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• W2124345503 abstract "Article1 July 2002free access NO sensing by FNR: regulation of the Escherichia coli NO-detoxifying flavohaemoglobin, Hmp Hugo Cruz-Ramos Hugo Cruz-Ramos The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Jason Crack Jason Crack Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences, University of East Anglia, Norwich, NR4 7TJ UK Search for more papers by this author Guanghui Wu Guanghui Wu The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Martin N. Hughes Martin N. Hughes Chemistry Department, King's College London, Strand, London, WC2R 2LS UK Search for more papers by this author Colin Scott Colin Scott The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Andrew J. Thomson Andrew J. Thomson Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences, University of East Anglia, Norwich, NR4 7TJ UK Search for more papers by this author Jeffrey Green Jeffrey Green The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Robert K. Poole Corresponding Author Robert K. Poole The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Hugo Cruz-Ramos Hugo Cruz-Ramos The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Jason Crack Jason Crack Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences, University of East Anglia, Norwich, NR4 7TJ UK Search for more papers by this author Guanghui Wu Guanghui Wu The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Martin N. Hughes Martin N. Hughes Chemistry Department, King's College London, Strand, London, WC2R 2LS UK Search for more papers by this author Colin Scott Colin Scott The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Andrew J. Thomson Andrew J. Thomson Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences, University of East Anglia, Norwich, NR4 7TJ UK Search for more papers by this author Jeffrey Green Jeffrey Green The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Robert K. Poole Corresponding Author Robert K. Poole The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK Search for more papers by this author Author Information Hugo Cruz-Ramos1, Jason Crack2, Guanghui Wu1, Martin N. Hughes3, Colin Scott1, Andrew J. Thomson2, Jeffrey Green1 and Robert K. Poole 1 1The Krebs Institute for Biomolecular Research, Department of Molecular Biology and Biotechnology, The University of Sheffield, Sheffield, S10 2TN UK 2Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences, University of East Anglia, Norwich, NR4 7TJ UK 3Chemistry Department, King's College London, Strand, London, WC2R 2LS UK *Corresponding author. E-mail: [email protected] The EMBO Journal (2002)21:3235-3244https://doi.org/10.1093/emboj/cdf339 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Nitric oxide (NO) is a signalling and defence molecule of major importance in biology. The flavohaemoglobin Hmp of Escherichia coli is involved in protective responses to NO. Because hmp gene transcription is repressed by the O2-responsive regulator FNR, we investigated whether FNR also senses NO. The [4Fe–4S]2+ cluster of FNR is oxygen labile and controls protein dimerization and site-specific DNA binding. NO reacts anaerobically with the Fe–S cluster of purified FNR, generating spectral changes consistent with formation of a dinitrosyl-iron–cysteine complex. NO-inactivated FNR can be reconstituted, suggesting physiological relevance. FNR binds at an FNR box within the hmp promoter (Phmp). FNR samples inactivated by either O2 or NO bind specifically to Phmp, but with lower affinity. Dose-dependent up-regulation of Phmp in vivo by NO concentrations of pathophysiological relevance is abolished by fnr mutation, and NO also modulates expression from model FNR-regulated promoters. Thus, FNR can respond to not only O2, but also NO, with major implications for global gene regulation in bacteria. We propose an NO-mediated mechanism of hmp regulation by which E.coli responds to NO challenge. Introduction Nitric oxide (NO) is a signalling and defence molecule of major importance, but the biochemical basis of resistance to NO and reactive nitrogen species (RNS) is poorly understood. NO and RNS pose distinct biochemical challenges by reacting under appropriate conditions with several key biomolecules including metalloproteins, thiol groups of proteins and low molecular weight thiols such as glutathione and homocysteine (Hcy) (Poole and Hughes, 2000). These cellular targets may themselves serve as signal transducers, sensing the presence of NO and RNS, resulting in changes in gene expression and consequent synthesis of protective enzymes. Regulation of transcriptional activation in some cases integrates the responses to free radicals of both oxidative and nitrosative stresses. Thus, in Escherichia coli, the soxRS and oxyR regulons, known to respond to superoxide anion and hydrogen peroxide, respectively, are also induced by NO (Nunoshiba et al., 1993; Hausladen et al., 1996; Ding and Demple, 2000). These global antioxidant regulators afford protection against NO or nitrosothiols (Nunoshiba et al., 1993; Hausladen et al., 1996), but the resistance mechanisms are not entirely known, despite their importance in allowing bacteria to combat, for example, macrophage microbicidal activity (De Groote et al., 1997). The most fully understood mechanism for detoxification of NO involves the bacterial flavohaemoglobin (Hmp). Hmp synthesis is activated primarily by NO or RNS, and the physiological function of Hmp in protecting E.coli from NO is established (Poole et al., 1996; Membrillo-Hernández et al., 1999; Poole and Hughes, 2000). Aerobically, Hmp detoxifies NO by acting as an NO oxygenase (Gardner et al., 1998a; Hausladen et al., 1998) or denitrosylase (Hausladen et al., 2001) and affords inducible protection of aconitase activity (Gardner et al., 1998b) and respiration (Stevanin et al., 2000). Anaerobically, Hmp serves as an NO reductase, generating nitrous oxide (Hausladen et al., 1998; Kim et al., 1999). Consistent with the role of Hmp in NO detoxification, the flavohaemoglobin-encoding gene of E.coli, hmp, is up-regulated by NO and RNS; this appears not to involve SoxRS (Poole et al., 1996). We have reported (Membrillo-Hernández et al., 1998) a mechanism of hmp gene regulation that involves interaction between S-nitrosothiols and Hcy. Intracellular Hcy is an important co-regulator of several genes involved in methionine biosynthesis, via its effects on MetR, a LysR family DNA-binding protein. One gene activated by MetR with Hcy as cofactor is glyA, which, in E.coli, is adjacent to hmp and divergently transcribed from it. Elevated Hcy levels, achieved either by exogenous Hcy or in certain met mutants, decrease hmp expression. Since Hcy has been shown to be nitrosated by S-nitrosoglutathione (GSNO) (Membrillo-Hernández et al., 1998), such nitrosating agents can deplete Hcy pool sizes, and are postulated to enhance MetR binding at a site proximal to hmp, and up-regulate hmp transcription (Membrillo-Hernández et al., 1998). It is important to recognize that this mechanism does not explain hmp regulation by NO itself. First, NO induces hmp expression anoxically, under which conditions NO will not nitrosate Hcy. Secondly, although the S-nitrosoHcy generated on reaction with GSNO breaks down to release NO, which could itself be the inducer, the reaction of Hcy with sodium nitroprusside (SNP) (which also induces hmp) forms a more stable species from which NO is not released. Thus other mechanisms for hmp regulation by NO, particularly anoxically, must be present. Anaerobically, the global regulator FNR (Kiley and Beinert, 1999; Green et al., 2001) is also involved in the regulation of hmp since an fnr mutation enhances hmp-lacZ expression (Poole et al., 1996). Inspection of the hmp promoter reveals a DNA-binding site for FNR in a repressing position, but how FNR contributes to NO-mediated regulation has remained obscure. Up-regulation of hmp in Salmonella is achieved by a third mechanism involving the iron uptake regulator Fur (Crawford and Goldberg, 1998). Here, we report an anaerobic regulatory mechanism of hmp induction directly mediated by NO, involving nitrosylation of the [4Fe–4S]2+ clusters in the FNR protein. NO sensing by FNR has far-reaching implications for a full understanding of its global regulatory functions. Results FNR is an NO sensor The starting point for this work was the observation that the oxygen-responsive global transcription factor FNR represses expression of the NO-detoxifying protein Hmp during anaerobic growth (Poole et al., 1996). FNR senses oxygen via the assembly–disassembly of [4Fe–4S]2+ clusters (Green et al., 1996; Khoroshilova et al., 1997; Kiley and Beinert, 1999). Iron–sulfur clusters of other proteins have been shown to react with NO, and thus the clusters of FNR might also constitute a mechanism to account for NO responsiveness of hmp expression. To address this possibility, the effects of NO on the [4Fe–4S]2+ clusters of FNR were investigated in vitro. The anoxic, reconstituted FNR protein (holo-FNR) showed the characteristic broad absorbance at ∼420 nm (Khoroshilova et al., 1995; Green et al., 1996; Lazzazera et al., 1996; Jordan et al., 1997) and was devoid of electron parmagnetic resonance (EPR) signals. On addition of the fast NO-releasing compound, proline NONOate, the broad absorbance at 420 nm was replaced progressively by a prominent shoulder at 360 nm and a decrease in absorbance at 420 nm with an isosbestic point near 480 nm (Figure 1A). A plot of ΔA360 versus NO concentration shows that the reaction is complete at a [NO]/[FNR] ratio of <3.0 (Figure 1A inset and B). Figure 1.Effect of NO on FNR. (A) Optical spectra of [4Fe–4S]2+ FNR (27 μM, monomer) before and after treatment with aliquots of 4.17 mM proline NONOate to give a final concentration of 140 μM NO (72 μM NONOate). Inset: a plot of ΔA360 versus the ratio of [NO]/[FNR]. When no further change was apparent, a sample was withdrawn for EPR analysis. (B) Optical spectra of the same FNR suspension acquired after sequential additions of proline NONOate. The lowest spectrum is FNR in the absence of NO and the uppermost one is at a [NO]/[FNR] ratio of 5.2. The [NO]/[FNR] ratio increases from 0, 0.2, 0.3, 0.6, 1.2, 2.2, 2.9, 3.7 to 5.2, giving rise to dose-dependent absorbance increases at 360 nm. Download figure Download PowerPoint Dinitrosyl-iron–cysteine (DNIC) complexes have relatively intense optical absorption bonds in the near UV. Figure 2A displays the absorption spectra of yellow and green DNICs having dimeric and monomeric structures (inset), respectively. The extinction coefficients for the two species are as follows: dimeric clusters ϵ310 = 9650/M/cm, ϵ362 = 8529/M/cm and ϵ750 = 100/M/cm; monomeric form ϵ362 ∼3580/M/cm, ϵ603 = 300/M/cm and ϵ772 = 312/M/cm. The dimeric DNIC is EPR silent, whereas the monomeric form has one unpaired electron, S = 1/2, with an axial g-tensor, g = 2.038, 2.024 and 2.06. The EPR spectrum of a sample taken from the titration of FNR with proline NONOate at a [NO]/[FNR] ratio of ∼5 shows just such an axial EPR spectrum (Figure 3A). This signal is strongly reminiscent of the DNIC of cysteine (Kennedy et al., 1997; Vanin et al., 1998). Integration of the g = 2.038 signal gave only 20% of an electron spin per [4Fe–4S]2+ cluster. A minor species appearing at g = 2.15 may arise from a thiyl radical (data not shown; Kennedy et al., 1997). The EPR spectra measured at low temperature (not shown) revealed a trace amount of a [3Fe–4S]1+ cluster at g = 2.01. Figure 2.Optical spectra of DNIC complexes. (A) Model complexes. The monomeric species (445 μM; solid line) displays characteristic absorption maxima at 397 nm (ϵ ∼3580/M/cm), 603 nm (ϵ ∼299/M/cm) and 772 nm (ϵ ∼312/M/cm) (Costanzo et al., 2001). The dimeric species (250 μM, dashed line) displays characteristic absorption maxima at 310 nm (ϵ ∼9650/M/cm), 362 nm (ϵ ∼8529/M/cm) and 750 nm (ϵ ∼100 M/cm). Inset: structures proposed for monomeric and dimeric DNIC species (Costanzo et al., 2001). (B) Comparison between experimental (dashed line) and simulated (black line) spectra of DNICs. The simulated spectrum consists of contributions from 20 and 80% of the monomeric and dimeric DNIC species, respectively, plus apoFNR. Download figure Download PowerPoint Figure 3.EPR spectra of [4Fe–4S]2+ FNR. (A) The FNR sample used in Figure 1, after treatment with 140 μM NO (72 μM proline NONOate; 27 μM FNR monomer). (B) Increasing magnitudes of EPR signals elicited by addition of aliquots of NO solution (NO and FNR concentrations were as indicated). The temperature was 77 K. Microwave power and frequency were in (A) 2.007 mW and 9.653 GHz, and in (B) 2.000 mW and 9.669 GHz, respectively. Modulation amplitude, frequency and receiver gain were in (A) 5 gauss (0.5 mT), 100 kHz and 5.02 × 105 and in (B) 10 gauss (1 mT), 100 kHz and 1.00 × 105, respectively. Download figure Download PowerPoint In a parallel set of experiments, sequential additions of NO solution or air-saturated buffer (∼230 μM O2) to reconstituted FNR also caused absorbance decreases at 420 nm. Some difficulty was experienced with slow protein precipitation causing baseline variations but, during this titration with NO, samples were withdrawn for EPR spectroscopy. The main species detected at 70 K again was typical for a monomeric DNIC (Figure 3B). For example, the EPR spectra of NO-treated FNR are very similar to the EPR spectra of the d7 form of DNIC-aconitase (Kennedy et al., 1997). At 10 K, varying intensities of the signal for the [3Fe–4S]1+ cluster (g = 2.02) were also seen, as previously observed when [4Fe–4S]2+ FNR is briefly exposed to oxygen (Green et al., 1996). The experimental EPR results (Figure 3) and optical spectra (Figure 1) suggest a mixture of the yellow and green forms of DNIC. We therefore simulated the optical spectrum by combining spectra of the monomeric and dimeric forms in the ratio 20:80. The result is shown in Figure 2B. The match to the experimental spectrum (Figure 2B, dashed line) is good, although the presence of some light-scattering precipitate causes background absorption sloping towards the UV. An FNR sample exposed to NO solution and lacking the 420 nm peak was flushed with N2 gas for 60 min, then used in a standard procedure previously shown to be effective for reconstitution of the Fe–S cluster (Jordan et al., 1997). Importantly, after overnight incubation, the protein showed the same optical features (420 nm maximum) as an NO-untreated, but reconstituted, sample (data not shown). These data strongly suggest that the [4Fe–4S]2+ cluster of FNR can react with NO to form a mixture of monomeric and dimeric DNICs, and that the NO-treated protein remains native and can be reconstituted. Thus the conversion of the native [4Fe–4S]2+ cluster to DNICs could provide the molecular basis of a direct NO-responsive switch. Interaction of FNR with wild-type and mutant Phmp A putative FNR box, TTGAG----ATCAA, with a strong resemblance to the consensus FNR-binding sequence is centred at position +5.5 in the Phmp region. To determine whether the putative FNR box is actually recognized by FNR, mutations were made in which the central G (underlined above) was mutated to A, and the corresponding C (also underlined), in the second half of the box, was changed to T. The interaction of FNR with its target sequence was then analysed by gel retardation. At holo-FNR concentrations as low as 0.4 and 0.5 μM, 9 and 20% of the wild-type Phmp, respectively, is retarded (Figure 4A). No retardation could be detected when the FNR box was mutated and incubated with holo-FNR (Figure 4D). Interestingly, wild-type Phmp was also retarded (11 and 26%) by dioxygen- and NO-treated FNR lacking the 420 nm peak, but this occurred only with higher protein concentrations (i.e. >>2 μM) (Figure 4B and C). The retardation profile suggests that holo-FNR, as well as O2- and NO-treated FNR, may bind Phmp in a similar, namely dimeric, form, but with different affinities. The protein–DNA dissociation constants (Kd) for the three FNR forms were determined by gel retardation and repressor–operator interaction analyses. Both titration methods showed similar results. O2- and NO-treated FNR had similar affinities for Phmp with apparent dissociation constants of Kd = 2.5–3 μM. The affinity for holo-FNR, Kd = 1 μM, is greater than that of either O2- or NO-treated FNR. Figure 4.DNA binding by holo-FNR, O2- and NO-treated FNR proteins. Gel retardation assays were performed by incubating the 32P-labelled Phmp DNA fragment (wild-type) with increasing amounts of (A) holo-FNR, (B) O2-treated FNR and (C) NO-treated FNR protein. The 32P- labelled Phmp DNA fragment (mutated) was incubated with increasing amounts of (D) holo-FNR. Arrows indicate the FNR–target DNA (black) and the FNR–Phmp complex (white). Protein concentrations (monomer) are shown. Download figure Download PowerPoint DNase I protection by holo-FNR, O2-treated and NO-treated FNR DNase I footprinting confirmed the different binding affinities of holo-FNR, O2- and NO-treated FNR obtained from titration experiments. A 26 bp region overlapping the FNR box and located between −6 and +20 in Phmp was protected by the addition of 0.2 μM holo-FNR (Figure 5A). Increasing protein concentrations also revealed hypersensitive sites. Once the FNR concentration reached 2–12 μM, an AT-rich hexamer centred at −15.5 appears to be protected from cleavage (Figure 5A and C). The same footprinting pattern was observed with both O2-treated FNR (data not shown) and NO-treated FNR (Figure 5B). Consistent with gel retardation results, 0.2 μM O2- or NO-treated FNR failed to protect the FNR box and protection was seen from 2 μM protein (Figure 5B). Again, the AT-rich hexamer and the hypersensitive sites were revealed by adding further protein. However, in the case of NO-treated FNR (Figure 5B), the hypersensitive +1G and +2A sites were substituted by a −1T site. Also, extensive non-specific DNA protection could be seen upstream of the hypersensitive sites found immediately 5′ to the AT-rich hexamer when higher concentrations of NO-treated FNR were used, indicating that this form of the protein not only has a lower affinity for DNA but also binds with reduced specificity (Figure 5B). Escherichia coli RNA polymerase (RNAP) was used in DNase I footprinting assays to determine whether the σ70-holoenzyme recognizes Phmp. By itself, RNAP bound to a 64 bp DNA region (−44 to +20) containing the −10 and −35 hexamers, as well as the FNR box (Figure 5C). Figure 5.DNase I protection by holo-FNR, NO-treated FNR and RNAP bound to Phmp. The 32P-labelled Phmp DNA fragment was digested with DNase I in the presence of various concentrations of (A and C) holo-FNR, (B) NO-treated FNR and (C) RNAP. Template strand patterns shown were obtained with no FNR (lanes -), 0.2 μM (lanes 1), 2 μM (lanes 2), 12 μM (lanes 3) and 20 μM (lanes 4) FNR protein (as monomer), and with 69 nM RNAP (lane 5). Lanes G provide a calibration for the GC base pairs in the Phmp region. Hypersensitive sites are marked with black or white arrowheads. Hollow boxes show the region protected by dimeric FNR binding. Striped boxes show the position of a DNase I-protected AT-rich hexamer. The filled boxes indicate the region bound by RNAP σ70 subunit. The −10 and −35 hexamers, the ribosome-binding site (rbs), the transcription start site (+1) and the FNR box are indicated on the Phmp sequence shown below. Download figure Download PowerPoint In vivo modulation of FNR activity by NO Regulation of hmp transcription by NO and RNS appears complex, and at least one regulatory mechanism involving MetR is known (Membrillo-Hernández et al., 1998; Poole and Hughes, 2000). In order to assess the modulation of FNR activity by NO in vivo, we used an E.coli strain carrying a single copy of a Phmp-lacZ fusion (RKP2178). Previously, a bolus addition of a solution of NO gas was used to demonstrate up-regulation of hmp but, in view of the reactivity of NO and the demonstration that E.coli cells consume NO, even anaerobically (Kim et al., 1999), we have used both NO gas solutions (see later, Figure 7) and NO provided by its release from NOC-5 and NOC-7. Figure 6A (traces 1–3) shows the predicted progress of NO release from both NOC-5 and NOC-7 (10 μM of each, final concentration). The release from NOC-5 (half-life of 25 min at pH 7.0, 37°C) is shown as trace 1, and from the donor with a much shorter half life, NOC-7 (5 min at pH 7.0, 37°C) as trace 2. Trace 3 is the sum of the calculated individual NO release profiles. The actual NO concentration in solution (trace 4) was monitored continuously during anaerobic growth with an NO electrode; during the first 10 min of culture, the measured and theoretical NO levels were in good agreement but thereafter NO concentration declined. Nevertheless, significant NO in the range of concentrations previously shown to up-regulate a Phmp-lacZ fusion (Poole et al., 1996) were sustained in the experiment. Figure 6.NO derepresses hmp expression in vivo by reacting with FNR. (A) Release of NO by the NOC mixture over 1 h. Traces 1 and 2 are the theoretical NO release patterns at 37°C (in PBS pH 7.4) by NOC-7 (half-life 5 min) and NOC-5 (half-life 25 min), respectively. Trace 3 is the total NO release by the NOC mixture (addition of traces 1 and 2). Trace 4 is the measured NO release by the NOC mixture under the same conditions of NO challenge in anoxic LB-containing cells (OD600 of ∼0.2) at 37°C. Anaerobic cultures of (B) RKP2178 [φ(hmp-lacZ)1] and (C) RKP2185 (RKP2178 but fnr-271::Tn10) were challenged with different amounts of NOC mixture. The β-galactosidase activity is the mean of three independent repetitions of the experiment; bars show standard deviations. Download figure Download PowerPoint Figure 7.NO modulates gene expression in vivo by reacting with FNR. Anaerobic cultures of E.coli strains carrying (A) the FNR-inducible promoter [FF(−71.5)::lacZ] or (B) the FNR-repressible promoter (FFgalΔ4::lacZ) were treated with different amounts of NO. β-galactosidase activity is the mean of three independent experiments. Download figure Download PowerPoint To determine whether these concentrations of NOC compounds or NO generated were cytotoxic, the anaerobic growth of cultures of strains RKP2178 [φ(hmp-lacZ)] or RKP2185 [φ(hmp-lacZ) fnr] (see below) was measured. NOC-5 and NOC-7 were added (5 or 10 μM of each) to exponentially growing cultures that had been pre-grown in sealed tubes to a Klett reading of 20–25. In the first 30 min after NOC addition, growth rates declined slightly but then resumed the rates characteristic of control cultures (data not shown). In an additional test of toxicity, measurements of viable cell numbers were made during a similar anaerobic experiment. Wild-type hmp+ cells were resistant to 500 μM of each NOC compound, whereas hmp mutant cells (strain RKP4600) were sensitive to 50 μM of each NOC compound (results not shown). We have also shown that E.coli can be grown at NOC concentrations as high as 5 mM (M.Binet, J.Laver, H.Cruz-Ramos, R.K.Poole, unpublished data). Thus, NOC compounds in cultures sustain NO in solution over 30 min at concentrations that up-regulate hmp yet are not toxic for growth. β-galactosidase activities were assayed following the addition of different concentrations of an NOC mixture. Increasing concentrations of NOC compounds elicited an increase in φ(hmp-lacZ) activity in a wild-type background (Figure 6B) but, in an fnr strain, NOC addition was without effect on the derepressed level of φ(hmp-lacZ) activity (Figure 6C). These results are consistent with NO reacting with FNR in vivo with consequent derepression of hmp transcription, but additional regulatory mechanisms are suggested by the fact that the derepressed level in the wild-type strain (Figure 6B) is lower than in the fnr mutant (Figure 6C). In order to assess (i) the modulation of FNR activity by NO in vivo in a simpler system, and (ii) the implications for regulation of other genes in the FNR regulon, we used the pRW50[FF(−71.5)::lacZ] and pRW50(FFgalΔ4::lacZ) fusion plasmids (Williams et al., 1998) providing relatively simple semi-synthetic promoters, at which induction or repression is totally dependent on anaerobic FNR binding. These constructs were used to transform Δlac and Δlac Δfnr strains. The β-galactosidase activities of the anaerobically grown strains were assayed following the addition of different volumes of NO solution. It was anticipated that any disruption by NO of FNR might reduce its inducing activity on FF(−71.5)::lacZ and derepress the FFgalΔ4::lacZ promoter fusion. The titration in vivo of the FNR-inducible promoter [FF(−71.5)] with NO showed a progressive decrease in β-galactosidase activity, reaching only 40% of the NO-untreated level at 25 μM NO (Figure 7A). The titration curve of the FNR-repressible promoter (FFgalΔ4) showed a bell-like shape (Figure 7B). Promoter derepression was observed from 5 to 10 μM NO (1.6- to 2.5-fold increase, respectively) and the highest reporter activity (∼5-fold increase) was reached with 25 μM NO. These data showed that NO is effective as a signal molecule in a relatively narrow concentration range. NO concentrations above 25 μM appear to destabilize lacZ transcripts. As expected, the FF(−71.5) promoter (strain RKP2761) was inactive in the absence of fnr (93 ± 13 Miller units of β-galactosidase when 10, 20, 30 or 50 μM NO was added; data not shown). Again, as expected, the FFgalΔ4 promoter (strain RKP2762) was active in the fnr background regardless of the presence of NO (210 ± 30 Miller units of β-galactosidase when 10, 20, 30 or 50 μM NO was added). In control experiments, β-galactosidase activities were measured on strains [RKP2672 (Δlac) and RKP2760 (Δlac Δfnr)] carrying the pRW50 plasmid but without the promoter-lacZ fusion; in all cases, the activities were low (<2 Miller units of β-galactosidase) and showed no pattern of variation during anaerobic growth in the presence of NO. Discussion Here we present direct evidence of FNR binding to the Phmp in vitro by gel retardation, repressor–operator interaction and footprinting studies. The FNR box centred at position +5.5 was bound specifically by holo-FNR. RNAP occupied a DNA region overlapping the FNR box. Thus, this regulatory element is located appropriately for repression of transcription initiation by promoter occlusion. Point mutations of the central GC pairs within the FNR box were sufficient to abolish protein–DNA interaction, demonstrating the presence of a single FNR-binding site in Phmp. Other FNR-repressed promoters, such as ndh, narX and fnr, contain more than one FNR box (Guest et al., 1996), and complex regulatory mechanisms involving FNR have been described (Meng et al., 1997; Marshall et al., 2001). To our knowledge, Phmp is the first described naturally occurring promoter where repression by the E.coli FNR protein appears relatively simple, i.e. involving a single FNR box in the repressing position. The DNA-binding activity of FNR was affected by either O2 or NO treatment, resulting in a lower Phmp affinity and specificity. Nevertheless, some residual DNA-binding ability was observed after treatment of the FNR protein with either O2 or NO. Such residual DNA-binding activity has been reported previously for O2-treated FNR (Lazzazera et al., 1996). However, in other work with different DNA targets, treatment of FNR with O2 (Jordan et al., 1997) and either O2 or NO (Wu et al., 2000) abolished DNA binding. Thus, it is likely that the DNA target influences the ability of O2- or NO-treated FNR to bind. The footprinting data for NO-treated FNR indicate similar, but not identical, DNA distortions occurring upon binding of O2-treated FNR. DNase I-hypersensitive sites arise from protein-induced bending that creates kinks in the DNA structure. The DNase I hypersensitivities and the protected AT-rich hexamer found immediately 5′ to the FNR box remained, whereas the +1 FNR half-site appeared less susceptible to the endonuclease when NO-treated FNR was bound. Whether this difference in DNA binding relates t" @default.
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• W2124345503 date "2002-07-01" @default.
• W2124345503 modified "2023-10-16" @default.
• W2124345503 title "NO sensing by FNR: regulation of the Escherichia coli NO-detoxifying flavohaemoglobin, Hmp" @default.
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