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Oxidative Stress and Cell Cycle Group, Universitat Pompeu Fabra, C/Dr. Aiguader 88, E-08003 Barcelona, Spain
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Abstract |
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CTD to detect and detoxify peroxides is impaired. Our results indicate that inactivation of Tpx1 by excess peroxides is not required for H2O2 signaling towards the Sty1 pathway, as expected from the floodgate model, and that the carboxy-terminal extension of Tpx1 concomitantly improves H2O2 scavenging and increases susceptibility to inactivation. |
Introduction |
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Upon severe H2O2 doses, the peroxidatic Cys of eukaryotic 2-Cys Prxs is selectively hyper-oxidized to Cys-sulfinic acid (SO2H) during catalysis, and this modification in-activates the peroxidase (Rabilloud et al. 2002; Yang et al. 2002). The sulfinic form of several eukaryotic Prxs can be enzymatically reduced to the active thiol form through the action of sulfiredoxins (Biteau et al. 2003; Woo et al. 2005) and/or sestrins (Budanov et al. 2004). This substrate-mediated inactivation of eukaryotic Prxs has been proposed to allow these enzymes to act as molecular floodgates, scavenging peroxides under basal conditions, but permitting higher concentrations during peroxide signal transduction (Woo et al. 2003).
It had been reported that bacterial Prxs are less sensitive to oxidative inactivation by a substrate excess than eukaryotic Prxs (Wood et al. 2003a). Sequence comparison and structural analysis of eukaryotic and prokaryotic Prxs helped to define the structural basis of sensitivity to substrate inactivation. The sequence alignment revealed two amino acid signatures—the GGLG and the YF motifs—inserted in two distal regions of Prx proteins, and present only in the peroxide-sensitive eukaryotic enzymes, not in bacterial homologues. Structural comparison indicated that these motifs of sensitive Prxs stabilize a fully folded, rigid conformation, in which the peroxidatic Cys in the sulfenic acid form is susceptible to substrate-mediated inactivation (Koo et al. 2002; Wood et al. 2003a).
The only 2-Cys Prx in Schizosaccharomyces pombe, Tpx1, seems to be an essential component of the complex signal transduction program of the fission yeast in response to oxidative stress. Thus, there are two main pathways, which respond to extracellular H2O2: the Pap1 and the Sty1 pathways (for reviews, see Vivancos et al. 2006; Veal et al. 2007). The Pap1 transcription factor is more sensitive to H2O2 than the MAP kinase Sty1 pathway, and thus Pap1 is known to induce adaptation, whereas Sty1 promotes survival responses. Tpx1 was described as the initial sensor of the Pap1 pathway, which responds to moderate H2O2 stress (Bozonet et al. 2005; Vivancos et al. 2005). Thus, oxidation of Tpx1 by H2O2 will in turn activate Pap1 by inducing the formation of an intramolecular disulfide bond in the transcription factor to hinder its nuclear export signal, what leads to its nuclear accumulation and to the Pap1-dependent gene response. Since Tpx1 is substrate inactivated, severe doses of H2O2 will reversibly inactivate Tpx1 and postpone Pap1 activation. The same severe oxidant dose will then fully activate the Sty1 pathway by an unknown mechanism. Sty1 activation also leads to the induction at the transcriptional level of a complex anti-stress program that includes the synthesis of the sulfiredoxin Srx1, which is charged with recycling Tpx1 and triggering the Pap1 pathway (Bozonet et al. 2005; Vivancos et al. 2005). This inactivation shunt avoids overlap between the Pap1 and Sty1 pathways upon severe H2O2 stress, but its biological relevance remains unknown. It has also been suggested a role of Tpx1 in proper activation of the Sty1 pathway in response to H2O2 (Veal et al. 2004). Furthermore, we have recently described that Tpx1 is essential for aerobic growth because of its role as a H2O2 peroxidase (Vivancos et al. 2006; Jara et al. 2007). Thus, Tpx1 functions in S. pombe as a peroxide scavenger and as an activator of signal transduction pathways. However, it is not clear whether the sensitivity to inactivation by an excess of peroxides is important for its role activating Pap1 and/or Sty1. In fact, the Gpx3-Yap1 redox-relay of Saccharomyces cerevisiae, orthologous to the Tpx1-Pap1 system, does not suffer substrate-mediated temporal inactivation: Gpx3 is a glutathione peroxidase, and it does not become inhibited at high concentrations of peroxides (Delaunay et al. 2002).
To test whether the temporal inactivation of Tpx1 by an excess of H2O2 is essential for proper induction of the Pap1 and Sty1 signal transduction pathways and for an optimal adaptation of fission yeast to oxidative stress, we have modified Tpx1 in S. pombe to produce a Prx less sensitive to oxidative inactivation. To do so, we have generated a C-terminal truncated Tpx1.CTD, and analyzed the effect of this mutant protein on cell growth and anti-stress gene induction. We show that Tpx1.CTD is indeed not inactivated at the same peroxide concentrations that inactivate the wild-type protein. Cells expressing this mutant Prx are sensitive to aerobic growth conditions, but gene induction by the Pap1 and Sty1 pathways is not disrupted. Instead, the truncated version of Tpx1 has an apparent Km for H2O2 tenfold higher than that of the wild-type protein, and thus the enzyme is less efficient than the wild-type form at detoxifying the H2O2 generated during oxidative metabolism.
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Results |
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To test the effect of the C-terminal extension on the sensitivity of Prxs to substrate-mediated inactivation, we constructed truncated versions of the tpx1 open reading frame (ORF) to yield proteins lacking 17 aa (Tpx1.CTD) or 9 aa (Tpx1.CTD183) of the CTD of the wild-type protein (Fig. 1A). We tested the sensitivity of strains expressing HA-tagged wild-type Tpx1 or one of the truncated forms to peroxide-mediated inactivation by measuring the Tpx1 monomer/covalent dimer ratio by non-reducing electrophoresis of trichloroacetic acid (TCA) extracts (Fig. 1B). Wild-type Tpx1 covalent dimers are barely observed in cells treated with H2O2 concentrations over 1 mM, whereas this concentration did not compromise dimer formation by the truncated proteins Tpx1.CTD or Tpx1.CTD183. We then confirmed the oxidation of Cys-48 of wild-type Tpx1 to the sulfinic acid state by immunodetection in TCA extracts with commercial anti-Prx-SO3 antibodies (Fig. 1C, left panel). The sulfinylated forms of Tpx1.CTD and Tpx1.CTD183 could not be detected in extracts from cells exposed to H2O2 concentrations from 0.2 to 5 mM (Fig. 1C, center and right panels). Since both truncated proteins behaved identically, Tpx1.CTD was used in all subsequent experiments.
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CTD activates Pap1 in response to low and high doses of H2O2
We tested whether the activation of Pap1 by Tpx1.CTD would retain the same kinetics and Srx1-dependence as the wild-type enzyme. As shown in Fig. 2A, the mutant Prx is able to trigger Pap1 activation 5 min after 1 mM oxidant stress, whereas wild-type Tpx1 is not. Furthermore, the absence of Srx1 did not alter the kinetics of Pap1 activation by Tpx1.CTD. We tested whether the C-terminal deletion affected cell sensitivity to peroxides or gene activation by the Pap1 or Sty1 pathways. Cells expressing Tpx1.CTD were more sensitive than a wild-type strain not only to 1 mM extracellular H2O2 (Fig. 2B, right panel), but also to aerobic growth (Fig. 2B, left panel). However, the induction of Sty1-dependent genes was not altered (Fig. 2C), whereas the induction of Pap1-dependent genes was faster than in a wild-type strain, as expected (compare kinetics of Fig. 2A and C).
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CTD are sensitive to aerobic growth conditions
We tested whether the peroxidase activity of Tpx1.CTD was compromised so that cells expressing only this truncated form would suffer from intrinsic oxidative stress during aerobic growth. Under anaerobic conditions, the growth profiles of wild-type cells and cells expressing Tpx1.CTD were the same (Fig. 3A), confirming that the expression of Tpx1.CTD affects the sensitivity of cells only to aerobic growth conditions. The sensitivity of a strain expressing Tpx1.CTD to grow under aerobic conditions is not as severe as that of a strain lacking tpx1, as expected (Fig. 3A). The impaired aerobic growth was reflected in increased protein carbonylation, which provides an index of protein damage resulting from oxidative stress; when shifted from anaerobic to aerobic growth, cells expressing Tpx1.CTD contained almost as many carbonylated proteins as a strain lacking Tpx1 (Fig. 3B). Even under anaerobic conditions, expression of Tpx1.CTD impairs cell survival upon extracellular H2O2 stress (Fig. 3A).
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CTD for H2O2 is tenfold higher than that of wild-type Tpx1
To verify whether Tpx1.CTD had a decreased affinity for H2O2, we measured the Tpx1 monomer/covalent dimer ratio as in Fig. 1B, but using lower doses of extracellular peroxide. Concentrations as low as 0.01 mM were sufficient to trigger disulfide formation and thus dimerization by wild-type Tpx1 (Fig. 4A, upper panel). In contrast, the minimum concentration of extracellular peroxide able to induce dimer formation by Tpx1.CTD was 0.05–0.1 mM (Fig. 4A, lower panel).
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CTD, and various concentrations of H2O2. The resulting Prx-dependent activities, obtained after subtracting the peroxidase activity linked to Trx1–Trr1 (see Experimental procedures), are shown in Fig. 4B. As shown before for other Prxs, the substrate-mediated inactivation of wild-type Tpx1 at high H2O2 concentrations distorted the typical Michaelis kinetics. Thus, we applied Lineweaver–Burk plots to the Tpx1 and Tpx1.CTD activities at concentrations from 5 to 50 µM in the case of wild-type Tpx1, and from 5 to 100 µM for mutant Tpx1.DCTD. Thus, the apparent Km of wild-type Tpx1 for H2O2 was c. 2 µM, as described earlier (Jara et al. 2007) (Fig. 4B, left panel). However, the affinity of Tpx1.DCTD for H2O2 (with an apparent Km of 20 µM) was significantly lower than that of the wild-type enzyme. The same apparent Km was observed for Tpx1.DCTD183 (data not shown). Very high concentrations of H2O2 can trigger sulfinic acid formation in Tpx1.DCTD
According to the in vitro peroxidase activities described above for Tpx1.DCTD (Fig. 4B, right panel), this mutant Prx also seems to be susceptible to inactivation by peroxides, since at concentrations above 5 mM the peroxidase activity was markedly diminished. Two further lines of evidence demonstrate that Tpx1.DCTD can be hyper-oxidized to the sulfinic acid form by an excess of peroxide: (i) Higher concentrations of H2O2 (5–25 mM) are less efficient than lower concentrations at activating Pap1 in cells expressing Tpx1.DCTD (Fig. 5A); and (ii) we could immunodetect hyper-oxidized Tpx1.DCTD–SO2H with anti-Prx-SO3 antibodies upon exposure of the enzyme to very high doses of H2O2 both in vivo (Fig. 5B) and in vitro (Fig. 5C).
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Discussion |
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Cells expressing Tpx1.DCTD are sensitive to aerobic, but not anaerobic, growth. This phenotype cannot be consequence of impaired Pap1 or Sty1 activation by peroxides, since deletion of neither pap1 nor sty1 genes compromises cell viability in the absence of extracellular stress (for a review, see Ikner & Shiozaki 2005). Instead, we have shown here that the mutant protein has a compromised sensitivity for its substrate, H2O2, with an apparent Km tenfold higher than that of the wild-type enzyme. Prxs are inefficient peroxidases, but are adapted to maintain low levels of peroxides based on their low Km and their abundance. In the case of Tpx1.DCTD, the affinity for H2O2 is impaired and therefore cells expressing it are handicapped and show markers of intracellular oxidative stress when grown in the presence of oxygen.
We have demonstrated here that disruption of the C-terminal structure of the Prx Tpx1 makes the enzyme resistant to hyper-oxidation to the sulfinic acid form, but the capacity of the mutated enzyme for H2O2 scavenging is impaired, as well. In fact, it had already been reported in vitro that proteolytic C-terminal truncation of Tpx1 yielded mutant protein forms more resistant to H2O2-mediated inactivation than the intact Tpx1, and that such truncations may have also affected the sensitivity of the protein to detoxify H2O2 (Koo et al. 2002). A later report suggested that two motifs unique to eukaryotic 2-Cys Prxs—the GGLG insertion in the center of the polypeptide and the CTD extension—define the structural origins of sensitivity to oxidant-mediated inactivation (Wood et al. 2003a). Here, we confirm in vitro (Koo et al. 2002) and in vivo that only one of these motifs, the C-terminal extension, is sufficient to modify the propensity of a eukaryotic Prx both for thiol to sulfenic acid oxidation (the basis of H2O2 peroxidase activity) and for sulfenic acid to sulfinic acid oxidation (the basis of Prx inactivation). It is important to point out that, when comparing Tpx1.DCTD and wild-type Tpx1, the extent of affinity loss for the thiol to sulfenic acid oxidation (tenfold higher apparent Km for the truncated protein) is less pronounced than the increased resistance to hyper-oxidation by sulfinic acid formation (20-fold higher H2O2 concentration is required to inactivate Tpx1.DCTD).
According to published reports, the prokaryotic Prxs, AhpC from E. coli and Salmonella thyphimurium, have a low Km for H2O2 similar to eukaryotic Prxs, but require at least 100-fold more H2O2 for inactivation (Wood et al. 2003a). It may be that the absence of the GGLG motif or another structural feature, in addition to the lack of C-terminal extension, of these robust enzymes maintains affinity without increasing the inactivation rate. This might involve bringing the peroxidatic and resolving Cys residues closer so as to favor disulfide formation after oxidation of the thiol group to sulfenic acid, the disulfide group being stable and resistant to further oxidation (Claiborne et al. 1999). A mutational analysis of prokaryotic Prxs and the isolation of a robust enzyme with an impaired Km for H2O2 would contribute to the biochemical analysis of this family of enzymes.
Prxs containing the GGLG and YF motifs are not exclusive from the Eukarya domain. We have found sequences coding for GGLG-containing Prxs in bacteria as ancient as Firmicutes. In these cases, the coded proteins do not have the C-terminal YF motif. In some genera of this phylum, these eukaryotic-like Prx genes co-exists with ahpC-like sequences. However, they show higher similarity to S. pombe Tpx1 than to their own ahpC-like Prxs. In less ancient bacterial groups such as Cyanobacteria, Chlorobi, Spirochetaetes or Proteobacteria, we have found sequences coding for Prxs containing both the GGLG and the YF motifs. Taking all the sequences together, BLAST searches always separate two families of enzymes, and members of both families can be found in prokaryotes, whereas eukaryotes seem to have inherited members of one class only. This suggests that Prxs, sensitive or robust to oxidative inactivation, evolved from a common ancestor and diverged to fulfill different cellular functions early in evolution. The reductase Srx1 is apparently absent from all prokaryotes and its presence probably denotes a further step in evolution.
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Experimental procedures |
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We used the wild-type strains 972 (h–), JA212 (h+ leu1) and PN513 (h– leu1 ura4), as well as other published strains such as EA40hap-p123.41x (h–leu1 tpx1::kanMX6 pHA-tpx1.41x) (Vivancos et al. 2005), EA56 (h+ leu1 srx1::kanMX6) (Vivancos et al. 2005), AV15 (h–atf1::kanMX6) (Zuin et al. 2005), AV18 (h–sty1::kanMX6) (Zuin et al. 2005), AV25 (h–pap1::kanMX6) (Zuin et al. 2005) and AV42 (h+ leu1 ura4 ade6-M210 tpx1::kanMX6) (Jara et al. 2007). In order to generate an S. pombe strain expressing Tpx1.DCTD, we transformed strain JA212 with a linear tpx1.DCTD::kanMX6 fragment, obtained by PCR amplification with ORF-specific mutagenic primers, designed to delete the last 17 aa of tpx1, and plasmid pFA6a-kanMX6 as a template, yielding strain AV33 (h+ leu1 tpx1.DCTD::kanMX6). We disrupted the srx1 ORF by transformation of strain PN513 with a PCR-amplified srx1::kanMX6 fragment, yielding strain EA55 (h– leu1 ura4 srx1::kanMX6). To construct a strain deleted in srx1 and expressing Tpx1.DCTD, we crossed strains AV33 and EA55, yielding strain AV37 (h+ leu1 tpx1.DCTD::kanMX6 srx1::kanMX6).
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 sachet (Merck, Whitehouse Station, NJ). In order to analyze sensitivity to H2O2 on plates, S. pombe strains were grown aerobically or anaerobically, as indicated, in EMM liquid media to an OD600 of 0.5. Cells were then diluted in water, and the number of cells indicated at the top of the panels in 4 µL was spotted onto YE5S solid media in the presence or absence of H2O2. The spots were allowed to dry, and the plates were incubated at 30 °C under aerobic or anaerobic conditions, for 3–5 days.
Plasmids
We used plasmids p123.41x (pHA-tpx1.41x) (Vivancos et al. 2005) and p124.41x [constructed as p123.41x, after PCR amplification of the tpx1 ORF from codon 1 to 175 with specific mutagenic primers including BamHI and SmaI restriction sites, and cloned into pRep.41x (Maundrell 1993); the plasmid expressed HA-Tpx1.CTD]. A similar strategy was used to construct p176.41x (carrying the tpx1 ORF from codon 1 to 183; the plasmid expresses HA-Tpx1.CTD183). The plasmids p123.41x, p124.41x and p176.41x carry wild-type and mutant tpx1 ORFs under the control of the nmt (no message in thiamine) promoter. In order to express S. pombe proteins in E. coli, the ORFs coding for Tpx1, Tpx1.CTD and Tpx1.CTD183 were digested from plasmids p123.41x, p124.41x and p176.41x with BamHI and SmaI, and subcloned into a modified GST-tagging fusion vector pGEX-2T-TEV, that encoded a TEV protease cleavage site between the tag and the cloned ORF. The resulting plasmids were called p210, p203 and p204, respectively. The trr1 and trx1 ORFs were amplified and cloned into the pGEX-2T-TEV expression vector as described before (Jara et al. 2007).
Preparation of S. pombe TCA extracts and immunoblot analysis
For in vivo redox state analysis of Pap1, TCA extracts were prepared as described before (Vivancos et al. 2005). Regarding Tpx1 analysis, tpx1 strains carrying plasmids p123.41x, p124.41x or p176.41x and expressing HA-Tpx1, HA-Tpx1.CTD and HA- Tpx1.CTD183, respectively, were pre-treated with thiamine for 5 h in order to obtain wild-type levels of Tpx1 protein, as described before (Vivancos et al. 2005). Analysis of the redox state of wild-type and truncated Tpx1 proteins (expressed from integrated tpx1 genes or episomal plasmids) was performed following the same protocol as for Pap1, except that after extract alkylation, samples were not dephosphorylated, then they were also diluted fivefold and electrophoretically separated by non-reducing SDS/PAGE. Proteins were immunodetected using polyclonal anti-Pap1 antibodies (Vivancos et al. 2004), monoclonal anti-HA antibodies (when detecting the HA-tagged forms of Tpx1) or polyclonal anti-Tpx1 (Jara et al. 2007). For the detection of sulfinylated Tpx1, the undiluted alkylated samples were separated by reducing SDS/PAGE, and immunodetected with anti-Prx-SO3 antibody (LabFrontier, Seoul, Korea) or anti-HA antibodies as a loading control.
RNA analysis
Total RNA from S. pombe cultures was obtained, processed and transferred to membranes as described before (Vivancos et al. 2004). Membranes were hybridized with the 
-32P-dCTP-labeled trr1, ctt1, srx1, gpx1 and cdc2 probes, containing the complete ORFs of the thioredoxin reductase-, catalase-, sulfiredoxin-, glutathione peroxidase- and cyclin-dependent protein kinase-coding genes.
Preparation of S. pombe extracts to measure protein carbonylation
Protein carbonylation was detected upon in vitro derivatisation of oxidized proteins with 2,4-dinitrophenylhydrazine (DNPH) (Levine et al. 1994; Cabiscol & Levine 1995). Cells from anaerobic pre-cultures were grown aerobically over a period of 12 h in 50 mL of rich media, and harvested at an OD600 of 0.5. Cell pellets were resuspended in carbonylation buffer, lysed with glass beads, and carbonyl groups of proteins in cell extracts were derivatised with DNPH. The modification was then immunodetected with anti-DNP antibodies, as described (Jara et al. 2007). As a loading control, Sty1 was detected using polyclonal anti-Sty1 antibodies (Jara et al. 2007).
Purification of recombinant Tpx1, Tpx1.CTD, Tpx1.CTD183, Trx1 and Trr1 proteins, and peroxidase activity assays
Bacteria strain FB810 (Benson et al. 1994) transformed with the pGEX-2T-TEV derivatives were inoculated into LB broth with 100 µg/mL of ampicillin and incubated at 37 °C for c. 16 h with vigorous shaking. GST-tagged proteins were purified as described (Jara et al. 2007). We released the S. pombe proteins without the GST tags after incubation with TEV protease (Invitrogen, Carlsbad, CA) (Jara et al. 2007).
NADPH oxidation was monitored as a decrease in optical density at 340 nm using a UltrospecTM 3100 pro UV/Visible Spectrophotometer in a 500-µL reaction mixture containing 50 mM HEPES–NaOH pH 7.0, 0.25 mM NADPH, 6 µg Trr1, 20 µg Trx1, 1 µg of Tpx1, Tpx1.CTD, or Tpx1.CTD183 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 previously described (Koo et al. 2002). We performed the same measurements in the absence of Tpx1 or Tpx1.CTD to obtain the peroxidase activity linked to Trx1–Trr1 (Hirota et al. 2002), and subtracted these values from the total.
Detection of in vitro over-oxidized Tpx1–SO2H or Tpx1.CTD–SO2H
In order to detect the Tpx1–SO2H forms in vitro, a reaction with the same conditions as described for the peroxidase activity assay was used with 5 µg of Tpx1 or Tpx1.CTD. After 10 min, the reaction was stopped by the addition of TCA 100% to a final 12.5% concentration. Proteins were pelleted, washed in acetone and dried. Samples were resuspended in loading buffer (50 mM Tris pH 6.8, 4% glycerol, 1.4% SDS, saturated bromophenol blue and 100 mM dithiothreitol). A measure of 2.5 µg of total Tpx1 or Tpx1.CTD was resolved in 12% reducing SDS/PAGE followed by Western blot analysis using anti-Prx-SO3 antibody (LabFrontier). As a loading control, fivefold diluted samples were used for Western blot analysis using anti-Tpx1 antibody.
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Acknowledgements |
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Footnotes |
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aThese authors contributed equally to this work.
bPresent address: Barcelona Science Park, Barcelona, Spain
* Correspondence: E-mail: elena.hidalgo{at}upf.edu |
References |
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Accepted: 11 November 2007
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