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1 Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield S10 2TN, UK
2 The School of Biosciences, The University of Birmingham, Birmingham B15 2TT, UK
Correspondence
Jeffrey Green
jeff.green{at}sheffield.ac.uk
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ABSTRACT |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Unden et al., 2002). This metabolic hierarchy reflects the amount of energy that can be conserved by each process, and is maintained by oxygen-, nitrate- and nitrite-responsive transcription factors. The E. coli FNR protein is an oxygen-responsive global transcription factor (Kiley & Beinert, 2003). Under anaerobic conditions FNR contains a [4Fe–4S] cluster that promotes dimerization of the protein and enhances site-specific DNA binding (Jervis & Green, 2007; Lazazzera et al., 1996). In the presence of oxygen, disassembly of the FNR iron–sulphur clusters is accompanied by the formation of FNR monomers and inhibition of DNA binding, thereby switching off FNR-activated genes (Kiley & Beinert, 2003; Lazazzera et al., 1996; Sutton et al., 2004). The DNA sequence recognized by FNR consists of an inverted repeat with a consensus sequence of TTGATnnnnATCAA (Eiglmeier et al., 1989). Once bound to DNA, FNR dimers activate transcription by recruiting RNA polymerase, or alternatively repress transcription by inhibiting the formation of productive promoter–RNA polymerase interactions (reviewed by Browning et al., 2003). There are two basic types of FNR-dependent promoter. Class I promoters possess FNR sites located close to –61.5, –71.5, –82.5 or –92.5 (Wing et al., 1995). This architecture permits the formation of only one protein–protein contact, between FNR and the C-terminal domain of the RNA polymerase 
-subunit (Browning et al., 2003). However, the more common architecture for FNR-activated promoters is that adopted by class II promoters, which have an FNR site centred at or close to 41.5 bp upstream of the transcript start site. This arrangement allows the formation of multiple protein–protein contacts between RNA polymerase and FNR (Browning et al., 2003).
As mentioned above, in the absence of oxygen, nitrate is the preferred electron acceptor (Guest et al., 1996). In E. coli the presence of nitrate is sensed by paralogous two-component systems NarXL and NarPQ (Stewart, 1994). The membrane-bound sensors NarX and NarP perceive the presence of nitrate and nitrite in the environment (Lee et al., 1999; Rabin & Stewart, 1992, 1993; Williams & Stewart, 1997). These sensors then act to control the phosphorylation state of the regulators NarL and NarP (Darwin & Stewart, 1996; Yamamoto et al., 2005), and there is a complex exchange of information between the sensors and the regulators to allow fine control of target-gene expression in response to nitrate and nitrite. The phosphorylated regulators bind specific DNA sequences related to the consensus TACYYMT (where Y=C or T, and M=A or C) to positively or negatively regulate gene expression (Darwin et al., 1997). In addition, there is differential recognition of NarL/P heptamer patterning, such that NarL recognizes all TACYYMT heptamers, whereas NarP only binds these sequences when they are arranged as an inverted repeat with a 2 bp spacing, the so-called 7-2-7 arrangement (Darwin et al., 1997). Thus, the Nar two-component systems work in conjunction with FNR to maintain a central metabolic hierarchy in E. coli. For example, in the absence of oxygen, nitrate promotes the NarL-mediated activation of the FNR-dependent nitrate reductase operon (narGHJI) and NarL-mediated repression of the FNR-dependent fumarate reductase operon (frdABCD) (Darwin & Stewart, 1996).
Upstream of the E. coli ydhY gene is a consensus FNR site. Transcript-profiling experiments have shown that the abundance of the ydhY and downstream transcripts is enhanced under anaerobic conditions in the presence of FNR (Constantinidou et al., 2006; Kang et al., 2005), and that the transcripts are less abundant in the presence of nitrate (Constantinidou et al., 2006). In vivo transcription studies using a ydhY–lacZ fusion have shown that ydhY expression is FNR-dependent, and in vitro transcription reactions in the presence of FNR yield a product with a size consistent with the presence of a class II FNR-dependent promoter upstream of ydhY (Kang et al., 2005).
The amino acid sequences of the proteins encoded by the ydhY–T operon suggest that they are components of an oxidoreductase. Thus, YdhX (222 aa) is a putative electron transfer protein with a Tat signal sequence (MSFTRRKFVLGMGTVIFFTGSASSLLA) (Berks et al., 2005), four [4Fe–4S] clusters and 50 % identity over 213 aa to the nitrite reductase protein NrfC from E. coli. The YdhV protein (700 aa) possesses DXXGL motifs that are associated with archaeal tungsten-containing oxidoreductases and is 28 % identical (41 % similar over 687 aa) to the aldehyde ferredoxin oxidoreductase of Pyrococcus furiosus (Kletzin et al., 1995). This protein lacks a Tat signal sequence and thus should be located in the cytoplasm, unless it is exported as a complex with YdhX. YdhU (261 aa) is a predicted cytochrome b-containing integral membrane protein that is 43 % identical to the thiosulphate reductase cytochrome b subunit of Salmonella typhimurium (Heinzinger et al., 1995). For Tat substrates such as YdhX, the assembly of the catalytic complex requires accessory proteins known as redox enzyme maturation proteins (REMPs), which are often co-transcribed with the genes encoding the oxidoreductase subunits (Turner et al., 2004). It is possible that the YdhW and YdhT proteins are REMPs. The YdhY protein (208 aa) is predicted to be ferredoxin-like in possessing four iron–sulphur clusters. Thus, it is possible that YdhY is involved in electron transfer reactions with YdhV.
In this study, it is shown that the ydhY gene is the first gene of a six-gene operon (ydhYVWXUT). The mechanism of regulation of ydhY–T expression by FNR, in response to oxygen availability, and by the NarXL and NarPQ systems, in response to nitrate and nitrite availability, revealed that NarL represses ydhY–T expression by binding to sites in the promoter region that overlap those occupied by FNR and RNA polymerase. Furthermore, it is shown that NarP recognizes a 7-2-7 site close to the transcript start to repress ydhY–T expression in the presence of nitrate and nitrite, providing new evidence in support of earlier transcript-profiling experiments that suggest that nitrate-activated NarP can act as a repressor (Constantinidou et al., 2006).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. Cultures of E. coli were grown in L broth (Lennox, 1955) or Evans defined minimal medium (Evans et al., 1970) supplemented with appropriate antibiotics (ampicillin, 100 µg ml–1; chloramphenicol, 20 µg ml–1; tetracycline, 25 µg ml–1). For β-galactosidase assays, L broth containing appropriate antibiotics was inoculated (1 : 50) from overnight cultures and incubated under aerobic (250 ml conical flasks containing 5 ml medium shaken at 250 r.p.m.) or anaerobic (sealed bottles filled to the neck) conditions at 37 °C until OD600 0.4–0.6 was reached. β-Galactosidase activities were then measured (Miller, 1972).
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). The ydhY promoter region (–189 to +176 relative to the transcription start) was amplified from the chromosome of E. coli MC1000 by PCR using appropriate oligonucleotides designed to contain EcoRI/BamHI restriction sites. The authenticity of the amplified fragment was verified by DNA sequencing after ligation into EcoRI/BamHI-digested pRS415 to create pGS1739 (Simons et al., 1987). The inserts were then transferred to 
RZ5 and the resulting (promoter–lacZ) fusions were introduced as single copies into the attachment site of isogenic E. coli strains (Simons et al., 1987). Lysogeny status was verified using the method of Powell et al. (1994). Site-directed mutagenesis of the FNR site (TTGATAACGATCAA to ATCATAACGATGAT; mutated bases shown in bold type) was achieved using PCR and appropriate synthetic oligonucleotides. Mutagenesis of the four possible NarL/P sites (heptamer –42, TAACGAT to TTAAAAT; heptamer –16, TAATCAC to ATAAATC; heptamer +6, TAACCAT to TTAAGGT; and heptamer +15, AGGATTA to TCAATTT, mutated bases shown in bold type) was also achieved using PCR. DNA sequencing after ligation into pRS415 verified the authenticity of the altered promoters before transfer to the E. coli chromosome as single-copy fusions.
For 5' random amplification of cDNA ends (RACE)-PCR transcript mapping, RNA was isolated from anaerobic cultures of E. coli MC1000 using Qiagen RNeasy mini kits according to the manufacturer's instructions. The transcript start was identified using 2 µg E. coli RNA per RACE reaction according to the manufacturer's instructions (Roche). The initial cDNA template was generated using oligonucleotide primer RydhY (TCTAATAGTGGAGGATCCACCGGGTTCATT). The cDNA was polyadenylated and then amplified using RydhY and the oligo(dT) anchor primer (Roche), yielding a single product of 
400 bp (as judged by agarose gel electrophoresis), which was subjected to DNA sequencing.
The extent of the ydhY operon was determined using the Reverse-iT One-Step RT-PCR kit (ABgene) and total RNA prepared as described above. The oligonucleotides used were: TATACCGCGGACACCTGCCG (forward primer ydhY); TGGCAGCACACAATCTACTG (forward primer ydhV); GCATCTGTATCGCCGGTTCG (forward primer ydhX); ACCGCATAACACACATTTCG (reverse primer ydhY); CTCTTAATATATTACCTGTC (reverse primer ydhV); CGAGAGAAACACGGCGCGCG (reverse primer ydhW); CCATCCCCAGAACAAATTTG (reverse primer ydhX); GCCACGAGCACGAAGCAAGG (reverse primer ydhU); CTCCGATACGCCATTCTCGC (reverse primer ydhT). Control reactions lacking reverse transcriptase used the ydhY forward primer and the ydhY reverse primer.
Gel retardation and footprinting assays.
Radiolabelled ydhY promoter fragments (–189 to +176 relative to the transcript start) were prepared by PCRs (50 µl total volume) with the same primers used to create the ydhY–lacZ fusion in the presence of [-32P]dATP (0.37 MBq). The amplified product was purified from Tris-acetate-buffered agarose gels (Sambrook & Russell, 2001) using a Qiagen gel extraction kit (Qiagen). The purified labelled promoter DNA was incubated with 8 µM FNR-D154A (FNR*), an FNR protein that retains the ability to bind DNA under aerobic conditions (Ziegelhoffer & Kiley, 1995), together with Tris-HCl, pH 8.0 (20 mM), glycerol (5 %, v/v), KCl (100 mM), BSA (0.1 mg ml–1), DTT (1 mM) and calf thymus DNA (3 µg) for 5 min, before separating the FNR* : DNA complexes from DNA by electrophoresis in polyacrylamide gels buffered with Tris/borate/EDTA (TBE) buffer (90 mM Tris, 90 mM borate, 2 mM EDTA). After electrophoresis the gels were transferred to filter paper (3MM, Whatman) and dried for autoradiography. For gel retardation assays with maltose binding protein (MBP)–NarL (0–4 µM) and MBP–NarP (0–4 µM), the Nar proteins were phosphorylated by incubation with acetyl phosphate (50 mM) for 45 min at 25 °C. The proteins were then incubated with radiolabelled ydhY promoter DNA (as above) in binding buffer: HEPES, pH 8.0 (20 mM), MgCl2 (5 mM), potassium glutamate (50 mM), DTT (1 mM) and calf thymus DNA (3 µg), for 15 min before separation of protein–DNA complexes on 6 % polyacrylamide gels buffered with TBE. After electrophoresis the gels were transferred to filter paper (3MM, Whatman) and dried for autoradiography. Footprinting reactions with FNR* (2 µM), phosphorylated MBP–NarL (0–6 µM) and phosphorylated MBP–NarP (0–6 µM) were done essentially as described by Darwin et al. (1997) with PydhY (–189 to +176) used to construct pGS1739 as the target DNA (see above). The promoter fragment was radiolabelled using Klenow fragment and an appropriate [32P]dNTP at either the BamHI site (NarP) or the EcoRI site (NarL). The MBP–NarL and MBP–NarP proteins were purified as previously described (Darwin et al., 1997).
Construction of a ydhY–T mutant.
A disruption in the ydhYVWXUT operon of W3110 was obtained by linear transformation based on the method of Yu et al. (2000). Oligonucleotides containing 3' sequences complementary to the first or last 20 bp of the chloramphenicol-resistance cassette of plasmid pACYC184 (Martinez et al., 1988) and 5'-end sequences flanking ydhY–T were constructed. Linear DNA carrying the resistance cassette and flanking regions was generated by PCR. E. coli strain W3110 containing the plasmid pTP223 (TetR) (Poteete & Fenton, 1984), which carries the red recombinase genes under the control of an IPTG-inducible promoter, were grown overnight at 37 °C and diluted (1 : 100) in L broth containing tetracycline (25 µg ml–1) and IPTG (2 mM) and grown to OD600 0.3. Electrocompetent cells were prepared and transformed with 5 µg PCR product then incubated in 1 ml L broth at 37 °C for 1 h before plating on selective medium. The resulting colonies (CmR) were immediately cured of pTP223 and the presence of the mutation was confirmed by PCR and DNA sequencing. Further transfer of the ydhY mutation into clean genetic backgrounds was achieved using bacteriophage P1vir-mediated transduction (Sambrook & Russell, 2001).
Biolog Phenotype Microarray (PM) studies.
Cultures of E. coli W3110 and the isogenic ydhY mutant (JRG5199) were grown for 16 h at 37 °C on R2A agar (Biolog) under anaerobic conditions. Colonies were picked from the plate and suspended in 15 ml IF-0 inoculating fluid (Biolog) to a density that matched the Biolog 85 % turbidity standard (OD420 0.12). The Biolog PM1 and PM2 phenotyping arrays, which contain different carbon sources, were directly inoculated with 100 µl bacterial suspension per well. For PM3 and PM4 (nitrogen, phosphorus and sulphur sources) the bacterial suspensions were supplemented with glycerol (20 mM) and ferric citrate (2 µM). All arrays were incubated anaerobically at 37 °C in a Labsystems iEMS Reader and were monitored hourly for 24 h by measuring the increase in OD590 due to reduction of the redox indicator tetrazolium violet. All fluids, standards and PM plates were purchased from Biolog. Data were collected from two independent screens in which the differences between the tetrazolium violet reduction rates under each condition were less than 10 %.
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RESULTS |
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), but the precise transcript start site has not been mapped. Here, RACE-PCR was used to identify the ydhY transcript start site (Fig. 1a), which placed the centre of the FNR site at –42.5, a class II location (Fig. 1b). Evidence for direct interaction of FNR with the proposed FNR binding site upstream of ydhY was sought using gel retardation assays. To simplify the experiments, an FNR protein (FNR-D154A; referred to as FNR*) that retains some DNA binding activity in the presence of air was used in the gel retardation assays (Ziegelhoffer & Kiley, 1995). These showed that FNR* was able to bind to the unaltered PydhY but not to the same promoter fragment carrying mutations in the predicted FNR site (Fig. 1c). Furthermore, mutation of the FNR site in a plasmid-based ydhY–lacZ fusion abolished FNR-mediated regulation (Fig. 1d), and binding of FNR* at the FNR site in the ydhY promoter was confirmed by DNase I footprinting (Fig. 1e). Thus, the data presented are entirely consistent with the findings of Kang et al. (2005).
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) raised the possibility that ydhY is the first gene of a six-gene operon (ydhYVWXUT). The lengths (in nucleotides) of the intergenic regions are: 20 ydhY–ydhV; 3 ydhV–ydhW; 12 ydhW–ydhX; –4 ydhX–ydhU; and 3 ydhU–ydhT (Fig. 2). Therefore, RT-PCR using RNA isolated from anaerobic cultures of E. coli was used to determine the approximate downstream limits of the ydhY transcript. Using pairs of primers in adjacent genes it was established that ydhY and ydhV are co-transcribed, as are ydhV, W and X, and ydhX, U and T (Fig. 2). This shows that the ydhY–T genes are co-expressed from the FNR-dependent class II promoter upstream of ydhY.
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ydhY–lacZ) were grown in the presence of fumarate, DMSO, trimethylamine-N-oxide, nitrate or nitrite. In the presence of glucose, anaerobic expression of ydhY–T was approximately twofold lower than that of cultures lacking glucose (Table 2). Furthermore, β-galactosidase activity from the single-copy ydhY–lacZ fusion under aerobic conditions in the absence of FNR was increased approximately twofold compared with the parent, confirming the observations made with the multi-copy plasmid-based reporter (Fig. 1d
). The mechanism of this apparent FNR-mediated repression under aerobic conditions is unknown at this time. Addition of fumarate did not significantly change ydhY–T expression under anaerobic conditions. However, supplementation of the medium with DMSO or trimethylamine N-oxide enhanced anaerobic ydhY–T expression, whereas anaerobic expression was significantly lower in the presence of nitrate or nitrite (Table 2).
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), suggesting a role for both two-component systems in the regulation of ydhY–T expression. This pattern of regulation, along with the predicted functions of the encoded proteins, is consistent with the suggestion that the ydhY–T operon is associated with anaerobic respiration, perhaps by contributing to the maintenance of redox balance in the presence of specific electron acceptors.
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). One heptamer, centred at –42 (heptamer –42), matches the consensus (indicated by underlining) at five positions (TAACGAT), and is in the middle of the FNR site (centred at –42.5), and thus is located in a similar position to a NarL heptamer (centred at –50), which overlaps the FNR site in the nrfA promoter (Darwin et al., 1997). A second heptamer (heptamer –16) that also matches the consensus at five positions is centred at –16 (TAATCAC). The two remaining heptamers (heptamers +6 and +15) both match the consensus at six positions and are arranged as a 7-2-7 inverted repeat centred at +10.5 (TAACCATaaAGGATTA). Whereas NarL can potentially recognize all these heptamers, NarP only interacts with heptamers in a 7-2-7 arrangement (Darwin et al., 1997). DNase I footprinting confirmed that NarP recognized the predicted 7-2-7 site (Fig. 3a). The MBP–NarP footprint covered 33 bp (–10 to +23) punctuated by two hypersensitive sites (+6 and +7), and thus the NarP footprint at the ydhY–T promoter is similar to that observed at nirB (Darwin et al., 1997). Like NarP, NarL clearly protected the 7-2-7 region, but in contrast to NarP, the NarL footprint extended further upstream to –29 and thus encompassed the –16 heptamer, with hypersensitive sites at +19 and –15 (Fig. 3b). There was no obvious protection across the –42 heptamer; although the intensity of the DNA fragments in this region appeared weaker, no clear boundaries could be defined. This might be due to the –42 heptamer being a poorer match to the Nar consensus than the 7-2-7 site. The extended region of protection observed with NarL compared to NarP was consistent with the presence of only one retarded NarP–ydhY complex in gel retardation assays, whereas with NarL, multiple complexes were resolved (Fig. 3c). The PydhY–NarP complex also appeared to be more stable than the PydhY–NarL complexes in the gel retardation assay (as evidenced by the smeared appearance of the bands corresponding to the NarL complexes).
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). The altered promoter regions were fused to lacZ in the reporter plasmid pRS415 and transferred to the chromosome of E. coli MC1000 (lac). After growth of E. coli MC1000 strains containing the various promoter mutants at 37 °C to OD600 0.4–0.6 under anaerobic conditions in L broth or L broth containing nitrate (40 mM) or nitrite (5 mM), promoter activities were determined by measuring β-galactosidase. The results (Table 4) confirmed the response of the ydhY–T promoter to the presence of nitrate and nitrite, as described above. Mutation of the –42 heptamer (overlapping the FNR site) impaired repression in the presence of nitrate, but not nitrite (Table 4), despite the absence of clear protection of the –42 heptamer by phospho-NarL in the footprinting assay. Mutation of the upstream element (+6) of the 7-2-7 site impaired repression by both nitrate and nitrite, showing that this site is important for both nitrate- and nitrite-mediated repression (Table 4). However, mutation of the downstream element (+15) of the 7-2-7 site resulted in a promoter with lower activity than that of the unaltered promoter in the absence of nitrate or nitrite. Repression by nitrate was unaffected by mutation of the +15 heptamer (Table 4). Although nitrite was a less effective repressor when the +15 heptamer was mutated, the effect was not as obvious as when the +6 heptamer of the 7-2-7 site was disrupted. A plausible explanation for these observations is that mutation of the +6 or +15 heptamers inhibits phospho-NarP binding, allowing nitrite-activated phospho-NarL to bind at the remaining unaltered heptamer (either at the +6 heptamer, where NarL acts as an effective repressor, or at the +15 heptamer, where NarL does not contribute significantly to repression). In this way the effect on nitrite-mediated repression of mutating the +6 heptamer will be more apparent than mutation of the +15 heptamer. Mutation of the –16 heptamer did not affect repression by nitrate, despite the protection afforded to this region of the promoter by NarL (Fig. 3b). However, mutation of the –16 heptamer enhanced repression in the presence of nitrite, suggesting that NarL binding at –16 antagonizes NarP action at the 7-2-7 site. Overall, the simplest explanation for the results obtained is that the 7-2-7 site is involved in nitrite repression, the–42 heptamer and the upstream element of the 7-2-7 site contribute to nitrate repression, and the –16 site modulates the response to nitrite. This is consistent with the known DNA binding properties of NarP (recognizes 7-2-7 sites) and NarL (recognizes both 7-2-7 sites and isolated heptamers) (Darwin et al., 1997).
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A ydhY–T mutant does not exhibit an anaerobic growth defect
Anaerobic batch cultures of E. coli W3110 and two independent ydhY–T operon mutants exhibited the same growth rates and growth yields when L broth, L broth plus glucose (0.4 %), or Evans minimal medium plus glucose (20 mM) was used as the medium (results not shown). Furthermore, measurement of final OD600 of cultures of W3110 and the ydhY–T mutant grown in L broth plus glucose (0.4 %) supplemented with nitrate (40 mM) or nitrite (5 mM) indicated that the growth of the mutant was not impaired in the presence of these alternative electron acceptors (not shown). Biolog PM phenotyping arrays use the reduction of a tetrazolium dye to integrate respiration of bacteria during an experiment (Bochner, 2003). Ninety-six-well plates containing a different medium in each well are inoculated and a colour change resulting from the reduction of the redox dye is used to monitor respiratory activity of the culture. This showed that the ydhY–T mutant was not significantly impaired on any of the carbon, nitrogen or phosphorous sources tested. However, reduction of the tetrazolium dye was lower for the ydhY–T mutant when several compounds (L-cysteine, D-cysteine, L-cysteic acid, hypotaurine and butane sulphonic acid) were supplied as the sole source of sulphur, but these differences were not apparent when measured as final OD600 readings in simple anaerobic growth tests in the presence of L-cysteine or D-cysteine in glucose–Evans minimal medium (results not shown). It has been shown elsewhere that E. coli is able to utilize tetrahydrothiophene 1-oxide (THTO) as an electron acceptor under anaerobic conditions. However, the genes required were not identified (Meganathan & Schrementi, 1987). Growth of the ydhY–T mutant JRG5199 in Evans minimal medium supplemented with 0.1 % casamino acids and either 0.3 % (w/v) glucose or 0.5 % (w/v) glycerol as carbon source in the presence of THTO (25 mM) was similar to that of the parental W3110 strain (results not shown). Similar results were obtained when 1 mM tungsten (possibly required for YdhV activity; see above) was supplied in place of molybdenum (not shown). Thus, it was concluded that although the ydhY–T operon appears to play a role related to anaerobic metabolism of some sulphur compounds, this does not include THTO utilization under the conditions tested.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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), because the equivalent chromosomal location in Salmonella enterica serovar Typhimurium is occupied by the ttr operon, encoding tetrathionate reductase (Hensel et al., 1997).
Consistent with the hypothesis that YdhY–T has a role in anaerobic metabolism, gel retardation, footprinting, site-directed mutagenesis and transcript-mapping experiments showed that the oxygen-responsive transcription factor FNR directly activates transcription of ydhY–T under anaerobic conditions by binding at a site located at position –42.5 relative to the transcript start, confirming and extending earlier work that shows that ydhY is expressed from a class II FNR-dependent promoter (Kang et al., 2005). Furthermore, ydhY–T expression is regulated by the NarXL and NarPQ nitrate- and nitrite-responsive two-component systems, confirming the results of previous transcription-profiling studies (Constantinidou et al., 2006).
DNA sequence analysis identified four potential Nar binding heptamers (Fig. 1b). Footprinting assays showed that NarP recognized the 7-2-7 site centred at +10.5 and that NarL recognized a larger region of the ydhY–T promoter that included the 7-2-7 site and –16 heptamer. No clear protection of the –42 heptamer by NarL was detected. However, site-directed mutagenesis indicated that the –16 heptamer and the +15 heptamer, although protected in the in vitro footprinting assays, were not essential for nitrate-responsive ydhY–T expression in vivo, whereas both the –42 and the +6 heptamer made significant contributions, despite the absence of strong protection by NarL of the –42 heptamer in DNase I footprints. This suggests that in vivo there are other factors that lead to occupation of the –42 heptamer by NarL in the presence of nitrate. The –42 heptamer is located in the middle of the FNR site and the overlap of the FNR and NarL sites suggests that in vivo these proteins compete for occupation of this region of the promoter, with FNR acting positively and NarL negatively to control ydhY–T expression. This simple mechanism should ensure that ydhY–T is maximally expressed in the absence of both oxygen and nitrate. In contrast to NarL, the nitrite-responsive NarP protein recognizes heptamers organized as 7-2-7 inverted repeats (Darwin et al., 1997), such as heptamers +6 and +15. Inactivation of either the +6 or +15 heptamer affected ydhY–T expression in the presence of nitrite, suggesting that NarP acts negatively at this site, presumably by competing with RNA polymerase. The enhanced repression observed in the presence of nitrite when the –16 heptamer was mutated suggests that the presence of NarL at the –16 site inhibits the action of NarP at the 7-2-7 site. Detailed in vitro analysis of NarL/NarP–ydhY promoter interactions will be necessary to further investigate this possibility. Mutation of the +6 heptamer site also impaired nitrate-mediated repression, suggesting that NarL also inhibits RNA polymerase binding by occluding the transcription start point. Thus, it appears that NarL and NarP adopt overlapping mechanisms to inhibit ydhY–T expression. Both compete with RNA polymerase for occupation of the ydhY promoter in the region of the transcript start, but in addition, NarL competes with FNR to inhibit FNR-mediated anaerobic activation of ydhY–T expression. This additional feature, i.e. NarL is capable of occupying more of the ydhY–T promoter than NarP, could explain why NarL appears to be more effective than NarP in inhibiting ydhY–T expression. NarL-mediated repression of the frdA promoter is achieved by NarL binding over a large region centred near the transcription start site and including the FNR site (Li et al., 1994). Similarly, at the FNR-activated NarL-repressed dmsA promoter NarL protects a large region that includes the sites for both FNR and RNA polymerase binding (Bearson et al., 2002). Thus, NarL-mediated repression of the ydhY promoter follows the general pattern established by the frdA and dmsA promoters.
Whilst several NarL-repressed FNR-dependent promoters have been studied, FNR-dependent promoters that are repressed by NarP are uncommon because NarP tends to act as an activator, although recent transcript-profiling experiments suggest that NarP might have a wider role as a repressor than previously thought (Constantinidou et al., 2006). The transcript-profiling experiments reveal that a number of transcripts are less abundant in cultures of the narXL mutant grown in the presence of nitrate (Constantinidou et al., 2006). Examination of a narXL narP double mutant indicates that the absence of NarP enhances the abundance of transcripts from 37 operons, mainly associated with hydrogen and dicarboxylate metabolism (Constantinidou et al., 2006). Like ydhY–T, several of the operons suggested to be NarP-repressed, on the basis of transcript profiling, possess credible matches to the NarP consensus site close to the transcript start site, consistent with NarP-mediated repression (Constantinidou et al., 2006). NarP has been reported to act as a negative regulator of the FNR-dependent class II fdnGHI promoter under low-nitrate conditions (Wang & Gunsalus, 2003), although this promoter has also been shown to be weakly activated by NarP (Rabin & Stewart, 1993). Nevertheless, in contrast to the ydhY–T promoter, where NarP-mediated repression requires a 7-2-7 site centred at +10.5, occupation of which will probably inhibit RNA polymerase binding, the fdnGHI promoter possesses a NarP-binding 7-2-7 site centred at –104.5 that is thought to antagonize activation by NarL (Wang & Gunsalus, 2003). Thus, the architecture of the ydhY–T promoter appears to be somewhat different from that of previously characterized FNR-, NarL- and NarP-regulated promoters, having evolved to be repressed in the presence of nitrate and nitrite by using NarP/NarL to inhibit RNA polymerase binding, and NarL to inhibit FNR binding. The relative degrees of repression of the ydhY–lacZ fusion in the presence of nitrite (approximately twofold) or nitrate (13-fold) suggest that NarL-mediated inhibition of FNR recruitment is a key component of the regulation of the ydhY–T promoter, and that NarP-mediated inhibition of RNA polymerase binding is partially overcome by the ability of FNR to recruit RNA polymerase.
Whilst it is clear that FNR, NarL and NarP control expression of ydhY–T, there are likely to be additional regulatory features. For example, expression is lowered in the presence of glucose (Table 2), and it is possible that a specific regulator that responds to the presence of the YdhY–T substrate is required for full activation. Identification of the substrate for the YdhY–T-catalysed reaction is thus an important goal for future work.
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ACKNOWLEDGEMENTS |
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Edited by: J. W. B. Moir
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Received 30 July 2007;
revised 11 October 2007;
accepted 9 November 2007.
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ATCATAACGATGAT), lanes 3 and 4] were incubated with (+) or without (–) 8 µM FNR* (FNR-D154A) for 10 min before separating FNR*–DNA complexes (C) from free DNA (F) in TBE-buffered polyacrylamide gels and subsequent visualization by autoradiography. (d) Anaerobic activation of ydhY expression is abolished by mutation of the consensus FNR site. Anaerobic cultures of MC1000 (lac) or JRG1728 (fnr lac) containing pGS1739 (ydhY–lacZ) or pGS1831 (ydhY–lacZ with the FNR site inactivated as above) were grown at 37 °C in L broth to OD600 0.4–0.6. Promoter activities were determined by measuring β-galactosidase activities. Values are means±SD of triplicate assays from three independent cultures. (e) FNR* DNase I footprint at the ydhY promoter. The ydhY promoter was radiolabelled on the top strand. The reaction in the track labelled ‘–’ contained no FNR* protein; the reaction in the track labelled FNR contained FNR* (2 µM dimer). The solid line below the panel indicates the region of protection. The gel was calibrated with a Maxam and Gilbert G track (G).
