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Journal of Bacteriology, January 2008, p. 487-493, Vol. 190, No. 2
0021-9193/08/$08.00+0     doi:10.1128/JB.01510-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

The GntR-Like Regulator TauR Activates Expression of Taurine Utilization Genes in Rhodobacter capsulatus

Jessica Wiethaus, Britta Schubert, Yvonne Pfänder, Franz Narberhaus, and Bernd Masepohl^*

Lehrstuhl für Biologie der Mikroorganismen, Fakultät für Biologie, Ruhr-Universität Bochum, 44780 Bochum, Germany

Received 19 September 2007/ Accepted 25 October 2007

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Rhodobacter capsulatus can efficiently grow with taurine as the sole sulfur source. The products of the tpa-tauR-xsc gene region are essential for this activity. TauR, a MocR-like member of the GntR superfamily of transcriptional regulators, activates tpa transcription, as shown by analysis of wild-type and tauR mutant strains carrying a tpa-lacZ reporter fusion. Activation of the tpa promoter requires taurine but is not inhibited by sulfate, which is the preferred sulfur source. TauR directly binds to the tpa promoter, as demonstrated by DNA mobility shift assays. As expected for a transcriptional activator, the TauR binding site is located upstream of the transcription start site, which has been determined by primer extension. Site-directed promoter mutations reveal that TauR binds to direct repeats, an unusual property that has to date been shown for only one other member of the MocR subfamily, namely, GabR from Bacillus subtilis. In contrast, all other members of the GntR family analyzed so far bind to inverted repeats.

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Sulfate is the preferred sulfur source of most bacteria, but in most natural environments, sulfate availability is limited. Sulfonates and sulfate esters are much more abundant than sulfate, and consequently, many bacteria have adapted mechanisms to use these compounds as alternative sulfur sources (10). Upon sulfate limitation, Escherichia coli and many other bacteria synthesize a set of sulfate starvation-induced proteins (references 10, 24, and 25 and references therein). Among these are ABC-type transporters involved in uptake of taurine (2-aminoethanesulfonate).

Phototrophic purple bacteria (Rhodospirillaceae), and especially Rhodobacter species, have long been used as model organisms to study anoxygenic photosynthesis, carbon assimilation, hydrogen metabolism, and nitrogen fixation (references 6-8 and 15 and references therein). In the absence of sulfate, Rhodobacter capsulatus can efficiently grow with taurine under photoheterotrophic conditions (16).

Two divergently transcribed gene clusters, tauABC and tpa-tauR-xsc (formerly orf459-orf484-orf590), are involved in taurine utilization by Rhodobacter capsulatus (16) (Fig. 1). The tauABC genes are predicted to encode an ABC transport system mediating taurine uptake. The tpa and xsc gene products exhibit strong similarity to taurine-pyruvate aminotransferase (Tpa) from Bilophila wadsworthia (catalyzing the initial transamination of taurine to 2-sulfoacetaldehyde during anaerobic taurine degradation) (13) and sulfoacetaldehyde acetyltransferase (Xsc) from Alcaligenes defragrans (converting 2-sulfoacetaldehyde into sulfite and acetyl phosphate) (21). Genes similar to R. capsulatus tauR were previously identified in close proximity to xsc genes in the genomes of Paracoccus denitrificans and other proteobacteria by database searches (4, 21), but none of these genes has been experimentally characterized so far. As shown in this study, TauR is a transcriptional activator essential for tpa expression.

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FIG. 1. Organization of the R. capsulatus tpa-tauR-xsc gene region. Mutant strains defective for tpa, tauR, and xsc contain Gm resistance cassettes, with the directions of transcription of the Gm resistance gene symbolized by arrows. Hybrid plasmid pBSRUB60 carrying a transcriptional fusion of lacZ to the tpa promoter is based on the mobilizable broad-host-range plasmid pBBR1MCS. Neither the Gm resistance cassette nor the lacZ gene is drawn to scale.

R. capsulatus TauR belongs to the GntR superfamily of bacterial transcription regulators that bind DNA by a helix-turn-helix motif. The N-terminal DNA-binding domain is well conserved for all members of the GntR family, whereas the C-terminal effector-binding and oligomerization domain is more variable and therefore was used to define six GntR subfamilies, namely, the FadR, HutC, MocR, YtrA, AraR, and PlmA families (14, 19). Members of the GntR family control diverse processes, such as fatty acid metabolism in E. coli (FadR), histidine utilization in Pseudomonas putida (HutC), rhizopine catabolism in Rhizobium meliloti (MocR), acetoin utilization in Bacillus subtilis (YtrA), arabinose metabolism in B. subtilis (AraR), and plasmid copy number control in Anabaena sp. strain PCC 7120 (PlmA). R. capsulatus TauR shows the highest similarity to the MocR subfamily, named after S. meliloti MocR (20).

In contrast to well-characterized members of the FadR, HutC, and YtrA subfamilies, which bind to inverted repeats in the promoter regions of their target genes, MocR-like proteins had previously been predicted to bind to direct repeats (19). To date, this has been shown for only one MocR-like protein, namely, GabR from B. subtilis (2, 3). GabR acts as both a negative autoregulator and an activator of genes involved in the utilization of -aminobutyrate (GABA) as the sole nitrogen source.

In this study, we analyzed the role of R. capsulatus TauR by genetic and biochemical means. TauR was shown to be essential for taurine-dependent tpa expression. Site-directed mutagenesis of the tpa promoter and DNA mobility shift assays suggest binding of TauR to direct repeats.

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Strains, plasmids, and growth conditions. The bacterial strains and plasmids used in this study are listed in Table 1. Methods for conjugational plasmid transfer between E. coli and R. capsulatus, procedures for selection of mutants, and growth conditions have been described earlier (reference 16 and references therein).

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TABLE 1. Bacterial strains and plasmids used in this study

RNA isolation and primer extension. R. capsulatus cultures were grown in RCV minimal medium with taurine as a sulfur source until late logarithmic phase. RNAs of these cultures were isolated using the Micro-to-Midi Total RNA Purification System (Invitrogen, Karlsruhe, Germany) following the instructions of the manufacturer. Primer extension was carried out as described previously (1) using oligonucleotide LP-tpa-2, complementary to the 5'end of tpa mRNA (Table 2), to map the transcription start site of tpa (Fig. 2).

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TABLE 2. Synthetic oligonucleotides used in this study

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FIG. 2. Transcription start site mapping of the tpa promoter. Primer extension was carried out with total RNA from R. capsulatus cells grown with taurine as the sole sulfur source. Primer LP-tpa-2 (binding to the 5' region of tpa) (Table 2) was used for reverse transcription. The resulting primer extension product is shown in lane PE. The corresponding sequencing reactions (A, C, G, and T) with plasmid pBSRUB48 served as length standards. The putative (–10) promoter region, the transcription start site (+1), the ribosomal binding site (RBS), and the translation initiation codon for the tpa gene product (Tpa) are indicated.

Construction of the P_tpa-lacZ reporter plasmid pBSRUB60 and β-galactosidase assays. A 461-bp DNA fragment (Fig. 3, fragment A) encompassing the tpa promoter (P_tpa) was PCR amplified with the primer pair UP-tpa-1/LP-tpa-2 (Table 2), using R. capsulatus chromosomal DNA as a template. The PCR product was blunt-end cloned into the SmaI site of vector plasmid pBluescript KS, leading to plasmid pBSRUB48, in which P_tpa was flanked by BamHI and HindIII restriction sites. Subsequently, the BamHI-HindIII fragment from pBSRUB48 carrying P_tpa was cloned into the mobilizable broad-host-range plasmid pBBR1MCS. Finally, the promoterless E. coli lacZ gene was inserted at the HindIII site, resulting in reporter plasmid pBSRUB60 (tpa-lacZ). R. capsulatus reporter strains carrying pBSRUB60 (tpa-lacZ) were grown in RCV minimal medium with taurine or sulfate as a sulfur source before β-galactosidase activities were determined as described earlier (16).

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FIG. 3. DNA mobility shift assays narrowing down the TauR binding site within the tpa promoter region. The transcription start site of the tpa gene (labeled +1) is indicated above the physical map. DNA fragment A (461 bp) encompassing the tauA-tpa intergenic region was generated by PCR amplification using the primer pair UP-tpa-1/LP-tpa-2 (Table 2). In addition, subfragments were generated by cutting fragment A with the indicated restriction enzymes. The DNA fragments were either preincubated with purified TauStrep protein (lanes 2, 4, 6, 8, and 10) or not (lanes 1, 3, 5, 7, and 9) prior to agarose gel electrophoresis. DNA fragments retarded by TauR are marked (A to E). A 50-bp DNA ladder (Fermentas, St. Leon-Rot, Germany) was used as a length standard (lane M).

Overexpression of R. capsulatus TauR protein in E. coli. The tauR coding region was PCR amplified with the primer pair UP-tauR/LP-tauR (encompassing recognition sites for SacII and SalI, respectively) (Table 2) using R. capsulatus chromosomal DNA as a template. Subsequently, the 1.5-kb SacII-SalI DNA fragment with tauR was cloned into the Strep-tag expression vector pASK-IBA3 (Table 1), resulting in hybrid plasmid pBSRUB29, carrying an in-frame tauR-Strep-TagII fusion (tauR_Strep). Plasmid pBSRUB29 was transformed into E. coli strain BL21(DE3), which served as a host for overexpression of the tagged R. capsulatus TauR protein (TauR_Strep). Purification of the recombinant protein was carried out as described previously (26).

Nonradioactive DNA mobility shift assays. A 461-bp DNA fragment encompassing P_tpa was obtained by PCR amplification with the primer pair UP-tpa-1/LP-tpa-2 using plasmid pBSRUB48 as a template (Table 2). P_tpa subfragments were generated by cutting the 461-bp amplification product with appropriate restriction enzymes (Fig. 3). Preincubation of promoter fragments with the TauR_Strep protein and agarose gel electrophoresis of DNA-protein complexes were carried out as described previously (18).

Site-directed mutagenesis and radioactive DNA mobility shift assays. Point mutations within the tpa promoter region (designated mut-1 to mut-4) (Fig. 4) were generated by overlap extension PCR (23) with appropriate primers (Table 2) and pBSRUB48 as a template. The PCR products were blunt-end cloned into the SmaI site of pBluescript KS, leading to hybrid plasmids pBSRUB94 (mut-1), pBSRUB95 (mut-2), pBSRUB96 (mut-3), pBSRUB97 (mut-4), and pBSRUB110 (mut-3/4). These plasmids served as templates for PCR amplification of 269-bp mutant tpa promoter fragments using the primer pair PJW72-U/PJW73-L (Table 2). Purification of the amplification products, ³²P end labeling, preincubation of labeled promoter DNA with the TauR_Strep protein, and polyacrylamide gel electrophoresis of DNA-protein complexes were carried out as described earlier (27). If required, 1 µM taurine or 100 µM pyridoxal 5'-phosphate was added to the preincubation mixture. Quantification of DNA mobility shift assays was done using the AIDA image analyzer software, version 4.19 (Raytest, Straubenhardt, Germany).

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FIG. 4. Site-directed mutagenesis of the tpa promoter. The heavy line below the physical map depicts a 269-bp PCR fragment (amplified with the primer pair PJW72-U/PJW73-L [Table 2]) used for DNA mobility shift assays (Fig. 5). Site-directed mutations (mut-1, mut-2, mut-3, and mut-4) within direct repeats (DR-1a, DR-1b, DR-2a, and DR-2b) were created by overlap extension PCR. Direct repeats DR-2a and DR-2b are centered at positions –72.5 and –49.5, respectively, relative to the transcription start site (+1).

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Mapping of the tpa transcription start site. Genetic analyses have indicated that the tpa, tauR, and xsc genes comprise a single transcription unit (reference 16 and this study) (Fig. 1). To assign the transcription start site of the tpa-tauR-xsc operon, primer extension experiments were performed with total RNA isolated from an R. capsulatus wild-type culture grown with taurine as the sole sulfur source (see Materials and Methods) (Fig. 2). The 5' end of the mRNA mapped between the TauR binding site (see below) (Fig. 3) and the ATG start codon for the tpa gene.

TauR activates tpa expression. To analyze the expression of the tpa-tauR-xsc operon, plasmid pBSRUB60 carrying a transcriptional tpa-lacZ fusion was introduced into wild-type R. capsulatus and tpa, tauR, and xsc mutant strains (see Materials and Methods) (Table 3). These mutant strains contain a gentamicin (Gm) resistance cassette, which constitutively drives transcription of downstream genes (reference 27 and references therein). The resulting reporter strains were grown in RCV minimal medium with taurine or sulfate as a sulfur source prior to determination of lacZ-mediated β-galactosidase activity, as described earlier (16). The results shown in Table 3 can be summarized as follows. (i) Expression of the tpa-lacZ fusion was taurine inducible, as expected from earlier studies (16). In contrast to R. capsulatus, the tpa and xsc genes in Rhodobacter sphaeroides belong to different operons. While Xsc and other enzymes involved in the utilization of taurine as a sole carbon or nitrogen source by R. sphaeroides are synthesized upon the addition of taurine, expression of the tpa gene is not induced by taurine (5). Thus, even closely related species, such as R. capsulatus and R. sphaeroides, have adopted various strategies for taurine utilization. (ii) Transcription from the tpa promoter was not inhibited by sulfate. In contrast, expression of the tauABC genes coding for a putative ABC-type taurine transporter was clearly repressed by sulfate (16). Nevertheless, small amounts of the TauABC transporter should be synthesized in the presence of sulfate in order to allow uptake of small amounts of taurine sufficient for TauR-dependent tpa activation. Alternatively, some taurine might enter the cells independently of the sulfate-inhibited TauABC transporter. However, utilization of taurine as a sulfur source seems to be primarily controlled by sulfate repression of taurine uptake, while utilization of (intracellular) taurine is not inhibited by sulfate. This might reflect the adaptation of R. capsulatus to natural environments, which are characterized by much lower and more varied concentrations of sulfate and taurine than the experimental conditions used in this study. (iii) Taurine-dependent induction of tpa expression strictly required TauR. (iv) In contrast to TauR, neither Tpa nor Xsc was essential for tpa expression. It remains unclear, however, why the xsc mutation led to elevated levels of tpa-lacZ expression in the presence of taurine plus sulfate. (v) A polar tpa mutation (with the Gm gene reading in the opposite direction from tpa; FFRUB47) prevented tpa-lacZ expression, suggesting that tauR expression derives from the tpa promoter. Thus, this finding implies cotranscription of tpa and tauR. (vi) Expression of tauR by the constitutive Gm gene promoter (FFRUB48) led to elevated levels of tpa-lacZ expression, indicating that tauR was overexpressed in the mutant. In this mutant, clear tpa expression occurred in the presence of sulfate as the sole sulfur source, suggesting that TauR mediates tpa activation to some extent independently of taurine availability. In the wild type, taurine-independent low-level expression of the tpa-tauR-xsc operon may account for basal levels of TauR required for positive autoregulation.

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TABLE 3. Taurine-dependent expression of tpa-lacZ in R. capsulatus

TauR binds to the tpa promoter. Since R. capsulatus TauR was essential for tpa-lacZ expression (Table 3), it seemed likely that TauR would bind to the tpa promoter. Interaction of TauR with the tpa promoter was analyzed by DNA mobility shift assays (see Materials and Methods). For this purpose, the TauR protein was overexpressed and purified as a C-terminally Strep-tagged recombinant protein from E. coli (see Materials and Methods). TauR_Strep was preincubated either with a 461-bp PCR fragment encompassing the tpa promoter or with subfragments created by digestion of the PCR product with different restriction enzymes. The protein-DNA complexes were analyzed by agarose gel electrophoresis, followed by ethidium bromide staining (Fig. 3). Since only one of two restriction fragments in the respective assays was retarded, the other fragment served as an internal control for TauR-specific binding. Comparison of retarded DNA fragments suggested that the TauR binding site is confined to an 88-bp BglI-TacI fragment within the tpa promoter region. As one would expect for the binding site of a transcriptional activator, the TauR binding site is located upstream of the transcription start site (Fig. 3).

Since tpa-lacZ expression was taurine inducible in vivo (Table 3), we asked whether taurine might influence in vitro binding of TauR to the tpa promoter. The tpa promoter region was PCR amplified using the primer pair PJW72-U/PJW73-L (Table 2) prior to radioactive 5' end labeling of the 269-bp PCR product (Fig. 4). TauR_Strep was preincubated with the radioactively labeled tpa promoter fragment in the presence or absence of taurine. Analysis of protein-DNA complexes by polyacrylamide gel electrophoresis revealed specific binding of TauR to the tpa promoter in the absence of taurine (Fig. 5A and B). TauR binding was not further enhanced by taurine. Thus, taurine-independent binding of the regulator to DNA and taurine-dependent gene activation are two separate events (see "Conclusions" below).

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FIG. 5. Influences of taurine and PLP on binding of TauR to the tpa promoter and specificity controls. (A) A 269-bp DNA fragment (marked in Fig. 4) carrying the TauR binding site was 32P end labeled prior to incubation with increasing amounts of TauRStrep (0, 0.02, 0.04, 0.07, 0.15, 0.22, 0.29, 0.44, 0.58, and 1.17 µM) in the absence or presence of taurine and/or PLP (see Materials and Methods). All binding assays were performed in the presence of poly(dI-dC) as the competitor DNA. (B) The decrease of band intensities for free probes, as shown in panel A, was quantified using the AIDA image analyzer software (Raytest, Straubenhardt, Germany). The band intensities of the free probes in the absence of TauR were set as 100%. (C) Unlabeled wild-type tpa promoter fragments (P-wt) were used as specific competitor DNA, while mutant tpa promoter fragments (P-mut-3/4, carrying the mutations mut-3 and mut-4) (Fig. 4) served as nonspecific competitor DNA. In detail, 0.58 µM TauR and radioactively labeled wild-type tpa promoter fragments were mixed with an 80-, 160-, 320-, 640-, 1,280-, 2,560-, 5,120-, or 10,240-fold excess of unlabeled competitor or noncompetitor DNA. All reactions were performed with 0.6 fmol 32P-labeled DNA.

Members of the MocR subfamily of transcriptional regulators contain a class I aminotransferase domain binding pyridoxal 5'-phosphate (PLP) (19). Since the putative PLP binding site is highly conserved in TauR (data not shown), we analyzed TauR-mediated gel retardartion of the tpa promoter in the presence and absence of PLP (Fig. 5A and B). As described above for taurine, PLP did not enhance in vitro binding of TauR to the tpa promoter (see "Conclusions" below).

Binding of TauR to the radioactively labeled tpa promoter fragment could be reversed by addition of increasing amounts of unlabeled tpa promoter DNA, but not by mutant tpa promoter DNA (Fig. 5C) (see below), thus demonstrating TauR binding specificity.

TauR binds to direct repeats. The 88-bp BglI-TacI fragment (encompassing the TauR binding site) contains two pairs of almost perfect direct repeats (DR-1a/DR-1b and DR-2a/DR-2b) (Fig. 4). To examine the roles of these sequences in TauR binding, site-directed mutations were created by overlap extension PCR (see Materials and Methods) (Fig. 4). As described above for the wild-type tpa promoter, DNA fragments carrying the mutations mut-1 to mut-4 were PCR amplified using the primer pair PJW72-U/PJW73-L (Table 2), radioactively labeled, and incubated with TauR_Strep.

The results of DNA mobility shift assays with increasing amounts of TauR_Strep are shown in Fig. 6. Binding of TauR to the tpa promoter was not influenced by the mutations mut-1 and mut-2 compared to the wild-type promoter. These findings suggest that the direct repeats DR-1a and DR-1b are dispensable for TauR binding. In contrast, binding of TauR was clearly diminished by the mutations mut-3 (in DR-2a) and mut-4 (in DR-2b). Interestingly, the effect of mut-4 was less pronounced than that of mut-3. Since the direct repeats DR-2a and DR-2b differ by 1 base pair (Fig. 4), one may speculate that TauR binds more strongly to DR-2a than to DR-2b. Nonetheless, a combination of both mutations (mut-3/4) completely abolished binding of TauR to the tpa promoter, strongly suggesting that both direct repeats, DR-2a and DR-2b, are involved in TauR binding.

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FIG. 6. DNA mobility shift assays with TauR and mutant tpa promoter fragments. (A) PCR fragments (269-bp) (marked in Fig. 4) carrying either the wild-type tpa promoter or variants mut-1, mut-2, mut-3, mut-4, and mut-3/4 were incubated with increasing amounts of TauRStrep (up to 1.17 µM). (B) The decrease of band intensities for free probes, as shown in panel A, was quantified using the AIDA image analyzer software (Raytest, Straubenhardt, Germany).

Conclusions. R. capsulatus TauR was shown to be essential for the activation of tpa expression. Since tauR is cotranscribed with the tpa gene, TauR appears to be a positive autoregulator. Interestingly, in vivo tpa expression was clearly enhanced by taurine, but taurine did not influence the binding of TauR to the tpa promoter in vitro. An unknown coactivator protein responding to taurine availability and binding to the direct repeats DR-1a/DR-1b immediately upstream of the TauR binding site might be required for responsiveness to taurine.

Alternatively, upon taurine-independent binding of TauR to the tpa promoter, TauR might be converted into its transcription-competent form by a reaction between taurine and PLP via the aminotransferase domain of TauR. Like other aminotransferases, TauR might transaminate taurine through conversion of PLP to pyridoxamine 5'-phosphate (PMP). In turn, TauR (carrying PMP) might activate tpa transcription. It remains speculative whether conversion of PLP to PMP requires pyruvate or another keto acid as an amino acid acceptor or if amination of PLP in TauR occurs independently of a keto acid. A similar model has been discussed previously for B. subtilis GabR, which binds to its target promoter independently of GABA and PLP but requires both GABA and PLP for activation of its target genes (2).

Binding of TauR to the tpa promoter involves direct repeats spanning the DNA region between –77 and –45 upstream of the transcription start site. Due to the close proximity between the TauR binding site and the putative –35/–10 promoter, one would expect direct interaction between TauR and RNA polymerase without the necessity of DNA bending.

Like TauR, GabR binds to direct repeats (2). However, the binding sites of the two regulators have different sequences and distances between the repeats. While the centers of the GabR binding sites (ATACCA) are separated by 34 bp (equivalent to three helical turns), the centers of the TauR binding sites (CTGGAC[T/C]TAA) are separated by 23 bp (equivalent to two helical turns). Thus, in both cases, the repeats are located on the same face of the DNA.

Binding of transcriptional regulators to direct repeats is an exception rather than the rule. To our knowledge, GabR from the gram-positive bacterium B. subtilis and TauR from the gram-negative bacterium R. capsulatus are the only MocR-like regulators that have been experimentally shown to bind to direct repeats. Thus, more work on this widespread but largely uncharacterized family of regulators is needed.

 
* Corresponding author. Mailing address: Lehrstuhl für Biologie der Mikroorganismen, Fakultät für Biologie, Ruhr-Universität Bochum, 44780 Bochum, Germany. Phone: 49 (0) 234 32 25632. Fax: 49 (0) 234 32 14620. E-mail: bernd.masepohl{at}rub.de

Published ahead of print on 2 November 2007.

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Journal of Bacteriology, January 2008, p. 487-493, Vol. 190, No. 2
0021-9193/08/$08.00+0     doi:10.1128/JB.01510-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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