation of Tpx1 in heat-shock adaptation, because the growth of tpx1 cells at 37°C under anaerobic conditions was identical to that of a wild-type strain (data not shown). We show here that Tpx1.C169S is able to support aerobic growth, providing the first evidence of in vivo peroxidase activity of a 2-Cys Prx lacking the resolving Cys. As shown for other family members, Tpx1.C169S is also able to detoxify peroxides in vitro. Nevertheless, Tpx1.C169S is not fully functional, as shown by the impaired survival of a strain expressing this protein when exposed to mild extracellular H2O2 stress. Whether this limited peroxidase activity is a consequence of a decreased catalytic efficiency, an increased sensitivity to inactivation by peroxides, or both is yet to be determined.
Several H2O2 scavengers contain only one conserved Cys residue; for example, this is the case with mammalian GSH peroxidase, NADH peroxidase, and members of the 1-Cys Prx family (Kang et al., 1998a; Poole et al., 2004; Rhee et al., 2005b). These proteins catalyze the reduction of peroxides, although the identity of the electron donor to reduce and recycle the Cys-SeOH or Cys-SOH to selenothiol or thiol groups, respectively, is still unknown for some of these H2O2 scavengers. We have demonstrated here that small thiols, such as DTT or GSH, can support the peroxidase activity of Tpx1.C169S in vitro, probably because these molecules can donate thiol groups to form disulfide linkages with the peroxidatic Cys of the other subunit (Tpx1.C48-SOH), and they can subsequently reduce the resultant disulfides. In our experiments, Tpx1.C169S also supports peroxidase activity via the Trx1 system. Poole and coworkers showed that the resolving Cys of the Prx AhpC is not essential for in vitro peroxidase activity when an excess of the reducing enzyme, AhpF, is supplied (Ellis and Poole, 1997). Even though it is unlikely than Trx1 provides the reducing capacity for the activity of Tpx1.C169S in vivo, because its concentration is normally 1000-fold lower than that of GSH (Danon, 2002), specific Tpx1–Trx1 protein interactions could mediate a more efficient reaction with this reducing system. Therefore, whether the intracellular GSH pool constitutes the in vivo thiol donor for Tpx1.C169S activity will need further investigation. The results presented here provide the first evidence that a 2-Cys Prx, Tpx1, does not require the resolving Cys for its peroxidase activity in vivo, and we propose that Tpx1.C169S can function as a 1-Cys Prx both in vitro and in vivo.
It has previously been reported that Tpx1 can be reversibly substrate-inactivated via the formation of a sulfinic acid on its peroxidatic Cys (Bozonet et al., 2005; Vivancos et al., 2005). We have shown in this report that Tpx1 is inactivated by high H2O2 doses in vitro and that this inactivation occurs concomitantly with the formation of Cys-48-SO2H in Tpx1. Our in vivo and in vitro data indicate that oxidation of Cys-48-SH to sulfinic acid occurs when Tpx1 exists as a covalent dimer, before the accumulation of the inactive Tpx1 monomer. Rhee and coworkers proposed that during H2O2 detoxification the major covalent dimer formed in Prx enzymes contains two disulfide bonds, but that several minor dimeric forms carrying only one disulfide are also likely to be formed, resulting in a variable compactness and different mobilities on nonreducing SDS-PAGE (Chae et al., 1994c). The sulfinylated dimer in our experiments migrates slower than the major, two disulfide-containing dimer, and it has the same electrophoretic mobility as the minor species carrying only one disulfide (data not shown). Therefore, only the minor band corresponding to the one disulfide-containing Tpx1 dimer undergoes sulfinylation. According to structural studies on Prxs, the peroxidatic and resolving Cys residues are situated far apart in the corresponding Prx dimers (Wood et al., 2003b), and formation of a disulfide probably requires and induces significant conformational changes. Perhaps formation of the first disulfide in a Tpx1 dimer temporarily prevents the second Cys-48-SOH from stabilizing within its structural pocket in the active site, increasing its susceptibility to hyperoxidation to Cys-SO2H.
|
ACKNOWLEDGMENTS |
|---|
and of Xavi Gomis-Ruth (Institut de Biologia Molecular de Barcelona, CSIC, Barcelona, Spain) for the pGEX-2T-TEV plasmid. This work was supported by Dirección General de Investigación, Spain, grants BMC2003-00220 and BFU2006-02610, and "Distinció de la Generalitat de Catalunya per a la Promoció de la Recerca Universitaria. Joves Investigadors" DURSI (Generalitat de Catalunya) to E.H. |
Footnotes |
|---|
* These authors contributed equally to this work. 
Address correspondence to: Elena Hidalgo (elena.hidalgo{at}upf.edu)
Abbreviations used: DNPH, dinitrophenylhydrazine; DTT, dithiothreitol; GSH, glutathione; ORF, open reading frame; PAGE, polyacrylamide gel electrophoresis; Prx, peroxiredoxin; TCA, trichloroacetic acid.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Benson, F. E., Stasiak, A., West, S. C. (1994). Purification and characterization of the human Rad51 protein, an analogue of E.coli. RecA. EMBO J 13, 5764–5771.
Biteau, B., Labarre, J., Toledano, M. B. (2003). ATP-dependent reduction of cysteine-sulphinic acid by S.cerevisiae sulphiredoxin. Nature 425, 980–984.[CrossRef][Medline]
Bozonet, S. M., Findlay, V. J., Day, A. M., Cameron, J., Veal, E. A., Morgan, B. A. (2005). Oxidation of a eukaryotic 2-Cys peroxiredoxin is a molecular switch controlling the transcriptional response to increasing levels of hydrogen peroxide. J. Biol. Chem 280, 23319–23327.
Budanov, A. V., Sablina, A. A., Feinstein, E., Koonin, E. V., Chumakov, P. M. (2004). Regeneration of peroxiredoxins by p53-regulated sestrins, homologs of bacterial AhpD. Science 304, 596–600.
Cabiscol, E. and Levine, R. L. (1995). Carbonic anhydrase III. Oxidative modification in vivo and loss of phosphatase activity during aging. J. Biol. Chem 270, 14742–14747.
Chae, H. Z., Chung, S. J., Rhee, S. G. (1994a). Thioredoxin-dependent peroxide reductase from yeast. J. Biol. Chem 269, 27670–27678.
Chae, H. Z., Kim, H. J., Kang, S. W., Rhee, S. G. (1999). Characterization of three isoforms of mammalian peroxiredoxin that reduce peroxides in the presence of thioredoxin. Diabetes Res. Clin. Pract 45, 101–112.[CrossRef][Medline]
Chae, H. Z., Robison, K., Poole, L. B., Church, G., Storz, G., Rhee, S. G. (1994b). Cloning and sequencing of thiol-specific antioxidant from mammalian brain: alkyl hydroperoxide reductase and thiol-specific antioxidant define a large family of antioxidant enzymes. Proc. Natl. Acad. Sci. USA 91, 7017–7021.
Chae, H. Z., Uhm, T. B., Rhee, S. G. (1994c). Dimerization of thiol-specific antioxidant and the essential role of cysteine 47. Proc. Natl. Acad. Sci. USA 91, 7022–7026.
Chang, T. S., Jeong, W., Choi, S. Y., Yu, S., Kang, S. W., Rhee, S. G. (2002). Regulation of peroxiredoxin I activity by Cdc2-mediated phosphorylation. J. Biol. Chem 277, 25370–25376.
Chang, T. S., Jeong, W., Woo, H. A., Lee, S. M., Park, S., Rhee, S. G. (2004). Characterization of mammalian sulfiredoxin and its reactivation of hyperoxidized peroxiredoxin through reduction of cysteine sulfinic acid in the active site to cysteine. J. Biol. Chem 279, 50994–51001.
Chuang, M. H., Wu, M. S., Lo, W. L., Lin, J. T., Wong, C. H., Chiou, S. H. (2006). The antioxidant protein alkylhydroperoxide reductase of Helicobacter pylori switches from a peroxide reductase to a molecular chaperone function. Proc. Natl. Acad. Sci. USA 103, 2552–2557.
Danon, A. (2002). Redox reactions of regulatory proteins: do kinetics promote specificity? Trends Biochem. Sci 27, 197–203.[CrossRef][Medline]
Ellis, H. R. and Poole, L. B. (1997). Roles for the two cysteine residues of AhpC in catalysis of peroxide reduction by alkyl hydroperoxide reductase from Salmonella typhimurium. Biochemistry 36, 13349–13356.[CrossRef][Medline]
Fujita, Y. and Takegawa, K. (2004). Characterization of two genes encoding putative cysteine synthase required for cysteine biosynthesis in Schizosaccharomyces pombe. Biosci. Biotechnol. Biochem 68, 306–311.[CrossRef][Medline]
Hirota, K., Nakamura, H., Masutani, H., Yodoi, J. (2002). Thioredoxin superfamily and thioredoxin-inducing agents. Ann. N.Y. Acad. Sci 957, 189–199.
Ikner, A. and Shiozaki, K. (2005). Yeast signaling pathways in the oxidative stress response. Mutat. Res 569, 13–27.[Medline]
Jang, H. H., et al. (2004). Two enzymes in one; two yeast peroxiredoxins display oxidative stress-dependent switching from a peroxidase to a molecular chaperone function. Cell 117, 625–635.[CrossRef][Medline]
Kang, S. W., Baines, I. C., Rhee, S. G. (1998a). Characterization of a mammalian peroxiredoxin that contains one conserved cysteine. J. Biol. Chem 273, 6303–6311.
Kang, S. W., Chae, H. Z., Seo, M. S., Kim, K., Baines, I. C., Rhee, S. G. (1998b). Mammalian peroxiredoxin isoforms can reduce hydrogen peroxide generated in response to growth factors and tumor necrosis factor-alpha. J. Biol. Chem 273, 6297–6302.
Koo, K. H., Lee, S., Jeong, S. Y., Kim, E. T., Kim, H. J., Kim, K., Song, K., Chae, H. Z. (2002). Regulation of thioredoxin peroxidase activity by C-terminal truncation. Arch. Biochem. Biophys 397, 312–318.[CrossRef][Medline]
Levine, R. L., Williams, J. A., Stadtman, E. R., Shacter, E. (1994). Carbonyl assays for determination of oxidatively modified proteins. Methods Enzymol 233, 346–357.[Medline]
Maundrell, K. (1993). Thiamine-repressible expression vectors pREP and pRIP for fission yeast. Gene 123, 127–130.[CrossRef][Medline]
Millar, J. B., Buck, V., Wilkinson, M. G. (1995). Pyp1 and Pyp2 PTPases dephosphorylate an osmosensing MAP kinase controlling cell size at division in fission yeast. Genes Dev 9, 2117–2130.
Moreno, S., Klar, A., Nurse, P. (1991). Molecular genetic analysis of fission yeast Schizosaccharomyces pombe. Methods Enzymol 194, 795–823.[Medline]
Poole, L. B., Karplus, P. A., Claiborne, A. (2004). Protein sulfenic acids in redox signaling. Annu. Rev. Pharmacol. Toxicol 44, 325–347.[CrossRef][Medline]
Quinn, J., Findlay, V. J., Dawson, K., Millar, J. B., Jones, N., Morgan, B. A., Toone, W. M. (2002). Distinct regulatory proteins control the graded transcriptional response to increasing H(2)O(2) levels in fission yeast Schizosaccharomyces pombe. Mol. Biol. Cell 13, 805–816.
Rhee, S. G., Kang, S. W., Chang, T. S., Jeong, W., Kim, K. (2001). Peroxiredoxin, a novel family of peroxidases. IUBMB Life 52, 35–41.[Medline]
Rhee, S. G., Kang, S. W., Jeong, W., Chang, T. S., Yang, K. S., Woo, H. A. (2005a). Intracellular messenger function of hydrogen peroxide and its regulation by peroxiredoxins. Curr. Opin. Cell Biol 17, 183–189.[CrossRef][Medline]
Rhee, S. G., Kim, K. H., Chae, H. Z., Yim, M. B., Uchida, K., Netto, L. E., Stadtman, E. R. (1994). Antioxidant defense mechanisms: a new thiol-specific antioxidant enzyme. Ann. N.Y. Acad. Sci 738, 86–92.[Medline]
Rhee, S. G., Yang, K. S., Kang, S. W., Woo, H. A., Chang, T. S. (2005b). Controlled elimination of intracellular H(2)O(2): regulation of peroxiredoxin, catalase, and glutathione peroxidase via post-translational modification. Antioxid. Redox. Signal 7, 619–626.[CrossRef][Medline]
Seaver, L. C. and Imlay, J. A. (2001). Hydrogen peroxide fluxes and compartmentalization inside growing Escherichia coli. J. Bacteriol 183, 7182–7189.
Veal, E. A., Findlay, V. J., Day, A. M., Bozonet, S. M., Evans, J. M., Quinn, J., Morgan, B. A. (2004). A 2-Cys peroxiredoxin regulates peroxide-induced oxidation and activation of a stress-activated MAP kinase. Mol. Cell 15, 129–139.[CrossRef][Medline]
Vivancos, A. P., Castillo, E. A., Biteau, B., Nicot, C., Ayte, J., Toledano, M. B., Hidalgo, E. (2005). A cysteine-sulfinic acid in peroxiredoxin regulates H2O2-sensing by the antioxidant Pap1 pathway. Proc. Natl. Acad. Sci. USA 102, 8875–8880.
Vivancos, A. P., Castillo, E. A., Jones, N., Ayte, J., Hidalgo, E. (2004). Activation of the redox sensor Pap1 by hydrogen peroxide requires modulation of the intracellular oxidant concentration. Mol. Microbiol 52, 1427–1435.[CrossRef][Medline]
Wong, C. M., Siu, K. L., Jin, D. Y. (2004). Peroxiredoxin-null yeast cells are hypersensitive to oxidative stress and are genomically unstable. J. Biol. Chem 279, 23207–23213.
Woo, H. A., Chae, H. Z., Hwang, S. C., Yang, K. S., Kang, S. W., Kim, K., Rhee, S. G. (2003a). Reversing the inactivation of peroxiredoxins caused by cysteine sulfinic acid formation. Science 300, 653–656.
Woo, H. A., Kang, S. W., Kim, H. K., Yang, K. S., Chae, H. Z., Rhee, S. G. (2003b). Reversible oxidation of the active site cysteine of peroxiredoxins to cysteine sulfinic acid. Immunoblot detection with antibodies specific for the hyperoxidized cysteine-containing sequence. J. Biol. Chem 278, 47361–47364.
Wood, Z. A., Poole, L. B., Hantgan, R. R., Karplus, P. A. (2002). Dimers to doughnuts: redox-sensitive oligomerization of 2-cysteine peroxiredoxins. Biochemistry 41, 5493–5504.[CrossRef][Medline]
Wood, Z. A., Poole, L. B., Karplus, P. A. (2003a). Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling. Science 300, 650–653.
Wood, Z. A., Schroder, E., Robin Harris, J., Poole, L. B. (2003b). Structure, mechanism and regulation of peroxiredoxins. Trends Biochem. Sci 28, 32–40.[CrossRef][Medline]
Yang, K. S., Kang, S. W., Woo, H. A., Hwang, S. C., Chae, H. Z., Kim, K., Rhee, S. G. (2002). Inactivation of human peroxiredoxin I during catalysis as the result of the oxidation of the catalytic site cysteine to cysteine-sulfinic acid. J. Biol. Chem 277, 38029–38036.
This article has been cited by other articles:
|
|
 |
|
  M. Jara, A. P. Vivancos, and E. Hidalgo C-terminal truncation of the peroxiredoxin Tpx1 decreases its sensitivity for hydrogen peroxide without compromising its role in signal transduction. Genes Cells, February 1, 2008; 13(2): 171 - 179. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||
-DNP, top) or anti-Sty1 antibodies as a loading control (
