Regulation of fructooligosaccharide metabolism in an extra-intestinal pathogenic Escherichia coli strain

Molecular Microbiology (2011) 81(3), 717–733 䊏

doi:10.1111/j.1365-2958.2011.07725.x First published online 22 June 2011

Regulation of fructooligosaccharide metabolism in an extra-intestinal pathogenic Escherichia coli strain mmi_7725 717..733

Gaëlle Porcheron,1 Emmanuel Kut,1 Sylvie Canepa,2 Marie-Christine Maurel2 and Catherine Schouler1* 1 INRA, UR1282 Infectiologie Animale et Santé Publique, F-37380 Nouzilly, France. 2 Laboratoire d’Analyses des Interactions Moléculaires, Plate-forme d’Analyse Intégrative des Biomarqueurs cellulaires et moléculaires (PAIB), UMR6175 Physiologie de la Reproduction et des Comportements, F-37380 Nouzilly, France.

Summary A gene cluster involved in the metabolism of prebiotic short-chain fructooligosaccharides (scFOS) has recently been identified in the extra-intestinal avian pathogenic Escherichia coli strain BEN2908. This gene cluster, called the fos locus, plays a major role in the initiation stage of chicken intestinal colonization. This locus is composed of six genes organized as an operon encoding a sugar transporter and enzymes involved in scFOS metabolism, and of a divergently transcribed gene encoding a transcriptional regulator, FosR, belonging to the LacI/GalR family. To decipher the regulation of scFOS metabolism, we monitored the fos operon promoter activity using a luciferase reporter gene assay. We demonstrated that the expression of fos genes is repressed by FosR, controlled by catabolite repression and induced in the presence of scFOS. Using electrophoretic mobility shift assays and surface plasmon resonance experiments, we showed that FosR binds to two operator sequences of the fos operon promoter region. This binding to DNA was inhibited in the presence of scFOS, especially by GF2. We then propose a model of scFOS metabolism regulation in a pathogenic bacterium, which will help to identify the environmental conditions required for fos gene expression and to understand the role of this locus in intestinal colonization.

Accepted 30 May, 2011. *For correspondence. E-mail catherine. [email protected]; Tel. (+33) 2 47 42 72 96; Fax (+33) 2 47 42 77 74.

© 2011 Blackwell Publishing Ltd

Introduction Fructooligosaccharides (FOS) are natural linear polymers comprising two to eight b-(2–1)-linked fructosyl units, usually attached to a terminal glucose residue (Roberfroid et al., 2010). Like many complex plant carbohydrates, these sugars are not hydrolysed by digestive enzymes, and therefore they reach the distal parts of the intestine intact where they are assimilated by the gastrointestinal microbiota, particularly probiotic bacteria (Gibson and Roberfroid, 1995; Roberfroid, 2001). This property of stimulating probiotic growth would be at the expense of pathogenic bacteria such as Escherichia coli or Samonella enterica serovar Typhimurium (Naughton et al., 2001; Sharp et al., 2001; Buddington et al., 2002). FOS have therefore been classified as prebiotics and are currently used commercially in food products and nutritional supplements for both humans and animals (Ritsema and Smeekens, 2003; Xu et al., 2003; Geier et al., 2009). Genes involved in FOS metabolism have mainly been described in non-pathogenic Gram-positive bacteria such as lactobacilli (Barrangou et al., 2003; Kaplan and Hutkins, 2003; Goh et al., 2006; Saulnier et al., 2007) and bifidobacteria (Ryan et al., 2005), which constitute part of the probiotic microbiota in the intestine. FOS uptake is mediated via an ATP-dependent binding cassette-type transporter in Lactobacillus acidophilus (Barrangou et al., 2003; Altermann et al., 2005) and Bifidobacterium longum (Parche et al., 2007; Gonzalez et al., 2008), via a phosphoenolpyruvate : carbohydrate phosphotransferase system (PTS) in Lactobacillus paracasei (Goh et al., 2006), and via a transporter of the major facilitator superfamily in Bifidobacterium breve (Ryan et al., 2005). However, it has recently been discovered that a pathogenic bacterium, the extra-intestinal avian pathogenic E. coli strain BEN2908, is also able to metabolize scFOS (short-chain FOS with two to four fructose units) (Schouler et al., 2009). Extra-intestinal pathogenic E. coli (ExPEC) strains are normal inhabitants of the gastrointestinal tract of humans and warm-blooded animals. They cause several extraintestinal diseases, including urinary-tract infections, neonatal meningitis, bacteraemia, septicaemia and respiratory-tract infections, the latter particularly in poultry (Russo and Johnson, 2000; Johnson and Russo, 2002; Smith et al., 2007). This respiratory disease is

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characterized by fibrinopurulent lesions such as air sacculitis, perihepatitis and pericarditis and is often associated with septicaemia and mortality (Dho-Moulin and Fairbrother, 1999; Barnes et al., 2003). The genomic region responsible for scFOS metabolism in the BEN2908 strain, called the fos locus, is found on the AGI-3 pathogenicity island, which possesses five loci bound by mobility-related genes (Chouikha et al., 2006). It has been shown that the fos locus improves chicken intestinal colonization by bacteria during the first 8 days post inoculation (Schouler et al., 2009). We can assume that scFOS metabolism helps the BEN2908 strain settle in its niche, and that, once established, this strain no longer requires this metabolism. This suggests a modulation of fos gene expression in vivo. The fos locus (GenBank accession No. AY857617) is composed of six genes putatively organized as an operon encoding for a putative major facilitator superfamily sugar transporter (FosT), two glycoside hydrolases of family 32 (FosE1 and FosE2), two proteins of unknown function (FosX and FosY) and a fructokinase (FosK), and a divergently transcribed gene encoding for a putative transcriptional regulator of the LacI/GalR family (FosR). Proteins of the LacI/GalR family repress transcription of the genes involved in carbon metabolism by binding to DNA in the absence of an inducer molecule. Each member of this family possesses an N-terminal DNA-binding domain, which forms a helix–turn–helix (HTH) motif, and a C-terminal domain involved in effector recognition and oligomerization. DNA operator sequences recognized by these regulators are inverted repeats with varying spacing (Weickert and Adhya, 1992; Fukami-Kobayashi et al., 2003; Swint-Kruse and Matthews, 2009). The presence within the fos region of the fosR gene encoding a putative transcriptional regulator of the LacI/GalR family strongly suggests that FosR could regulate the expression of fos genes. The inducer of FosR could either be the substrate transported and metabolized by proteins of the fos region, or a degradation product of this substrate, in other words, an scFOS or one of its derivatives. In order to better understand the role of the fos locus of the ExPEC strain BEN2908 in the colonization of its reservoir, we analysed the genetic organization, features and regulatory elements of the fos region and investigated the role of the transcriptional regulator FosR in the regulation of fos gene expression and the ability of scFOS and other carbon sources to induce this expression.

To provide further evidence that these genes are cotranscribed, we performed RT-PCR analysis of the region from fosT downstream to fosK using primer pairs connecting the 3′ end of each ORF to the 5′ end of the following ORFs. The results indicate that fosT to fosK was transcribed as a single mRNA (Fig. 1). We also showed that fosR was transcribed (Fig. 1). Fragments were also generated using primer pairs downstream of fosK (as far as 604 bp downstream of fosK) and within the fosK coding region (Fig. 1). No Rho-independent transcriptional terminator was detected in silico downstream fosK. Therefore, either a low level of transcription continues through a currently unidentified transcriptional terminator, or no transcriptional terminator is present immediately downstream of fosK, allowing the transcription to continue after fosK. Transcriptional start sites were mapped for the fos operon and fosR using 5′ RACE. The transcriptional start site of the fos operon was identified 25 bp upstream of the fosT start codon, and that of fosR was mapped 50 bp upstream of its start codon (Fig. 2A). Putative -10 and -35 regions were identified for both the fos operon and fosR. Regions upstream of the PfosT and PfosR start sites contain a putative s70 promoter with -10 (TACCAT) and -35 (GTGTCC) regions [in bold, nucleotides corresponding to the consensus sequence (Lewin, 2000)], and a 17-base spacer; -10 (TAGAAT) and -35 (TGCATA) regions and an 18-base spacer respectively (Fig. 2A). In addition to these two promoter sequences, in silico analysis revealed other noteworthy sequences within the IR between fosT and fosR (Fig. 2). We identified two palindromic sequences, O1 (nucleotides -4 to -25) and O2 (nucleotides -69 to -86). These sequences exhibit all the features of a LacI family binding site (Camas et al., 2010). Moreover, O1 is homologous to O2, and the latter is identical to the operator sequences recognized by ScrR (Jahreis and Lengeler, 1993; Novichkov et al., 2010) and CscR (Bockmann et al., 1992; Jahreis et al., 2002), the LacI/GalR family transcriptional regulators of sucrose metabolism in E. coli (Fig. 2B). We also identified a palindromic sequence (nucleotides -91 to -112) (Fig. 2A) similar to the consensus sequence recognized by CRPcAMP complexes [19 nucleotides corresponding to the consensus sequence (Lewin, 2000)]. All these elements strongly suggest that the transcription of fos genes could be under the control of the putative regulator FosR and catabolite repression.

Results

FosR, as a dimeric regulator, represses expression of fos genes

Promoter mapping, transcriptional organization and features of the intergenic region (IR) of the fos locus The fosT to fosK genes are transcribed in the same direction and appeared to be organized as an operon (Fig. 1A).

FosR (GenBank Accession No. AAW51730.1) is a 331 amino acid protein and has a calculated molecular mass of 37 kDa (41 kDa with the 6-His tag). FosR is homologous to transcriptional regulators of the LacI/GalR family © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 719

Fig. 1. Co-transcription within the fos locus. A. Gene organization of the fos locus. IS1, transposase; fosK, fructokinase; fosY, unknown function; fosE2, glycoside-hydrolase of family 32; fosX, unknown function; fosE1, glycoside-hydrolase of family 32; fosT, sugar transporter of the major facilitator superfamily; and fosR, transcriptional regulator of the LacI/GalR family. Transcripts are shown as thin lines below the gene cluster. B. Co-transcription of fos genes was revealed by RT-PCRs using primer pairs connecting the 3′ end of one ORF to the 5′ end of the following one. RT-PCRs were performed in the absence of reverse transcriptase to check for DNA contamination (lanes indicated by a – sign). Amplicons were analysed by electrophoresis using 1% agarose gels. Letter combinations below the agarose gels represent the respective genes joined by the PCR product. f and ff represent regions between IS1 and the end of fosK. M = DNA size marker.

Fig. 2. The regulatory elements of the intergenic region between fosT and fosR. A. The intergenic region between fosT and fosR. The transcriptional start sites are indicated by bent arrows, the -10 and -35 promoter elements are boxed, and the ribosome binding sites (RBS) are underlined. fosT and fosR start codons are indicated in bold. CRP-cAMP recognition sequence is indicated in italics. Operator sequences O1 and O2 are indicated by inverted arrows. B. Sequence alignment of O1 and O2 with sucrose transcription factor binding sites. Letters in bold corresponds to the LacI family binding sites. cscKBo and cscAo are operator sequences recognized by CscR. The binding site of ScrR was obtained from the RegPrecise database. Nucleotides in italics differ from binding sites of CscR and ScrR. N = any nucleotide; W, A or T. © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

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FosR binds to the IR between fosT and fosR on two operator sequences

Fig. 3. Oligomerization state of FosR. FosR recombinant protein was applied to a Ni-NTA column and treated with glutaraldehyde. Eluates were analysed by Western Blot. 1, molecular weight marker; 2, FosR protein; 3, FosR protein after glutaraldehyde treatment.

with a HTH DNA-binding motif (amino acids 2 to 56, cd01392) and a PBP1_sucrose_transcription_regulator domain (amino acids 63 to 330, cd06288) involved in ligand binding and oligomerization. As members of the LacI/GalR family form homodimers or tetramers, we investigated the oligomerization state of FosR using a cross-linking experiment. As shown in Fig. 3, the fraction treated with glutaraldehyde contains proteins with a molecular weight of approximately 80 kDa, corresponding to FosR dimers (Fig. 3, lane 3). No bands of higher molecular weight were observed on SDS-PAGE and Western Blotting. We then investigated the role of FosR in the regulation of fos gene expression. To measure the activity of the fos promoter, we constructed the plasmid pQF52 carrying the luc gene, which encodes firefly luciferase under the control of the whole IR. This plasmid was then introduced into the BEN2908 and BEN2908DfosR strains, and the expression of luciferase was monitored (Fig. 4A). To ensure that the expression measured depended only on the regulatory elements within the IR, we confirmed that strains carrying plasmid pQF51 (luc gene without the IR) had no luciferase activity (data not shown). In a FOS-free medium poor in carbohydrate sources, such as LB-Miller medium (Sezonov et al., 2007), luciferase expression in strain BEN2908DfosR was up to 51-fold higher than in BEN2908, indicating that the fos-promoter activity is much lower in the wild-type strain than in the fosR mutant. This clearly demonstrates the ability of FosR to repress fos gene expression.

The presence of an HTH motif within FosR strongly suggests that FosR is able to bind to DNA. We therefore performed electrophoretic mobility shift assays (EMSAs) to investigate the ability of FosR to bind to the IR between fosT and fosR. Two DNA shifts were observed when FosR was incubated with a DNA fragment containing the whole IR (Fig. 5A), suggesting that FosR is able to bind to two different sequences of this fragment (Fig. 5B). No shift was observed when an excess of the same unlabelled DNA fragment was added, indicating specific binding of FosR to the IR (Fig. 5B). Moreover, no shift was observed with a DNA fragment containing part of the fosR gene (Fig. 5A and B). The ability of FosR to bind to the IR was further confirmed using surface plasmon resonance (SPR) analysis. As shown in Fig. 5C, interactions between FosR and immobilized DNA were detected. The KD value of interaction between FosR and the IR obtained from three independent experiments was 3.67 nM ⫾ 0.24 (Fig. 5D). Finally, we carried out EMSA and SPR experiments to check the ability of FosR to bind to DNA fragments containing either the putative operator sequence O1 or O2. For these analyses, DNA fragments of the same size as the IR (O1 and O2, Fig. 5A) were used in order to be consistent with the first Biacore experiments using the IR fragment. A DNA shift was observed with the fragments containing both the operator 1 and operator 2 sequences (Fig. 5B). This shift was less marked for O1 and O2 than for the IR fragment because the binding sequences are at the beginning of the fragments. Thus there was less DNA curvature than if the binding had been in the middle of the fragment, explaining the shorter delay. SPR analyses also demonstrated the ability of FosR to bind to these two fragments (Fig. 5C), but with different affinities. Three independent experiments revealed that the KD value for the interaction between FosR and the fragment containing O1 was 9.14 nM ⫾ 1.35, whereas the interaction between FosR and the fragment containing O2 gave a KD value of 1.42 nM ⫾ 0.21 (Fig. 5D). These KD values indicate that the DNA/FosR complexes were relatively stable (Marushima et al., 2009). Finally, these observations demonstrate that FosR directly represses the expression of fos genes by binding specifically to the fos promoter region on two sites, with a stronger affinity for the operator 2 than operator 1.

Fig. 4. Analysis of the fos operon promoter activity. Growth curves (OD450) and relative luminescence intensities (RLU/OD450) of BEN2908, BEN2908DfosR, BEN2908DfosT and BEN2908DfosRDfosT strains carrying pQF52 grown with shaking at 37°C in (A) LB, or M9 minimal medium supplemented with (B) 0.2% scFOS (or 0.01% of glucose + fructose + sucrose, 0.074% of GF2, 0.106% of GF3 and 0.02% of GF4) (C) 5 mM GF2 (D) 0.2% D-glucose (E) 0.2% D-fructose (F) 5 mM GF2 + 0.02% D-glucose (G) 5 mM GF2 + 0.02% D-fructose (H) 0.2% D-glucose + 5 mM cAMP. The RLU average values and standard deviations indicated result from three independent experiments. © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 721

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Fig. 5. Interaction of FosR with the promoter region of the fos operon. A. Schematic representation of fosT, the intergenic region and fosR. The thick solid lines represent the four DNA fragments used in EMSA and SPR experiments: IR (240 bp), O1 (240 bp), O2 (238 bp) and fosR (240 bp). B. EMSAs of FosR binding to the promoter region of fos operon. Labelled DNA fragments were incubated with increasing concentrations of FosR. Some reactions, indicated with a + sign, were performed in the presence of 125-fold excess of unlabelled DNA fragments. C. SPR analysis of FosR binding to the promoter region of fos operon. Biotinylated DNA fragments (IR, O1 and O2) were immobilized on a CM5 sensor-chip. FosR was injected over a range of concentrations [3.125, 6.25, 12.5 (in duplicate), 25, 50 (in duplicate) and 100 nM] at a flow rate of 50 ml min-1, with an association time of 180 s and a dissociation time of 180 s. Black lines represent DNA–protein interactions, from the lowest (at the bottom) to the highest (at the top) concentration of FosR. Red lines represent the global fit of the entire data set to the 1:1 Langmuir binding model. D. The kinetic constants obtained from three independent SPR experiments.

scFOS induce fos gene expression by inhibiting the binding of FosR to the IR Regulators of the LacI/GalR family generally bind lowmolecular weight effectors that abolish their DNA-binding activities (Weickert and Adhya, 1992). To investigate the possible effectors of FosR, we performed EMSA and SPR

experiments to analyse the ability of FosR to bind to the IR in the presence of scFOS. For the EMSAs, the first step allowed FosR to bind to DNA and after that we added carbohydrates to the reaction. The addition of 0.1 mM of GF2, 1 mM of GF3, 3 mM of GF4 and 10 mM of sucrose removed the binding of FosR to the IR (observed by the loss of shift) (Fig. 6A). Binding was not inhibited by the © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 723

Fig. 6. Inhibition of FosR binding to the intergenic region by scFOS. A. EMSAs of FosR binding to the intergenic region. Labelled IR fragment was incubated with 120 nM of FosR with increasing concentrations of the indicated carbohydrates (0, 0.01, 0.03, 0.1, 0.3, 1, 3, 10 and 30 mM). P = DNA fragment without FosR. B. SPR analysis of FosR binding to the intergenic region. Biotinylated IR fragment was immobilized on a CM5 sensor-chip. FosR (200 nM) was pre-incubated with increasing concentrations of the indicated carbohydrates (0.02, 0.05, 0.1, 0.2, 0.5, 1, 2, 5, 10 and 20 mM) and injected at a flow rate of 10 ml min-1, with an association time of 120 s. The percentage of FosR bound to DNA was the ratio of RU levels measured 50 s after the end of the injection with and without carbohydrate.

addition of glucose, fructose, maltose (a disaccharide of glucose) or raffinose (a trisaccharide consisting of galactose, glucose and fructose) (data not shown). As the fosK gene encodes a putative fructokinase, EMSAs were also performed in the presence of D-fructose-1-phosphate and D-fructose-6-phosphate but no inhibition of binding was observed (data not shown). These results were confirmed by SPR analysis using an inhibition binding assay. This involved first pre-incubating 200 nM of FosR with various concentrations of carbohydrates and then measuring the decrease in FosR binding to DNA. We verified that the carbohydrates tested did not bind to DNA at high concentrations (data not shown). As shown in Fig. 6B, Biacore experiments also demonstrated that GF2 was the best

inhibitor of the interaction between FosR and the IR. We also attempted to obtain affinity constants for the interaction between FosR and carbohydrates using the affinity in solution method. In these experiments, a fixed concentration of FosR (200 nM) was mixed with varying concentrations of carbohydrates and left to reach equilibrium. The free concentration of FosR was then determined by injecting the sample containing the FosR-carbohydrate mixture over a ligand that binds to FosR but not to carbohydrate and not to the FosR-carbohydrate complex (i.e., the IR). It can be assumed that the measuring procedure itself did not significantly disturb the equilibrium of the sample mixture. The experimental set-up required a preliminary calibration curve for FosR bound to DNA in order to cal-

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culate the concentration of FosR not bound to carbohydrates, and thus to estimate the affinity constants between FosR and carbohydrates. The plot of free FosR against total carbohydrate concentrations allowed us to estimate a KD value between FosR and GF2 of 23 mM, whereas for GF3 and sucrose it was 154 and 635 mM respectively (Fig. S1). Overall, these results demonstrate that scFOS, and particularly GF2, are effectors that inhibit the binding of FosR to the IR. This inhibition could lead to the expression of fos genes. We therefore investigated fos operon promoter activity in three genetic contexts (strains BEN2908, BEN2908DfosT and BEN2908DfosR) in minimal media containing either scFOS or GF2 alone. The BEN2908DfosT strain is unable to metabolize scFOS, thus we can assume that when this strain stopped growing in the medium containing scFOS (Fig. 4B), all the fructose and glucose in the medium had been consumed and only scFOS remained. As shown in Fig. 4B, induction of luciferase expression in the wild-type strain and in the strain in which the fosR gene had been deleted began when only GF2, GF3 and GF4 remained in the media. This indicates that scFOS removed the repression by FosR, thereby inducing the activity of the fos operon promoter and thus fos gene expression. Moreover, luciferase expression was also induced in BEN2908 and BEN2908DfosR in a medium in which the only carbon source was GF2 (Fig. 4C), indicating that GF2 alone is able to induce fos operon promoter activity. Expression of fos genes is sensitive to cAMP-dependent catabolite repression A putative CRP-cAMP recognition sequence was identified in the IR, suggesting that fos gene expression could depend on catabolite repression. We thus investigated fos operon promoter activity in minimal media containing either glucose or fructose as the sole carbon source. As shown in Figs 4D and 7, the presence of D-glucose strongly repressed expression in strains BEN2908, BEN2908DfosT and BEN2908DfosR, whereas this repression was less marked in the presence of D-fructose (Figs 4E and 7). The activity of the fos operon promoter was up to 2.75-fold higher in BEN2908, 2.84-fold higher in BEN2908DfosR, and 3.81-fold higher in BEN2908DfosT with D-fructose than with D-glucose (Table S2). Moreover, 0.02% of D-glucose in a medium containing GF2 strongly repressed luciferase expression in the three strains until all the glucose had been metabolized (when BEN2908DfosT stopped growing) (Fig. 4F). On the other hand, 0.02% of D-fructose in a medium containing GF2 did not inhibit luciferase expression (Fig. 4G) as demonstrated by the fact that its expression was induced in the presence of D-fructose. These results demonstrate that D-glucose, but not D-fructose, induces catabolite repression of fos gene

Fig. 7. Maximum activity of the fos operon promoter with D-glucose, D-fructose and scFOS. Maximum relative luminescence intensities (RLU/OD450) of strains (A) BEN2908 (B) BEN2908DfosR and (C) BEN2908DfosT (black bars), BEN2908DfosR (white bars) and BEN2908DfosRDfosT (grey bars) carrying pQF52 in M9 minimal medium containing the indicated carbohydrates as the sole carbon source. The RLU average values and standard deviations indicated result from three independent experiments. Asterisks indicate significant differences in mean luciferase expression between two media or two strains revealed by a Student’s t-test. ***P < 0.005; **P < 0.02; *P < 0.05.

expression. It is noteworthy that fos operon promoter activity was strongly repressed in the presence of D-glucose, even in the BEN2908DfosR strain (Figs 4D and 7B). In addition, fos operon promoter activity in BEN2908 and BEN2908DfosR were higher in media containing D-glucose (scFOS, GF2 + glucose) than in those without (GF2 alone, GF2 + fructose) (Fig. 7A and B, Table S2). This suggests that the complete assimilation of D-glucose produces a © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 725

Fig. 8. CRP* induces fos operon promoter activity. Growth curves (OD450) and relative luminescence intensities (RLU/OD450) of CA8306 (in black) and CA8404 (in white) strains carrying pQF52 grown with shaking at 37°C in LB. The RLU average values and standard deviations indicated result from three independent experiments.

signal to increase fos gene expression when FosR is inactive. We therefore investigated whether the presence of cAMP could induce this expression. As shown in Figs 4H, 7A and B, the addition of 5 mM cAMP to a medium containing D-glucose led to a higher luciferase expression in BEN2908 and BEN2908DfosR, than when the medium contained only D-glucose (a 2.31-fold and 6.63-fold increase, respectively) (Table S2). This demonstrates that the expression of fos genes depends on the level of cAMP. Moreover, in BEN2908DfosR, similar and high level of luciferase expression was observed in LB-Miller medium and in a minimal medium containing D-glucose and cAMP (Fig. 4A and H). This maximal activity of the fos promoter in LB-Miller medium could be due to a high intracellular cAMP concentration. To confirm then the ability of CRP-cAMP complex to induce fos gene expression, we introduced the plasmid pQF52 into the CA8306 and CA8404 strains. Both strains can not synthesize cAMP because of a mutation in the cya gene encoding adenylate cyclase. CA8404 is a catabolite repression mutant expressing a cAMPindependent receptor protein mutant (CRP*) whereas CA8306 expresses wild-type CRP. As shown in Fig. 8, luciferase expression in CA8404 was up to 36-fold higher than in CA8306. This clearly demonstrates the ability of CRP*, and thus CRP-cAMP complex, to induce fos gene expression. Influence of D-fructose on fos gene expression in BEN2908DfosR We observed that fos operon promoter activity was induced later in BEN2908DfosR than in BEN2908 in media containing scFOS and GF2 + D-fructose at the early stage of growth (Fig. 4B and G). Luciferase expression was even lower in the former strain than in the wild-type strain in the scFOS medium, and was repressed in a medium containing only D-fructose (Fig. 4E). This observation is surprising, as the absence of FosR should lead to concomitant or earlier fos promoter activity. More-

over, induction rates of fos operon promoter activity were lower in BEN2908DfosR than in BEN2908 in a medium containing scFOS and D-fructose than in another medium (with or without glucose) (Table S2), whereas there was no difference in the induction rate of these two strains between the GF2 + D-glucose and GF2 media. We also observed that in the presence of D-fructose alone, growth of BEN2908DfosT was less than that of BEN2908, which in turn was less than that of BEN2908DfosR (Fig. 4E). This suggests that D-fructose could enter the cell via FosT. We therefore constructed a BEN2908DfosRDfosT strain in which we introduced the pQF52 plasmid to test whether the possible entry of D-fructose via FosT could influence fos promoter activity in media containing D-fructose. As shown in Figs 4B, E and G and 7C, we observed that luciferase expression was always higher in these media in this strain than in BEN2908DfosR. These results were clearly due to the presence of D-fructose, because the luciferase expression of BEN2908DfosRDfosT was no higher than that of BEN2908DfosR in media containing D-glucose (Figs 4D, F and 7C). This demonstrates that possible entry of D-fructose via FosT influences fos gene expression in BEN2908DfosR, probably by another regulator binding to operator sequences.

Discussion Most studies on FOS metabolism have involved the characterization of enzymes such as b-fructofuranosidases, probably because of their technological and industrial interest for the production of prebiotics (Ehrmann et al., 2003; Ryan et al., 2005; Goh et al., 2007). Data concerning the regulation of FOS metabolism are still scarce. It has been shown that the expression of genes involved in FOS metabolism in L. acidophilus, B. breve and L. paracasei are induced by mixtures of scFOS (containing GF2, GF3 and GF4) and repressed by glucose (Barrangou et al., 2003; Ryan et al., 2005; Goh et al., 2007). None of these studies have characterized the transcriptional regu-

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lators present at these loci or identified the sugars with the greatest inducing effect. This study is the first to characterize a regulator of FOS metabolism, and we have shown that fos gene expression depends on the presence of scFOS in the medium and also on GF2 as the sole carbon source. Moreover, we have demonstrated that GF2 is the effector that binds to FosR with the strongest affinity. This binding results in the removal of FosR-DNA-binding activity and consequently leads to the transcription of fos genes. FOS naturally occur in many plants such as wheat. In this plant, which is an important constituent of poultry feed, GF2 is relatively abundant (1.06 mg g-1 dry matter) (Hussein et al., 1998). The fact that fos gene expression depends on the presence of GF2 corroborates the involvement of these genes in the bacterial colonization of the chicken intestine (Schouler et al., 2009). Although the LacI/GalR family has been well described, only a few affinity constants between regulators and their effectors are known. This study determined an affinity constant between FosR and GF2 of 23 mM, which is extremely similar to the 22 mM found between GalR and D-galactose (Chatterjee et al., 1997). However, this value is higher than the KD values between allolactose and LacI (6 mM) (Yildirim and Mackey, 2003) and between IPTG and LacI (1.3 mM) (Spotts et al., 1991). Members of the LacI/GalR family possess a domain allowing oligomerization of the protein, and some, such as LacI, are able to form tetramers, while others, for example GalR, form dimers. In our study, we found that FosR, like GalR, is a dimeric regulator (Fig. 3). Indeed, FosR, like GalR, does not possess a leucine minizipper motif, which has been shown to promote tetramer formation of regulators such as LacI (Weickert and Adhya, 1992). We have also shown that FosR is able to bind to two operator sequences with different affinities at the nM scale, which is also similar to the KD value of 4 nM between GalR and its operator OE (Chatterjee et al., 1997) but higher than the affinity constant of 0.02 nM between LacI and its operator sequence (Spotts et al., 1991). Overall, this suggests that the regulation model of FosR is closer to that of GalR than to that of LacI. To increase transcription repression, LacI/GalR proteins cause DNA looping. Tetrameric regulators can bind to DNA at two operator sites (one per dimer) to generate highly stable loops. Dimeric regulators can bind to two different operator sites, with looping between these sites mediated by protein–protein interactions that can be promoted by DNA bending proteins and/or by DNA supercoiling (Swint-Kruse and Matthews, 2009). In the Gal repressosome, DNA looping is mediated by the binding of the histone-like protein HU to an architecturally critical position on the DNA, via a specific GalR–HU interaction, facilitating GalR–GalR interaction (Kar and Adhya, 2001; Roy et al., 2005; Semsey et al., 2006). As FosR is a dimeric regulator, we can assume that a DNA loop occurs

between the two operator sites O1 and O2 through the binding of a currently unidentified histone-like protein. We have demonstrated in this study that promoter activity of the fos operon is controlled by catabolite repression, as is the case for many promoters regulated by the LacI/ GalR family (Weickert and Adhya, 1992). The overall regulation of catabolite repression in E. coli involves the transcription activator CRP (cyclic AMP receptor protein), the signal metabolite cAMP, adenylate cyclase and EIIAGlc (IIA component of the glucose-specific PTS). In the presence of glucose, EIIAGlc is preferentially dephosphorylated because its phosphate is drained towards the carbohydrate. Phosphorylated EIIAGlc is able to activate adenylate cyclase, and thus cAMP synthesis. Once cAMP has been synthesized, it binds to CRP, and the resulting complex activates the promoters of many catabolic genes and operons (Saier, 1989; Brückner and Titgemeyer, 2002; Deutscher, 2008; Görke and Stülke, 2008). However, new insights indicate that cAMP is not the main contributor to the regulation of the lactose operon (Inada et al., 1996; Narang, 2009). There are also operon-specific mechanisms such as inducer exclusion, responsible for catabolite repression of the lactose operon, whereby EIIAGlc, nonphosphorylated in the presence of glucose, binds to and inactivates LacY, the lactose transporter (only in the presence of lactose) (Osumi and Saier, 1982; Nelson et al., 1983; Deutscher, 2008; Görke and Stülke, 2008). This is an important mechanism that also applies to the transport of other secondary carbon sources such as maltose, melibiose, raffinose and galactose (Saier, 1989; Titgemeyer et al., 1994; Görke and Stülke, 2008). There is no glucose repression in a mutant that lacks the Lac repressor (Inada et al., 1996), whereas there is in the mutant lacking the Fos repressor (Fig. 4D and F). Moreover, the addition of cAMP to a medium containing D-glucose results in increased fos gene expression (Fig. 4H), and fos promoter activity is higher in a strain expressing CRP* compared with a strain expressing wild-type CRP (in absence of cAMP) (Fig. 8). This suggests that, in contrast to what has been described for the lactose operon, the main mechanism leading to the glucose repression of the fos genes is dependent on the cAMP concentration and the binding of CRP-cAMP complex to the fos promoter region. However, further study on scFOS transport by FosT in the presence of D-glucose should indicate a collaborative implication of inducer exclusion in glucose repression of fos genes. In this study, we also observed that fos promoter activity was slightly repressed in the BEN2908DfosR strain, particularly in media containing D-fructose (Fig. 4B, E and G). As the operator sequences O1 and O2 are identical to operator sequences recognized by sucrose repressors ScrR and CscR (Fig. 2B), we checked for a possible role of these regulators in the regulation of fos gene expression. No sucrose PTS system is present in BEN2908 and no © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 727

cscR gene [belonging to the csc sucrose operon (Jahreis et al., 2002)] was found on the BEN2908 genome, indicating that sucrose repressors do not interfere with fos promoter activity. Another mechanism of catabolite repression in enteric bacteria involves the catabolite repressor/ activator (Cra) protein, originally identified as FruR, the LacI/GalR family repressor of fructose metabolism (Ramseier et al., 1995; Saier and Ramseier, 1996). Shimada et al. showed that 178 promoters of genes involved in carbon and energy metabolism are under the control of Cra. Operator sequences of the fos operon promoter possess similarities with the Cra binding site (Fig. S2). Action of the Cra protein is counteracted by inducers such as D-fructose-1-phosphate and D-fructose-1,6bisphosphate, but not by D-fructose (Jahreis and Lengeler, 1993; Saier and Ramseier, 1996; Shimada et al., 2011). Following our observation that the expression of fos genes in BEN2908DfosR was slightly repressed in media containing D-fructose, we investigated the role of this sugar in the expression of these genes. We found that fosT deletion in this strain resulted in higher fos promoter activity in the same media (Fig. 7C). Moreover, the differences in growth of BEN2908 and its derivatives in a medium containing D-fructose suggested that this sugar could enter the cell via both the FosT transporter and the fructose PTS system. One hypothesis explaining these results could be that non-phosphorylated D-fructose entering via FosT would not be able to remove repression induced by Cra. This regulator could thus bind to operator sequences of the fos promoter (unoccupied in the BEN2908DfosR strain) and repress its activity in this strain. In sum, this study enabled us to determine a regulatory model of scFOS metabolism in BEN2908. In the absence of scFOS, FosR could first bind to operator sequence 2 of the IR, repressing fos gene expression, and then bind to operator sequence 1 to repress this expression completely. A DNA loop between these sequences could then form, making the promoter site inaccessible to RNA polymerase. In the presence of GF2, the latter could bind to FosR and undergo a conformational change incompatible with DNA binding, as described for proteins of the LacI/GalR family (Weickert and Adhya, 1992; Swint-Kruse and Matthews, 2009). This would lead to fos gene expression. Moreover, in the presence of scFOS and glucose, EIIAGlc would be non-phosphorylated and would not be able to activate adenylate cyclase and cAMP synthesis. In these conditions, fos gene expression is repressed even in the presence of the effector. After metabolism of all the glucose, EIIAGlc would be phosphorylated and thus activate adenylate cyclase, cAMP synthesis and the formation of cAMPCRP complexes. The binding of this complex to the IR appears to increase fos gene expression. However, both inactivation of FosR (by GF2) and higher intracellular cAMP concentration are necessary for increased fos gene

expression. These data should lead to a better understanding of the environmental conditions required for gene expression enabling prebiotic metabolism in a pathogenic E. coli strain.

Experimental procedures Bacterial strains, plasmids and growth conditions Bacterial strains and plasmids used in this study are listed in Table 1. Escherichia coli strain BEN2908, O2:K1:H5, is a nalidixic acid-resistant derivative of strain MT78, which was isolated from the trachea of a chicken with a respiratory infection (Dho and Lafont, 1982; Germon et al., 2005; Chouikha et al., 2006; Chanteloup et al., 2010). The fosT isogenic mutant of BEN2908 was also used in this study (Schouler et al., 2009), as CA8306 and CA8404 E. coli strains (Brickman et al., 1973; Sabourn and Beckwith, 1975). E. coli strains XL1-Blue and TOP10 were used for cloning and gene expression. Strains were routinely grown in LB-Miller medium at 37°C with agitation. When necessary, antibiotics were used at the following concentrations: nalidixic acid 30 mg ml-1, ampicillin 100 mg ml-1 or kanamycin 50 mg ml-1. For expression analysis using luciferase, overnight LB cultures of strains carrying pQF52 plasmid were washed twice and resuspended in the same volume of M9 minimal medium (Miller, 1972). The strains were then inoculated to an optical density at 450 nm of 0.05 and cultured at 37°C in 25 ml of M9 medium supplemented with either 0.2% scFOS (Profeed P95; Beghin Meiji, France), 0.2% D-glucose (Sigma) ⫾ 5 mM cAMP (Sigma), 0.2% D-fructose (Sigma), 5 mM GF2 (Wako Chemicals GmbH, Germany) ⫾ 0.02% D-glucose, or 0.02% D-fructose. scFOS powder is a mixture containing small quantities of glucose, fructose and saccharose (5%), and larger amounts of GF2 (kestose, 37%), GF3 (nystose, 63%) and GF4 (fructofuranosyl nystose, 10%). Electrocompetent E. coli cells were obtained using Tung and Chow’s method (Tung and Chow, 1995).

Nucleic acid manipulation Restriction enzymes and T4 DNA ligase were used according to the manufacturer’s protocol (New England Biolabs; Promega). Plasmids were purified with Nucleospin Plasmid and Nucleobond PC100 kits and E. coli chromosomal DNA was purified with Nucleospin Tissue kit, according to the manufacturer’s protocols (Macherey-Nagel). PCR products were purified directly or from agarose gels with a Nucleospin Extract II kit (Macherey-Nagel) according to the manufacturer’s protocol. PCRs were performed with Applied Biosystems model 9700 apparatus, using 1 U Taq DNA polymerase (New England Biolabs) in 1 ¥ buffer, 200 mM of each deoxyribonucleoside triphosphate (Ozyme, France) and 1 mM of each primer in a 50 ml reaction volume. The oligonucleotides used in this study are listed in Table S1. For Southern blot and EMSAs, DNA fragments were transferred after electrophoresis to a Hybond-N + membrane (Amersham, GE Healthcare Life Sciences). For Southern blot, probes were labelled with peroxidase, and hybridized

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728 G. Porcheron et al. 䊏

Table 1. Bacterial strains and plasmids. Strain or plasmid E. coli strains BEN2908 BEN2908DfosT::kan BEN2908DfosR BEN2908DfosRDfosT::kan CA8306 CA8404 TOP 10 XL1-Blue Plasmids pKD4 pKD46 pCP20 p3121 pGEM-T easy vector pGEM::RIfosT-RDRBSfosT pGEM::luc pQF50 pQF51 pQF52 pBAD/HisA pBAD/HisA::fosR

Relevant characteristics

Source or reference

Extra-intestinal pathogenic strain; O2:K1:H5; Nalr Fos+ Fim+ Iut+ IbeA+ AGI-3+, avian origin Isogenic deletion mutant of BEN2908; Nalr Kanr FosIsogenic deletion mutant of BEN2908; Nalr Fos+ Isogenic deletion mutant of BEN2908DfosR; Nalr Kanr Fosl -, e14-, relA1, spoT1, DcyaA854, thi-1 l -, e14-, relA1, rpsL136 (Strr), crp-1004, spoT1, DcyaA854, thi-1 mcrA D (mrr-hsdRMS-mcrBC) DlacX74 deoR recA1 araD139D (ara-leu) 7697 galK rpsL endA1 nupG [F80lacZDM15] Strr recA1 endA1 gyrA96 thi-1 hsdR17 supE44 relA1 lac [F′ proAB lacIqZDM15 Tn10 (Tetr)]

Dho and Lafont (1982)

oriRg Ampr Kanr oriR101 repA101(ts) araBp-gam-bet-exo Ampr Ampr Kanr Luc+ Ampr pGEM-T easy containing the intergenic region between fosT and fosR without RBS of fosT pGEM-T easy containing luc gene and its RBS from p3121 oripMB1 oripRO1600 LacZ+ Ampr pQF50 deleted of lacZ gene and containing luc gene and its RBS pQF51 containing the intergenic region between fosT and fosR without RBS of fosT oripBR322 Ampr pBAD/HisA containing the fosR gene

DNA fragments were revealed using an enhanced chemoluminescent kit (RPN 3000, Amersham Pharmacia Biotech) as recommended by the manufacturer. For EMSAs, the membrane was treated with a DIG Gel Shift Kit according to the manufacturer’s protocol (DIG Gel Shift Kit, 2nd Generation, Roche). Finally, hybridized DNAs were revealed by chemiluminescent detection using an image acquisition device (Chemistart 5000, Vilber-Lourmat). For RT-PCR of the fos locus, total RNA was extracted from bacterial cells taken during the mid-exponential phase of growth in M9 medium containing 5 mM GF2 with the RNeasy mini kit (Qiagen) according to the manufacturer’s protocol. To provide immediate stabilization of RNA, 2 volumes of RNA protect (Qiagen) were added to a volume of bacterial cells. After 5 min at room temperature, solution was centrifuged at 5000 g for 15 min and pellets were stored at -80°C till RNA extraction. Residual DNA was removed by on-column DNaseI digestion using the RNase-Free DNase Set according to the manufacturer’s protocol (Qiagen).

Construction of plasmids and mutant strains Expression of recombinant FosR: To construct the pBAD/ HisA::fosR plasmid, the fosR gene was amplified from BEN2908 genomic DNA by PCR using primers GP2 and GP6. The resulting PCR product was digested with HindIII and SacI and inserted into the same restriction sites of pBAD/ HisA. The resulting pBAD/HisA::fosR plasmid was further introduced by electroporation into E. coli TOP10 for protein expression. Construction was verified by sequencing using pBADfor and pBADrev primers.

Schouler et al. (2009) This study This study Brickman et al. (1973) Sabourn and Beckwith (1975) Invitrogen Stratagene

Datsenko and Wanner (2000) Datsenko and Wanner (2000) Datsenko and Wanner (2000) Gerlach et al. (2007) Promega This study This study Farinha and Kropinski (1990) This study This study Invitrogen This study

fos promoter reporter system: To construct pGEM::RIfosTRDRBSfosT, the IR between fosT and fosR without the RBS of fosT was amplified from BEN2908 genomic DNA by PCR using primers GP28 and GP30. The resulting PCR fragment was then inserted into the pGEM-T easy vector (Promega). Recombinant plasmid was then introduced into E. coli XL1Blue by electroporation, and construction was verified by sequencing using primers GP21 and GP22. To construct pGEM::luc, the whole luc gene and its RBS were amplified from plasmid p3121 by PCR using primers GP19 and GP20 (Gerlach et al., 2007). The resulting PCR fragment was then inserted into the pGEM-T easy vector. Recombinant plasmid was then introduced by electroporation into E. coli XL1-Blue, and construction was verified by sequencing using primers GP21 and GP22. To construct pQF51, pGEM::luc was digested with BlpI and HindIII and the resulting restriction fragment (containing luc gene and its RBS) was inserted into the BlpI/HindIIIdigested pQF50 (resulting in the loss of lacZ) (Farinha and Kropinski, 1990). Recombinant plasmid was then introduced into E. coli XL1-Blue by electroporation, and construction was verified by sequencing using primers GP25 and GP26. Finally, recombinant plasmid was introduced by electroporation into BEN2908, BEN2908DfosT, BEN2908DfosR and BEN2908DfosRDfosT. To construct pQF52, pGEM::RIfosT-RDRBSfosT was digested with BamHI and HindIII, and the resulting restriction fragment was inserted into the BamHI/HindIII-digested pQF51, resulting in a plasmid allowing the expression of firefly luciferase under the control of the IR between fosT and fosR of the E. coli strain BEN2908. Recombinant © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 729

plasmid was then introduced into E. coli XL1-Blue by electroporation, and construction was verified by sequencing using primers GP26 and GP27. Finally, recombinant plasmid was introduced by electroporation into BEN2908, BEN2908DfosT, BEN2908DfosR, BEN2908DfosRDfosT, CA8306 and CA8404. fos mutants: To construct the BEN2908DfosR strain, the method developed by Datsenko and Wanner was used (Datsenko and Wanner, 2000). Briefly, using a Red recombination procedure, fosR was deleted and replaced by a kanamycin resistance (Kanr) cassette. The Kanr cassette was obtained by PCR amplification of plasmid pKD4 using primers cat180 and cat181, which contain extensions homologous to the 3′ end of the IR between fosT and fosR and to the 3′ end of fosR. The replacement of fosR was confirmed by PCR and sequencing using the primer pairs cat159/askana1 and skana/cat157, which enable detection of the left and right arms of the insertion respectively. The insertion of the Kanr cassette was also confirmed by Southern blot, using primers cat51 and askana2 to generate a probe of the kanamycin resistance gene from plasmid pKD4. The Kanr cassette was then removed using plasmid pCP20. The deletion of fosR was finally confirmed by PCR and sequencing, using primers cat157 and cat159. The same method was used to construct the BEN2908DfosRDfosT strain. Briefly, fosT was deleted from the BEN2908DfosR strain and replaced by a Kanr cassette. This was obtained by PCR amplification of plasmid pKD4 using primers cat147 and cat148, which contain extensions homologous to the beginning of fosE1 and to the 5′ end of the IR between fosT and fosR. The replacement of fosT was confirmed by PCR and sequencing using the primer pairs GP_45AS/skana and askana1/GP7, which enable detection of the left and right arms of the insertion respectively. The integrity of the IR was also verified by PCR and sequencing using primers GP3 and cat157. The insertion of the Kanr cassette was also confirmed by Southern blot, using primers cat51 and askana2 to generate a probe of the kanamycin resistance gene from plasmid pKD4.

RT-PCR To determine co-transcription within the fos locus, primer pairs connecting the 3′ end of one ORF to the 5′ end of the following one were used. RNA was reverse transcribed using Superscript Reverse Transcriptase III (Invitrogen) and one of the specific reverse primers of interest (GP_41AS2, GP_41AS, GP_42AS, GP_43AS, GP_44AS, GP_45AS, GP_finAS, GP_finAS2, GP_finAS3, GP18). Reaction mixtures contained 1 pmol of primer and 60 ng of RNA in a total volume of 10 ml. Reactions were performed for 60 min at 55°C followed by 15 min at 70°C to inactivate the reverse transcriptase following the manufacturer’s instructions. cDNA was then diluted by adding 20 ml of DEPC-treated water (Ambion) and used for PCR amplification with the reverse primers and appropriate forward primers (GP_41S, GP_42S, GP_43S, GP_44S, GP_45S, GP_46S, GP_46S2, GP17). Control RT-PCRs, omitting reverse transcriptase, were performed to check for DNA contamination of the RNA preparations.

Determination of the transcriptional start site in PfosT and PfosR The transcriptional initiation sites were determined using the 5′ Rapid Amplification of cDNA Ends (RACE) method (Sambrook et al., 1989). The first cDNA strand was synthesized using primer GP_45AS to determine the transcriptional start site of the fos operon, and primer GP18 to determine the transcription start site of the fosR gene. After treatment with 1 U of RNaseH (Invitrogen) for 20 min at 37°C, polyA tails were added to the 3′ ends of the cDNAs. Briefly, cDNAs were first incubated for 3 min at 94°C in a buffer containing 10 mM Tris-HCl pH 8.3, 1.5 mM MgCl2, 50 mM KCl and 0.2 mM dATP, and 80 U of terminal transferase were added to the reaction and then incubated for 30 min at 37°C. The cDNAspolyA were then used as templates in PCR reactions with oligo d(T)-anchor primer (5′/3′ RACE Kit, 2nd Generation, Roche) and either the nested fosT primer GP_46AS2 or the nested fosR primer GP_47AS2. A second PCR reaction was performed using the anchor-primer and the second nested fosT primer GP_46AS3 for PfosT, or the second nested fosR primer GP_47AS3 for PfosR. The products were cloned into pGEM-T easy vectors. Recombinant plasmids were then introduced into E. coli XL1-Blue by electroporation, and the sequences of the cloned fragments were obtained using primers GP21 and GP22 to determine the transcriptional start site.

Expression and purification of recombinant FosR FosR protein was expressed as an N-terminal His6-tagged protein. A 50 ml culture of E. coli TOP10 carrying pBAD/ HisA::fosR was grown in LB-Miller containing ampicillin at 37°C to an optical density at 600 nm of 0.5 and then supplemented with L-Arabinose to a final concentration of 0.002%. After 4 h, cells were harvested, suspended in 1 ml of lysis buffer (50 mM Tris pH 8, 150 mM NaCl, 20 mM imidazole, 1 mM EDTA, 0.1% Triton X-100, 1 mM PMSF, 0.25 mg ml-1 lysozyme) and incubated overnight at -80°C. After thawing, lysate was supplemented with 10 mg ml-1 DNaseI and 20 mM MgSO4, then incubated for 1 h at room temperature with agitation and centrifuged at 14000 g at 4°C for 30 min. Cleared lysate was applied to a nickel-nitrilotriacetic acid (Ni-NTA) column (Qiagen), which was then washed three times with washing buffer (50 mM NaH2PO4, 300 mM NaCl, 50 mM imidazole, pH 8). The His6-tagged FosR protein was finally eluted with elution buffer (50 mM NaH2PO4, 300 mM NaCl, 250 mM imidazole, pH 8) and analysed by SDS-PAGE. Imidazole was then eliminated by dialysis or through a Sephadex G-25 (Interchim) desalting column.

Determination of FosR protein oligomerization state The oligomerization state of FosR protein was determined by glutaraldehyde cross-linking (Fadouloglou et al., 2008). Briefly, 500 mg of FosR protein were diluted in a final volume of 4 ml in buffer A (10 mM imidazole, 50 mM NaCl, 50 mM NaH2PO4, pH 8) and applied to a Ni-NTA column (Qiagen), which was then washed once with buffer A and once with buffer B (50 mM NaCl, 50 mM NaH2PO4, pH 8). Glutaralde-

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730 G. Porcheron et al. 䊏

hyde (Sigma) was then diluted in buffer B to a final concentration of 0.05% and this solution was applied to the column, which was then washed with 0.5 M Tris-HCl pH 8 to stop the reaction. FosR protein was finally eluted with buffer E (300 mM imidazole, 50 mM NaCl, 50 mM NaH2PO4, pH 8) and the oligomerization state was analysed by SDS-PAGE and Western-blotting, using a mouse monoclonal anti-His (Eurogentec) and an anti-mouse IgG conjugated with peroxidase (Sigma). Western blots were revealed using West Pico Chemiluminescent Substrate (Pierce) as recommended by the manufacturer. Chemiluminescence was detected using an image acquisition device (Chemistart 5000, VilberLourmat).

EMSA Four DNA fragments were used for EMSA: (i) IR, the 240 bp IR between fosT and fosR, was amplified by PCR from BEN2908 genomic DNA using primers GP3 and GP7; (ii) O1, the 240 bp fragment between the 3′ end of fosT (nucleotide 166 of fosT) and the IR (nucleotide -49) containing only operator 1, was amplified by PCR from BEN2908 genomic DNA using primers GP8 and GP9; (iii) O2, the 238 bp fragment between the IR (nucleotide -43) and the 5′ end of fosR (nucleotide 65 of fosR) containing only operator 2, was amplified by PCR from BEN2908 genomic DNA using primers GP10 and GP11; (iv) fosR, the negative control, a 240 bp internal fragment of fosR (nucleotides 409–649), was amplified by PCR from BEN2908 genomic DNA using primers GP14 and GP15. PCR products were then labelled at their 3′ end by incubation with terminal transferase and digoxigenin11-dUTP (DIG-11-dUTP) following the manufacturer’s protocol (DIG Gel Shift Kit, 2nd generation, Roche). Binding reactions (20 ml) contained increasing amounts of purified FosR protein (0 to 300 nM), 1 mg poly[d(I-C)], a constant amount of labelled DNA fragment (0.8 ng), and sometimes an unlabelled fragment in excess (100 ng) in a buffer composed of 20 mM HEPES, pH 7.6, 1 mM EDTA, 10 mM (NH4)2SO4, 1 mM DTT, Tween 20, 0.2 % (w/v), 30 mM KCl, and 5 mM MgCl2. After incubation at room temperature for 15 min, reactions were either stopped immediately with 5 ml of loading buffer [0.25 ¥ Tris-Borate-EDTA (TBE) buffer, 60%; glycerol, 40%; bromophenol blue, 0.2% (w/v)], or were first reincubated for 6 min at room temperature with varying concentrations of carbohydrates [GF2, GF3, GF4 (Chemicals GmbH, Germany), D-glucose, D-fructose, maltose, D-raffinose, sucrose, D-fructose-1-phosphate and D-fructose-6-phosphate (Sigma)] and then stopped with loading buffer. Samples were finally separated on a 5% polyacrylamide native gel in TBE 0.25 ¥ at 4°C.

immobilized at 8000–10000 resonance units (RU) on each Flow Cell of a CM5 sensor chip (GE Healthcare), using a standard amine coupling protocol following the manufacturer’s instructions. The different biotinylated double-stranded DNAs were then injected until an immobilization level of ~300 RU (kinetic experiments) to ~600 RU (binding inhibition experiments) was reached. Kinetic analyses were carried out at a flow rate of 50 ml min-1. Concentrations of 3.12–100 nM of FosR protein were injected over the DNA surfaces for 180 s in one or two replicates. Dissociation was studied for 180 s. Regeneration of the surfaces was performed with a 10 s injection of 0.2% SDS followed by a 60 s running buffer wash. Data were corrected for non-specific binding by double referencing with responses from DNA control surface (containing fosR fragment) and from blank injections. Data were fitted to a 1:1 Langmuir interaction model with a local fit for Rmax using Biacore T100 evaluation software (version 2.01). Oligosaccharide binding inhibition experiments were performed using an ‘affinity in solution’ method (Adamczyk et al., 2000; Cochran et al., 2003). 200 nM FosR protein was preincubated with 0.02–20 mM oligosaccharides and this sample was injected over the DNA surface for 120 s at a flow rate of 10 ml min-1. The DNA surface was then regenerated with SDS solution. A 3.12–200 nM FosR protein calibration curve enabled calculation of the free FosR concentration and thus calculation of affinity constants between oligosaccharide and FosR protein using Biacore T100 evaluation software. Determination of affinity in solution provides an alternative to steady-state affinity measurements. The affinity in solution approach is used to determine the free concentration of one interactant in equilibrium mixtures containing known total interactant concentrations.

Luciferase measurements Promoter activities of the fos operon in different media were determined by firefly luciferase expression along the growth curve. Samples of 100 ml were taken every 45 min, and light emission (relative light unit, RLU) was recorded with a luminometer (Lumat LB 9507, Berthold). A luciferase Assay System kit (Promega) was used with some modifications. Briefly, samples were incubated with 300 ml of freshly lysed buffer (1 ¥ CCLR, 1.25 mg ml-1 lysozyme, 2.5 mg ml-1 BSA) for 10 min with agitation at room temperature. Solutions were quick-frozen in liquid nitrogen and then immediately incubated at 37°C. After thawing, samples were incubated for 10 min with agitation at room temperature. Finally, RLU was measured by incubating 25 ml of cell lysate with 50 ml of Luciferase Assay Reagent.

In silico analysis SPR analysis The same four DNA fragments used for EMSA were used for SPR analysis, except for the 240 bp IR between fosT and fosR, which was amplified by PCR from BEN2908 genomic DNA using primers GP16 and GP7. SPR experiments were performed on a Biacore T100 (GE Healthcare) at 25°C, with HBS as running buffer (10 mM HEPES, pH 7.4, 150 mM NaCl, 0.05% Tween 20). Neutravidin (Thermo Scientific) was

To find the rho-independent terminator at the 5′ end of the fos locus, the online TransTerm tool of the Nano+Bio-Center was used (Ermolaeva et al., 2000). FosR domains were predicted with NCBI’s Conserved Domain Database (Marchler-Bauer et al., 2009). Operator sequences 1 and 2 of the IR between fosT and fosR were compared with the transcription factor of the LacI family binding sites using the RegPrecise database (Novichkov et al., 2010). © 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 717–733

Regulation of FOS metabolism in Escherichia coli 731

Acknowledgements This work was funded by the Era-NET PathoGenoMics European program (grant ANR-06-PATHO-002–01), INRA and the Institut Fédératif de Recherche 136 (Agents Transmissibles et Infectiologie). G. Porcheron is a pre-doctoral fellow of INRA (MICA)/Région Centre (France). We are grateful to I. Virlogeux-Payant (INRA, Nouzilly, France) for the gift of the pQF50 plasmid. We thank Josef Deutscher for fruitful discussions. We are grateful to Beghin-Meiji (Neuilly/Seine, France) for its generous donation of Profeed P95.

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