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Address correspondence to Jakob R. Winther, Dept. of Physiology, Carlsberg Laboratory, Gamle Carlsberg Vej 10, DK-2500 Copenhagen Valby, Denmark. Tel.: 45-3327-5282. Fax: 45-3327-4765. email: jrw{at}crc.dk
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Abstract |
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Key Words: green fluorescent protein; glutathione; glutaredoxin; redox; oxidation
Abbreviations used in this paper: 4-DPS, 4,4'-dithiodipyridine; GSH, reduced glutathione; GSSG, oxidized glutathione; NEM, N-ethyl-maleimide; NPM, N-(1-pyrenyl)maleimide; rGrx1p, recombinant His-tagged yeast Grx1p; rxYFP, redox sensitive YFP; TCA, trichloroacetic.
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Introduction |
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Glutathione homeostasis is maintained through a poorly understood balance between synthesis, uptake, usage, breakdown, and excretion. The biosynthetic pathway has been studied in some detail in the yeast Saccharomyces cerevisiae. Here, synthesis takes place in the cytosol, in two ATP-dependent condensation reactions catalyzed by -glutamylcysteine synthetase, encoded by the GSH1 gene, and glutathione synthetase (GSH2), respectively (Ohtake and Yabuuchi, 1991; Grant et al., 1997). Although deletion of the GSH1 gene confers an auxotrophic requirement for glutathione, GSH2 can be disrupted without fatal consequences, suggesting that
-glutamylcysteine can functionally replace GSH (Grant et al., 1996, 1997; Spector et al., 2001). Uptake of GSH and GSSG from the culture medium is mediated by the high affinity glutathione transporter Hgt1p. As a gsh1 hgt1 double mutant is unable to grow on glutathione as the sole sulfur source, it is likely that Hgt1p is the only glutathione-transporting activity in the yeast plasma membrane (Bourbouloux et al., 2000; Hauser et al., 2000). Glutathione catabolism is thought to be initiated by the hydrolytic cleavage of the unusual
-peptide bond between glutamate and cysteine, whereby cysteinylglycine and glutamate are released. This reaction is catalyzed by
-glutamyl transpeptidase (for review see Taniguchi and Ikeda, 1998). A single
-glutamyl transpeptidase homologue, which is the product of the ECM38 gene, has been identified in yeast (Jaspers and Penninckx, 1984). However, additional pathways must be capable of degrading GSH, as an ECM38 knockout mutant is able to survive on glutathione as the only source of sulfur (Kumar et al., 2003).
A major pool of glutathione appears to be sequestered in the yeast vacuole, where it is actively imported as glutathione and glutathione S-conjugates by the yeast cadmium factor protein (Ycf1p), and to a lesser extent by its paralogue Bpt1p (Li et al., 1996; Rebbeor et al., 1998; Klein et al., 2002). Whether GSH, and perhaps in particular GSSG, is also delivered to the vacuole by vesicular transport as a "spillover" from the secretory pathway is intriguing, but has not yet been explored.
In the present paper, we apply a previously designed GFP-based sensor for disulfide bond formation (Ostergaard et al., 2001) to study the cytosolic thiol-disulfide redox status. The fluorescence of the sensor is modulated by the reversible formation/reduction of a solvent-exposed disulfide bond. Using yeast as a model organism, we show that this sensor is uniquely sensitive to the glutathione redox pair and not targeted by the thioredoxin pathway. Moreover, it possesses a dynamic range that is perfectly suited for measuring the glutathione redox status under normal as well as redox-compromised conditions.
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Results |
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Rapid equilibration of rxYFP in the cytosol
The fluorometric redox titration method is a steady-state technique. As such it provides a snapshot of the intracellular distribution of oxidized and reduced sensor, governed by the balancing action of synthesis of nascent reduced rxYFP and its subsequent oxidation. To evaluate the extent to which the measured steady-state distributions correlated with the equilibrium distribution, the in vivo kinetics of rxYFP oxidation was assessed by pulse-chase immunoprecipitation analysis in the wild-type, glr1, and glr1 met15 mutants, with intracellular GSSG/GSH ratios ranging from low to highly elevated. Cells were labeled with [35S]Met/Cys for 10 min and then chased for 0, 15, 30, and 60 min at which times thiol reactions were quenched by addition of TCA. Subsequently, free thiols were blocked by treatment with the alkylating reagent NEM. SDS-PAGE and phosphorimaging of immunoprecipitated rxYFP revealed that equilibrium was rapidly attained, with a half-time around 10 min irrespective of the intracellular GSSG/GSH ratio (Fig. 2). The fundamental difference between steady state and equilibrium became most apparent in the glr1 mutant, with an intermediate GSSG/GSH ratio. Here, the proportion of oxidized rxYFP was 73% according to the pulse-chase end point distribution, as opposed to 56% by the fluorometric method (compare Fig. 1 B with Fig. 2). This finding showed that whereas the fluorometric method constituted a highly efficient means of monitoring changes in the intracellular GSSG/GSH ratio, pulse-chase end point redox distribution analysis was required to obtain an accurate measure of the glutathione redox status.
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Applying this equation on the equilibrium data obtained from the pulse-chase analysis allowed us to estimate the absolute cytosolic concentrations of GSH and GSSG (Fig. 3 A). Because the precise pH of the cytosol was not known, values for the two predefined parameters Kox and kox were measured in vitro in the pH range from 6.1 to 7.9 (Table II). Using a nonlinear least squares procedure and Kox and kox parameters determined at pH 7, a close fit to the pulse-chase data was obtained which yielded GSH and GSSG concentrations of 13 and 0.17 mM, respectively. Due to the robustness of Kox to changes in pH around 7 (Table II) the estimated concentrations only displayed a minor pH dependency. Accordingly, if the fit was extended to cover pH values ranging from 6.7 to 7.3, which should include most of the cytosolic pH values reported in the literature (Melvin and Shanks, 1996; Breeuwer and Abee, 2000), GSH and GSSG concentrations were found to lie within a range of 1019 mM and 0.10.3 mM, respectively.
The wild-type cytosolic glutathione redox potential is surprisingly reducing
As mentioned above, steady-state measurements will underestimate the fraction of oxidized rxYFP compared with pulse-labeling end points. As the latter reflect the equilibrium of rxYFP with the glutathione redox pair, the intracellular glutathione redox potential, in turn, can be derived from the end point redox distribution. Using the Nernst equation and 265 mV as the standard redox potential (E°') of rxYFP calculated from Kox at pH 7.0 (Table II), we found the redox potential of the cytosolic glutathione pool in wild-type yeast cells to be 289 mV (Table I). In our whole-cell measurements the amount of GSH relative to cell density only displayed very little variation (<10% from wild type to glr1 grx1 grx2) in spite of large differences in GSSG/GSH ratios (Table I). This strongly argued that the GSH concentration (13 mM) that could be derived from the glr1 grx1 grx2 mutant was also valid for the wild type. Thus, using this concentration and a redox potential of 289 mV as calculated above, the concentration of GSSG in the cytosol of wild-type yeast was estimated to be 4 µM; a value considerably lower that would be anticipated on basis of glutathione measurements on whole-cell extracts.
Targeting of rxYFP to other cellular compartments
One of the obvious virtues of using a GFP-based sensor is its genetic nature, which, in principle, allows it to be targeted to noncytosolic compartments like the ER. We would like to note that we have found that several signal peptides commonly used for secretion of heterologous proteins (-factor, carboxypeptidase Y, invertase) did not promote efficient translocation of rxYFP (unpublished data). We have nevertheless constructed a variant that by several criteria appears to be translocated efficiently. However, the redox potential of rxYFP is, in its current version, not optimally suited for reliable redox measurements in this or other secretory compartments.
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Discussion |
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We found that at steady state 10% of rxYFP was oxidized in the cytosol of wild-type yeast. The degree of oxidation was consistent from determinations on a growing culture of yeast cells by fluorescence, from quantification of Western blots (unpublished data), and from pulse-labeling experiments. By introduction of various mutations and changing the growth medium we were able to manipulate the ratio between total GSSG and GSH from 4% in the wild type to 225%. The most extreme situation was attained by combining the glr1 mutation with a mutation in the sulfur assimilation pathway (met15) and growing the cells on GSSG as the sole sulfur source (Fig. 1 C). To our knowledge, such a GSSG/GSH ratio has not previously been measured in any living cells. Interestingly, the cellular glutathione homeostasis maintains essentially the same total pool of GSH whereas the GSSG varies (see Table I). Moreover, there is a consistent correlation between the GSSG/GSH ratio in the various mutants and the measured fraction of oxidized rxYFP, which increases to >90% in the most extreme case, suggesting a direct effect of the glutathione redox status on the redox status of rxYFP (Fig. 1 D).
As any protein is initially produced in the reduced state, the steady-state fraction of oxidized rxYFP might reflect a complete, but very slow, oxidation. To determine the oxidation kinetics and equilibrium redox distribution of rxYFP, pulse-labeling experiments were performed, followed by immunoprecipitation and separation of oxidized and reduced forms by nonreducing SDS-PAGE. In these experiments it could be shown that equilibrium was reached fairly rapidly in the wild type and in the glr1 and glr1 met15 mutants (Fig. 2), much more rapidly than in vitro kinetics would predict for an uncatalyzed reaction (Table II). We find that glutaredoxins catalyze the rapid equilibration of the sensor with the intracellular glutathione buffer, a conclusion we base on two observations. In vivo the rate of oxidation is significantly slowed down in glr1 grx1 grx2 mutant cells, deleted for the two cytosolic dithiol-type glutaredoxins (Fig. 2 and Fig. 3 A). In support of this, we show that glutaredoxin can catalyze the oxidation of reduced rxYFP as well as the reverse reaction in vitro (Fig. 4). Although this makes perfect sense from a biochemical point of view, it exposes a rather unconventional role for a glutaredoxin in vivo, where it is normally attributed to reduction of disulfide bonds (Grant, 2001). Similar in vitro experiments using yeast cytosolic thioredoxins showed catalysis of neither oxidation nor reduction (Fig. S1). This is an important observation as reduction of rxYFP by thioredoxin could in principle have formed a rapid kinetic link to NADPH oxidation through thioredoxin reductase. Instead, however, we find that the properties of rxYFP make it uniquely suited to the glutathione redox pair in the cytosol and that equilibration with this redox buffer is catalyzed by glutaredoxins.
Our estimates of absolute concentrations of GSH and GSSG are based on only two assumptions: (1) a cytosolic pH between 6.7 and 7.3; and (2) the absence of other catalysts than glutaredoxin 1 and 2. Although we have no direct evidence for the latter, it seems, however, highly reasonable given the slow equilibration kinetics observed in the glr1 grx1 grx2 triple knockout. It should also be emphasized that the presence of additional redox catalysts would imply an even lower cytosolic GSSG concentration than the one estimated here. If the GSH concentration of this mutant is close to that of the wild type, as supported by whole cell measurements, we find the cytosolic GSSG concentration of the wild type to be surprisingly low, 4 µM. Although it may be intuitively pleasing to know the absolute concentrations of oxidized and GSH in the cytosol, these figures may be difficult to interpret in a cytosolic environment where conditions are probably far from ideal due to high concentrations of proteins and low molecular weight solutes. In terms of disulfide bond stability the glutathione redox potential is, however, the central redox parameter on which to gauge the cytosolic redox environment. We find a value of 289 mV, which can be derived from the redox status of rxYFP using the Nernst equation. Although there is no clear consensus in the literature, this figure is significantly lower than previous estimates (e.g., 232 mV determined in a work by Hwang et al., 1992). Indeed, GSSG/GSH ratios measured on whole-cell extracts is considerably higher (4% in the wild type) than what we find in the cytosol (0.03%). Even under the most extreme conditions in the grl1 met15 strain grown on 1 mM GSSG the cytosol remains fairly reducing (Table I,
227 mV, which is equivalent to
4% GSSG), suggesting that the vast majority of the more than twofold excess of GSSG measured in cell extracts of this strain is sequestered (Table I). It is not unlikely that there is a considerable excess of GSSG in the secretory pathway including the vacuole and ER (Hwang et al., 1992; Li et al., 1996; Cuozzo and Kaiser, 1999). As redox potentials and GSH/GSSG ratios for the cytosolic compartment have generally been derived from whole-cell extracts, similar, albeit less extreme, sequestration may explain the difference between the redox potential we find in vivo and published data (Hwang et al., 1992; for review see Schafer and Buettner, 2001). Our results, which are very robust as discussed above, predict a significantly more reducing environment in the cytosol than previously estimated. In recent work, a GFP-based redox sensor has been applied to mitochondria and cytosol of mammalian cells in culture (Dooley et al., 2004; Hanson et al., 2004). However, it has not been determined what redox pair is measured in these cells nor has the kinetics for reaching equilibrium been investigated.
The engineered disulfide bond in rxYFP is of moderate stability as compared with many structural disulfide bonds in secretory proteins (Gilbert, 1995). Therefore, it is quite remarkable that it is found to be 10 and 53% oxidized under steady-state conditions in yeast and E. coli, respectively (Ostergaard et al., 2001). This suggests that the conspicuous absence of disulfide bonds in cytosolic proteins is a consequence of an evolutionary selection and not simply the result of reducing conditions.
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Materials and methods |
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Chromosomal deletions of GLR1 and GRX1 genes in M4975 (Table III) were generated by standard transformation with linear DNA fragments containing >300 bp of sequence derived from the regions immediately 5' and 3' to the start and stop codons, respectively, separated by an appropriate selection marker (Table III) from the YDp-series of plasmids (Sikorski and Hieter, 1989). The DNA fragments were constructed by PCR splicing by overlap extension (Horton et al., 1989).
For heterologous overexpression of Grx1p, the yeast GRX1 gene was amplified from genomic DNA using primers (5'-GGCGGCGCATATGGTATCTCAAGAAACTATC and 5'-GCCGCCCTCGAGATTTGCAAGAATAGGTTCTAAC) incorporating a 5' NdeI site overlapping the initiating Met codon and a 3' XhoI site. The cut amplicon was subsequently ligated into the corresponding sites of pET-24a(+) (Novagen) to give pHOJ167, expressing Grx1p with a COOH-terminal hexa-histidine tag.
Analytical techniques
The concentration of GSH (Sigma-Aldrich) in stock solutions was determined using 5,5'-dithiobis(2-nitrobenzoic acid) and 13,600 M1cm1 as the molar extinction coefficient () of nitrothiobenzoate at 412 nm (Ellman, 1959). Stock solutions of GSSG (Sigma-Aldrich) were quantified from its absorbance at 248 nm (
= 382 M1cm1; Chau and Nelson, 1991). Reduction of rxYFP and rGrx1p was performed by a 12-h incubation in the presence of 5 mM DTT at pH 7.0. Reductant was subsequently removed by gel filtration on a prepacked NAP-5 column (Sephadex G-25; Amersham Biosciences) equilibrated with an appropriate buffer. Before all in vitro experiments, buffers were thoroughly purged with argon to prevent interference from dissolved molecular oxygen.
Expression and purification of Grx1p
A 1-liter culture of Rosetta(DE3) (Novagen) cells harboring pHOJ167 was grown to the mid-log phase in terrific broth at 37°C (Sambrook et al., 1989), at which time IPTG was added to 1 mM. After 3 h of induction, cells were harvested, lysed, and rGrx1p purified on a His-Bind resin (Novagen) according to the manufacturer's protocol. To remove trace amounts of contaminating proteins, the elutate, dialyzed overnight against 20 mM potassium phosphate, pH 7.5, was subjected to anion exchange chromatography on a Resource Q column (Amersham Biosciences). rGrx1p, eluted with a linear gradient from 00.5 M NaCl in dialysis buffer, was >95% pure as judged by SDS-PAGE and Coomassie staining. The protein was stored at 80°C in 100 mM potassium phosphate, pH 7.0, 1 mM EDTA. Treatment with the thiol-specific alkylating reagent 4-acetamido-4'-maleimidylstilbene-2,2'-disulfonic acid (Molecular Probes) before and after incubation with DTT revealed that the purified rGrx1p was completely oxidized.
Rate and equilibrium constants for the reaction rxYFPred + GSSG rxYFPox + 2 GSH
Reactions were performed at 30°C in buffers containing 1 mM EDTA and either 100 mM K-MES (pH 6.16.7) or 100 mM K-MOPS (pH 7.07.9). The apparent second order rate constant (kox) for oxidation of rxYFP by GSSG was determined by diluting reduced rxYFP 300-fold into 2 ml of prewarmed buffer to a final concentration of
0.3 µM. When a stable base line was attained, the reaction was initiated by the addition of a high molar excess of GSSG to ensure pseudo first-order conditions, and the reaction was followed by the change in fluorescence at 523 nm. At each pH, a minimum of four independent measurements were performed at varying concentrations of GSSG. The rate constant kox was then obtained by a linear fit to the estimated pseudo first-order rate constants.
Equilibrium constants (Kox) at varying pH were determined directly in the fluorometer by incubating 0.2 µM reduced rxYFP in 2.5 ml MES or MOPS buffer (as above) containing 30150 µM GSSG and 19 mM GSH, adjusted to give 2070% oxidized rxYFP at equilibrium. To reduce the time required to reach equilibrium, reactions were performed in the presence of 5 µM rGrx1p. As shown in Fig. 4, rGrx1p functions as a simple redox-catalyst by speeding up the forward and reverse reactions to exactly the same extent. At equilibrium, the proportion of reduced redox sensor (fred) was determined by relating the fluorescence signal to that of the fully oxidized and reduced sensor obtained by addition of 20 µl 1.0 M GSSG to the reaction mixture followed by 20 µl 2.5 M DTT. To determine the exact amount of GSSG in the redox buffer at equilibrium, 180 µl of the reaction mixture were removed before addition of GSSG and DTT, quenched by addition of formic acid to 20%, and then analyzed by HPLC essentially as described previously (Takahashi and Creighton, 1996). At each pH, Kox was estimated from the relationship fred = 1/(1+ Kox·[GSSG]/[GSH]2) and reported as the mean of at least three independent reactions at varying [GSSG].
Equilibration of reduced rxYFP in a glutathione redox buffer
Progression of rxYFP toward equilibrium in a glutathione redox buffer can be described by the following differential equation
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Measurement of low molecular weight thiols
The intracellular content of GSH and GSSG was quantified by HPLC after derivatization with N-(1-pyrenyl)maleimide (NPM) using a protocol (Winters et al., 1995) optimized for yeast. Cells were grown in SC medium (when supplemented with either 1 mM GSH or GSSG, methionine was omitted from the medium) for more than five generations in the exponential phase before analysis. At an OD600 of 0.81.0, between 15 and 30 OD600 U were harvested by filtration onto a 25-mm glass filter membrane (Advantec) and immediately washed with 5 ml water. The filter was then rapidly transferred to 1 ml prewarmed 1% (wt/vol) 5-sulfosalicylic acid (Sigma; Anderson, 1985) and boiled for 4 min, interrupted by vigorous shaking every minute. The supernatant containing the extracted low molecular-weight thiols was separated from filter and cell debris by centrifugation at 15,000 g for 15 min (4°C) and kept on ice until further analysis. For determination of reduced thiols, 20 µl of the supernatant (appropriately diluted) were combined with 50 µl 100 mM Tris-HCl, pH 8.0, and 20 µl 0.1 M NaOH, raising the pH to 8, and immediately derivatized by addition of 750 µl 2.67 mM NPM (Sigma-Aldrich) dissolved in acetonitrile. After 10 min of incubation at room temperature, the reaction was quenched by addition of 2 µl 2 M HCl. To determine cellular disulfides, free thiols were blocked by adding 20 µl of the cell extract to a tube containing 50 µl 100 mM Tris-HCl, pH 8.0, 20 µl 0.1 M NaOH, 20 µl 10 mM NEM (Sigma-Aldrich), and 110 µM water. The mixture was incubated 10 min at 70°C. Excess NEM was subsequently quenched by addition of 20 µl 14.3 mM ß-mercaptoethanol (Sigma-Aldrich) followed by 10-min incubation at 70°C. Finally, disulfides were reduced by addition of 10 µl 20 mM tris-(2-carboxyethyl)phosphine (Sigma-Aldrich). After 10 min incubation at 70°C, the released thiols were labeled with NPM as described above. Quantification was performed by HPLC using a ReliaSil C18-AQ column (5 mm, 250 x 4.6 mm; Column Engineering Inc.). A 20-µl aliquot was injected onto the column and eluted isocractically in 65% acetonitrile, 0.1% acetic acid, 0.1% o-phosphoric acid (all vol/vol in water) at a flow rate of 0.5 ml/min. The NPM-derivatized thiols were detected by fluorescence with excitation and emission at 330 and 375 nm, respectively. Thiol concentrations were determined by relating the integrated peak areas to a standard curve based on known amounts of glutathione. Linearity was observed in the range from 300 fmol to 16 pmol thiol. The slope of the GSSG standard curve was found to be exactly twice that of the GSH standard curve, implying quantitative reduction of disulfides by tris-(2-carboxyethyl)phosphine. The reliability of the method was ascertained by the full recovery of exogenously added GSH or GSSG to cell extracts.
Determining the redox state of rxYFP in yeast by fluorescence
Strains were grown in SC-Leu medium (when supplemented with 1 mM GSH or GSSG, methionine was also omitted from the medium) for more than five generations before measurement. At an OD600 of 0.250.45, 840 µl of the culture was transferred to a prewarmed cuvette (30°C) and fluorescence monitored continuously using a Perkin-Elmer Luminescence Spectrometer LS50B equipped with an XF3074 emission filter (Omega Optical Inc.). Excitation wavelength was set at 512 nm (4-nm slit width). The pH of the medium was raised to 7 by addition of 60 µl 1.5 M K-MOPS, pH 7.9, 15 mM EDTA, immediately followed by 20 µl 4.6% (wt/vol) digitonin (Merck). After complete permeabilization of the cells (<3 min), observed as a slight drop in fluorescence intensity before return to a stable baseline (denoted Finit), the oxidation state of rxYFP was determined by reading the fluorescence after successive addition of 50 µl 6.3 mM 4-DPS (Sigma-Aldrich; Fox) and 100 µl 1 M DTT (Fred) to respectively fully oxidize and reduce the protein. The fraction of oxidized rxYFP was then calculated from the expression 1 (Finit Fox)/(Fred Fox), taking into account dilution by the added reagents.
Pulse labeling and immunoprecipitation
Before metabolic labeling, strains were grown overnight in SC-Leu-Met medium (when necessary, supplemented with 1 mM GSH or GSSG) to a final OD600 of 0.5 and then resuspended in the same medium at an OD600 of 0.5. After 30 min of preincubation at 30°C, cells were radiolabeled with 120 µCi [35S]Met/Cys (Pro-Mix; Amersham Biosciences) per ml of culture for 10 min and then chased by addition of excess methionine and cysteine. At indicated time points, 950 µl of cells were withdrawn and quenched by addition of 5% TCA (Sigma-Aldrich). Cells were spun down and washed twice with 0.5 ml ice-cold acetone. The pellet was allowed to dry and then redissolved in 100 µl of lysis buffer (100 mM K-MOPS, pH 7.0, 2% SDS, 1x complete protease inhibitor cocktail; Roche Diagnostics) containing 40 mM NEM. Cell lysis and immunoprecipitation were performed as described previously (Norgaard et al., 2001). rxYFP was precipitated using 10 µl polyclonal rabbit anti-GFP antibody (Molecular Probes) per time point.
Online supplemental material
Fig. S1 shows that yeast thioredoxins do not react with rxYFP. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200402120/DC1.
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Acknowledgments |
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Submitted: 20 February 2004
Accepted: 14 June 2004
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