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1 Program in Pathobiology, University of Washington, Seattle, WA, USA
2 Seattle Biomedical Research Institute, Seattle, WA, USA
3 Molecular and Cellular Biology Program, University of Washington, Seattle, WA, USA
4 Department of Microbiology and Immunology, University of Illinois at Chicago, Chicago, IL, USA
Correspondence
Nancy E. Freitag
nfreitag{at}uic.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Present address: Department of Molecular Microbiology, Washington University, St. Louis, MO 63110, USA.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Gray et al., 2006; Leimeister-Wachter et al., 1990; Miner et al., 2008; Pizarro-Cerda & Cossart, 2006; Scortti et al., 2007). PrfA is a 27 kDa protein that recognizes and binds a 14 bp DNA palindrome present in the promoters of its target genes (Freitag et al., 1992; Mengaud et al., 1989). PrfA regulates the expression of gene products required for L. monocytogenes invasion of host cells, intracellular growth, and cell-to-cell spread, and is absolutely essential for bacterial virulence (Freitag, 2006; Scortti et al., 2007).
Based on sequence and structural homology, PrfA has been identified as a member of the Crp/Fnr family of transcriptional activators (Eiting et al., 2005; Korner et al., 2003; Ripio et al., 1997). Proteins within this family generally become activated following the binding of small-molecule cofactors. Crp, for example, undergoes an allosteric change after binding cAMP and becomes a site-specific DNA binding protein that recognizes target promoters and interacts with RNA polymerase (RNAP) (Busby & Ebright, 1999; Kim et al., 1992; Kolb et al., 1993; Lawson et al., 2004). Crp appears to exist in an equilibrium between an active form that efficiently binds DNA target sequences and an inactive form that does not. Cofactor cAMP binding by Crp shifts the equilibrium toward the active form, either by stabilizing this form or by destabilizing the inactive form of the protein (Youn et al., 2007). PrfA may exist in an analogous equilibrium state such that binding of a cofactor is required to shift PrfA to a high-activity form capable of high-affinity DNA binding. Although it is generally believed that a PrfA cofactor exists, this cofactor has not yet been identified.
Mutations in crp have been identified that result in an active form of Crp in the absence of cAMP cofactor (Garges & Adhya, 1985; Harman et al., 1986; Kim et al., 1992; Youn et al., 2006, 2007). Structural and functional studies of these mutants (known as Crp* mutants) have led to the identification of regions of Crp that are important for activity, and it has been observed that Crp* mutants exhibit a conformation that resembles that of wild-type Crp bound to cofactor (Harman et al., 1986). Similar to crp*, several prfA mutations have been identified that appear to result in activation of PrfA in the absence of cofactor (known as prfA* mutants) (Miner et al., 2008; Mueller & Freitag, 2005; Ripio et al., 1997; Shetron-Rama et al., 2003; Vega et al., 2004; Wong & Freitag, 2004). Strains with prfA* mutations express high levels of PrfA-dependent gene products under conditions in which gene expression is usually repressed. The prfA* mutations identified thus far are not functionally equivalent, and significant differences in bacterial virulence have been reported for L. monocytogenes strains containing different prfA* alleles (Miner et al., 2008; Mueller & Freitag, 2005; Scortti et al., 2007; Shetron-Rama et al., 2003). This study describes a biochemical comparison of wild-type PrfA with five different PrfA* mutants (including a novel prfA* mutation) to elucidate the effects of specific amino acid substitutions on distinct aspects of PrfA function.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. L. monocytogenes strains were grown in brain heart infusion (BHI) medium and Escherichia coli strains were grown in Luria broth (LB) at 37 °C with shaking. Strains containing high-activity prfA* alleles such as prfA L140F and prfA G145S have previously proven difficult to construct using standard allelic-exchange techniques (Port & Freitag, 2007; Wong & Freitag, 2004; M. D. Miner and others, unpublished observations). However, a modified approach for generating isogenic mutants was developed and used successfully as follows: prfA L140F and prfA G145S were introduced into L. monocytogenes NF-L1124 containing a transcriptional fusion of gus and neo (Karow & Piggot, 1995) downstream of actA in the bacterial chromosome. Selection for the mutant strains was then facilitated based on an increased level of neomycin resistance in the presence of the prfA* allele, as conferred by the PrfA-dependent promoter actA. In addition, to prevent expression of the introduced prfA* mutations from the plasmid vector used for allelic exchange, prfA coding sequences missing the ATG start codon were amplified by PCR and inserted into the temperature-sensitive plasmid shuttle vector pKSV7 (generating plasmid pNF1147), and the L140F and G145S mutations were then separately introduced via the QuikChange Site-Directed Mutagenesis kit (Stratagene) according to the manufacturer's instructions with the primers listed in Table 2 [generating plasmids pNF1162 (prfA L140F) and pNF1161 (prfA G145S)]. To enrich for mutants containing the prfA L140F or prfA G145S chromosomal replacement, 5 µg neomycin ml–1 was used to select for prfA*-induced neomycin resistance on the final day of allelic exchange at 40 °C. Following allelic exchange, L. monocytogenes strains containing the desired mutations within prfA were confirmed by sequencing of PCR fragments derived from chromosomal DNA.
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) was transformed into chemically competent XL1 Red E. coli hypermutator bacterial cells (Stratagene). Selected transformants were inoculated into LB at 1 : 1000 dilution and grown with shaking to stationary phase at 37 °C. Cultures were repeatedly diluted and grown to stationary phase for a total of 10 cycles. The pNF1019 plasmid was then purified from XL1 Red and introduced via electroporation into conjugation-competent SM10 cells. Transfer of pNF1019 from SM10 into L. monocytogenes 
prfA was carried out as described previously (Wong & Freitag, 2004). Transconjugant prfA mutants exhibiting enhanced actA expression were identified as blue colonies on BHI plates containing 7.5 µg chloramphenicol ml–1, 200 µg streptomycin ml–1, 50 µg 5-bromo-4-chloro-3-indolyl-β-D-glucuronic acid (X-gluc) ml–1 and 5 µg neomycin ml–1.
Generation and purification of recombinant PrfA* proteins.
DNA fragments containing prfA and prfA* ORFs were amplified using PCR of L. monocytogenes genomic DNA isolated from NF-L1124 (prfA WT), NF-L1177 (prfA G145S), NF-L1166 (prfA L140F), NF-L1214 (prfA Y63C), NF-L1213 (prfA Y154C) or NF-L924 (prfA E77K) using primers listed in Table 2
. The PCR fragments were then cloned into pET100 using a Champion pET Directional TOPO Expression kit (Invitrogen) as per the manufacturer's instructions. Plasmids containing the prfA and prfA* ORFs were transformed into BL21 Star (DE3) expression cells and PrfA/PrfA* protein production was induced by addition of 1 mM IPTG for 1.5 h. Protein extracts containing recombinant PrfA/PrfA* proteins were passed over a nickel-affinity column, and PrfA was eluted with 200–500 mM imidazole buffer and dialysed into PBS (137 mM NaCl, 10 mM potassium phosphate, 2.7 mM KCl, pH 7.4) with 10 % glycerol (v/v). Purified protein was visualized and assessed for purity following separation on SDS-PAGE gels and Coomassie staining, and also confirmed by Western blot analysis using an anti-PrfA polyclonal antibody (Greene & Freitag, 2003).
Limited proteolysis.
One microgram of purified wild-type PrfA and each PrfA mutant was incubated with 300 ng trypsin (Sigma) or 250 ng subtilisin (Sigma) in Sigma 10x Multicore buffer for the indicated times at 37 °C. Reactions were terminated by the addition of 1 µl PMSF, and samples were then boiled for 5 min, run on 12 % acrylamide gels in MES Buffer (50 mM MES, 50 mM Tris base, 0.1 % SDS, 1 mM EDTA, pH 7.3; Invitrogen) for small-band separation, and visualized by Coomassie stain.
Electrophoretic mobility shift assays (EMSAs).
Primers used to PCR-amplify DNA fragments (
100 bp) containing the hly and actA promoters from L. monocytogenes genomic DNA are listed in Table 2. Primers labelled with cy5.5 label on the 5' ends were purchased from Operon Biotechnologies. To generate a DNA fragment for use as a non-specific competitor for DNA binding assays, primers were used to amplify the prfA ORF (370 bp), as this region lacks PrfA binding sites (primer sequences listed in Table 2). Extracts from NF-L890 (prfA) were made as follows: bacteria from 1 l cultures grown to mid-exponential phase in BHI were collected using centrifugation for 10 min at 6000 g and resuspended in 20 ml ice-cold PBS, and bacterial cells were disrupted by a triple passage through a French press. EMSA were performed as described elsewhere (Böckmann et al., 2000). EMSA reaction mixtures consisted of the following: 40 ng labelled DNA probe (hlycy5.5 or actAcy5.5), PrfA protein (as indicated), 1 µg BSA ml–1, and 50 mM dIdC in TB buffer [10 mM Tris-HCl, pH 8, 10 mM MgCl2, 5 mM CaCl2, 1 mM EDTA, 0.2 mM DTT, 10 % glycerol (v/v)] in a final reaction volume of 20 µl. For experiments including cell extracts, 1 µl (3 µg) extract was added to each reaction mixture. Sample reactions containing all components except labelled DNA were incubated for 15 min at room temperature. The labelled DNA probe was then added and samples were incubated for 3 min at 37 °C, followed by 27 min on ice. Samples were then loaded onto a 5 % acrylamide gel (0.5xTris-boric acid-EDTA) and run at a constant current of 20 mA for approximately 3 h in the dark at 4 °C. Gels were then visualized as in-gel Western blots using the Odyssey Imager (Li-Cor Biosciences) with the cy5.5-labelled fluorescent probes visualized at 700 nm. The His- and Express-tags were found to have minimal effects on PrfA and PrfA* protein function in comparison to purified PrfA protein without the tags (M. D. Miner, unpublished data), as has been reported elsewhere (Böckmann et al., 1996, 2000).
Measurement of β-glucuronidase (GUS) activity.
GUS activity was measured as previously described (Shetron-Rama et al., 2003) with minor changes. Briefly, L. monocytogenes cultures grown overnight at 37 °C in BHI were diluted 1 : 50 and grown with shaking at 37 °C for 8 h. OD600 was measured for each time point and two 500 µl culture aliquots were collected for all strains, except for the prfA L140F (NF-L1166), prfA G145S (NF-L1177) and prfA Y63C (NF-L1214) mutant strains, for which two 50 µl aliquots were collected (reflective of the increased GUS activity present in these three highly activated prfA* strains). Bacterial cells were recovered by microcentrifugation and the supernatants were removed. Bacterial pellets were resuspended in 100 µl (aliquot 1) or 1 ml (aliquot 2) ABT buffer (0.1 M potassium phosphate, pH 7.0, 0.1 M NaCl, 0.1 % Triton). GUS activity was measured as described elsewhere, with the substitution of 4-methylumbelliferyl-β-D-glucuronide in place of 4-methylumbilliferyl-β-D-galactoside (Sigma) (Youngman, 1987). Data were derived from duplicate samples taken from three independent experiments.
Measurement of haemolytic activity.
Haemolytic activity was measured as described elsewhere, with minor modifications (Camilli et al., 1989). Briefly, bacteria were grown without shaking overnight in BHI at 30 °C, the bacterial supernatants were recovered following centrifugation, and twofold serial dilutions of the supernatants were incubated with PBS-washed sheep red blood cells (RBCs; 0.3–10 %) for 30 min at 37 °C. After incubation, RBCs were recovered by centrifugation to measure 50 % lysis and supernatants were read in a spectrophotometer plate reader at A450.
Protein chemical cross-linking.
Purified proteins (500 ng) were incubated with either 10 µM sulfo-ethylene glycol bis[succinimidylsuccinate] (S-EGS) or bis[sulfosuccinimidyl] suberate (BS3) in 0.2 M triethylamine (TEA), pH 8.0, for 1 h at room temperature. Samples were then heated at 85 °C for 10 min in SDS sample buffer (1 % SDS, 10 % glycerol, v/v, 10 mM Tris-Cl, pH 6.8, 1 mM EDTA, and 0.05 mg ml–1 bromphenol blue dye) containing 5 % β-mercaptoethanol, run on SDS-PAGE and transferred to nitrocellulose. Rabbit polyclonal antibody directed against PrfA was used for Western blot analysis at 1 : 4000 dilution followed by incubation with goat-anti-rabbit-IRDye 680 at 1 : 10 000 (Li-cor Biosciences). Membranes were visualized on an Odyssey Imager (Li-cor Biosciences).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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prfA strain. The actA-gus-neo-plcB transcriptional fusion within the L. monocytogenes chromosome was used for the identification of prfA* mutations based on the enhanced expression of neomycin resistance and blue colony colour in the presence of the GUS substrate X-gluc on indicator plates. Plasmid pPL2-prfA containing a copy of wild-type prfA and its promoters was propagated in the E. coli mutator strain XL1 Red as described in Methods, and then transformed into conjugation-competent E. coli SM10 cells for conjugal transfer into prfA/actA-gus-neo-plcB L. monocytogenes. Transconjugants with prfA* mutations were selected based on enhanced neomycin resistance and dark-blue colony colour on plates containing neomycin and X-gluc. Out of approximately 40 000 transconjugants screened, two mutants were identified with approximately 200-fold and 185-fold higher levels of actA expression (based on GUS activity in broth culture) in comparison with the wild-type prfA strain. DNA sequencing of the mutant prfA alleles revealed a leucine to phenylalanine substitution at amino acid position 140 [prfA L140F, a previously described mutation (Wong & Freitag, 2004)], and a novel tyrosine to cysteine amino acid substitution at residue 63 (PrfA Y63C). The position of the Y63C substitution with respect to the 3D structure of PrfA and in relation to other identified prfA* mutations is shown in Fig. 1.
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; Wong & Freitag, 2004; M. D. Miner and others, unpublished observations). However, the use of the NF-L1124 strain containing the actA-gus-neo-plcB transcriptional fusion enabled selection of isogenic chromosomal mutants using enhanced neo expression to select for prfA* alleles (see Methods). As a result, it was possible to directly compare in vitro expression of PrfA-dependent genes and gene products for all five prfA* strains along with the wild-type parent strain. For the actA gene product, wild-type L. monocytogenes is known to express low-to-undetectable levels of actA during growth in BHI broth culture, whereas expression levels increase over 200-fold upon PrfA activation in the host cell cytosol (Bubert et al., 1999; Freitag & Jacobs, 1999; Moors et al., 1999; Shetron-Rama et al., 2002). High-level actA expression in broth culture was observed for L. monocytogenes strains containing the prfA G145S, prfA L140F and prfA Y63C alleles (200-fold higher than the levels expressed by strains containing wild-type prfA) (Fig. 2a); in comparison, moderate actA expression was observed for the prfA E77K and prfA Y154C mutant strains (10-fold increase in expression over wild-type).
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). Consistent with the observed increases in actA expression, the prfA G145S, prfA L140F and prfA Y63C strains exhibited the highest levels of LLO activity, followed by prfA E77K and prfA Y154C (Fig. 2b). Taken together, these data indicate a hierarchy of prfA* mutant activity based on the patterns of PrfA-dependent gene expression: high-activity mutants (prfA G145S, prfA L140F and prfA Y63C) and moderate-activity mutants (prfA E77K and prfA Y154C).
PrfA* mutants are conformationally distinct from the wild-type protein
Limited proteolytic digestion of proteins serves as a useful tool for rapid detection of protein conformational changes, and it has been used to distinguish between active and inactive forms of Crp (Harman et al., 1986; Tan et al., 1991). Limited protease digestion of Crp* mutants results in cleavage patterns that resemble those observed for Crp when bound to cAMP (Harman et al., 1986; Tan et al., 1991). To detect any conformational alterations associated with PrfA* mutations, each mutant protein was purified and subjected to limited trypsin digestion (Fig. 3). PrfA G145S protein served as a positive control for the assay, as a conformational change in this protein has already been demonstrated by crystal structure analysis (Eiting et al., 2005). As anticipated, PrfA G145S was found to exhibit enhanced susceptibility to protease digestion in comparison with wild-type PrfA (Fig. 3). Similar to PrfA G145S, PrfA Y63C and PrfA E77K exhibited similar patterns of enhanced susceptibility to proteolysis. Interestingly, the highly activated PrfA L140F did not exhibit enhanced susceptibility to proteolysis, but the substitution of phenylalanine for leucine in this mutant occurs adjacent to a trypsin cleavage site, and may thus interfere with protease digestion. These results strongly suggest that the presence of the prfA* mutations alters PrfA conformation.
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; Velge et al., 2007), and it was recently shown that wild-type PrfA (27 kDa) migrates in SDS-PAGE gels at a molecular mass of approximately 60 kDa following incubation with a chemical cross-linking agent (Velge et al., 2007). To assess the ability of the different PrfA* proteins to form dimers, purified proteins were incubated with two distinct chemical cross-linking agents and analysed on SDS-polyacrylamide gels. Two cross-linkers were used: S-EGS and BS3, both of which react with free amine groups but differ in the lengths of the linker arms [16 Å (1.6 nm) for S-EGS and 11 Å (1.1 nm) for BS3]. Interestingly, wild-type PrfA was found to form dimers more readily than any of the PrfA* mutants, with the ratio of dimer to monomer for S-EGS and BS3 being 0.27 and 0.44 respectively (Fig. 4). The moderately active PrfA Y154C and PrfA E77K showed reduced dimer formation in comparison to wild-type, with dimer to monomer ratios of approximately 0.05 for S-EGS and 0.11 for BS3 (Fig. 4). Unexpectedly, the most highly active PrfA* mutants PrfA G145S, PrfA L140F and PrfA Y63C exhibited the lowest ratio of dimer to monomer following chemical cross-linking (Fig. 4).
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; Vega et al., 2004). Subsequent to these studies, Eiting et al. (2005) demonstrated by surface plasmon resonance that PrfA G145S binds to the hly promoter with an 18-fold higher affinity than wild-type PrfA. Consistent with these observations, both of the highly activated PrfA* mutants PrfA G145S and PrfA L140F were observed to readily bind DNA fragments containing the hly promoter (Fig. 5). Wild-type PrfA was found to bind only weakly to the hly promoter, with detectable binding observed in the presence of 2 µg PrfA protein and no binding evident in the presence of 500 ng protein (Fig. 5). In contrast, the highly active PrfA* mutants PrfA G145S and PrfA L140F bound target DNA with very high affinity, such that all available DNA was bound at low protein concentrations (500 ng) (Fig. 5). PrfA Y63C, PrfA Y154C and PrfA E77K also demonstrated higher-affinity DNA binding in comparison with wild-type protein, but to a lesser degree than either PrfA G145S or PrfA L140F (Fig. 5). Binding in all cases was specific, as the addition of cold specific competitor DNA but not non-specific competitor DNA eliminated the PrfA-dependent mobility shift (M. D. Miner, unpublished data).
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). Consistent with the promoter preference previously observed for the wild-type protein, the mutant PrfA proteins were observed to bind the actA promoter with a lower apparent affinity than the hly promoter; however, the relative binding affinity hierarchy for the proteins was similar to that observed for hly DNA (Fig. 5). Binding was detectable with significantly lower amounts of both PrfA G145S and PrfA L140F protein in comparison to wild-type PrfA.
Examination of PrfA*–DNA–RNA polymerase complex formation in bacterial cell extracts
Activation of target gene expression requires both binding of PrfA to target promoter sites and recruitment of RNAP. To examine the ability of PrfA* mutants to form complexes with target promoter DNA fragments and RNAP, purified PrfA and PrfA* mutant proteins were incubated with DNA in the presence of cell extracts derived from an L. monocytogenes prfA deletion strain. As mentioned above for purified protein incubated with DNA, wild-type PrfA exhibited weak binding of the hly promoter in comparison with PrfA* mutants (Eiting et al., 2005; Mauder et al., 2006; Vega et al., 2004) (Fig. 6a, CIII complexes). However, in the presence of bacterial cell extracts, wild-type PrfA formed DNA–RNAP complexes with an affinity apparently equal to that of the PrfA* proteins (Fig. 6a, CI complexes). These results suggest that PrfA binding to the hly promoter is enhanced by the presence of RNAP and/or other components within bacterial cell extracts. The PrfA* mutants appeared to form PrfA–DNA–RNAP complexes that were roughly equivalent in amount to those formed using wild-type PrfA (Fig. 6a, CI complexes), and the absence of visible CII bands (RNAP–DNA complexes) suggests that RNAP is limiting under these assay conditions. Similar results were observed with the actA promoter (Fig. 6b).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Miner et al., 2008; Mueller & Freitag, 2005; Ripio et al., 1997; Scortti et al., 2007; Shetron-Rama et al., 2003; Vega et al., 2004; Wong & Freitag, 2004). This study represents, to our knowledge, the first biochemical comparison of multiple PrfA* mutant proteins with differing levels of activation. Our results suggest that prfA* mutations have the capacity to activate PrfA via a variety of structural and functional modifications.
Overall, the expression levels of PrfA-dependent gene products in vitro appeared in this study to correlate most strongly with the binding affinity of PrfA for target DNA (Fig. 5). prfA* mutations that conferred the highest levels of PrfA-dependent gene expression in vitro exhibited the highest affinity of DNA binding as detected by EMSA, with moderately active prfA mutant alleles correspondingly exhibiting more moderate increases in DNA binding. Interestingly, although wild-type PrfA formed very low amounts of protein–DNA complexes with either the hly or actA promoter fragments in comparison with PrfA* proteins (Fig. 5), PrfA–RNAP–DNA complexes were readily formed for both promoter fragments with RNAP present in cell extracts (Fig. 6). Earlier studies by Mauder et al. (2006) have suggested that the efficiency of binding of PrfA to its binding site alone (CIII formation) does not necessarily indicate its potential to initiate transcription at a PrfA-dependent promoter. Their conclusions were based on in vitro transcription assays using purified PrfA proteins (including PrfA G145S) and partially purified RNAP with linear DNA templates. However, substantially less PrfA G145S is required to form either PrfA–DNA or PrfA–RNAP–DNA complexes than wild-type PrfA (Mauder et al., 2006; Fig. 5); thus, it seems reasonable to speculate that under conditions in which PrfA concentrations are limiting, activated PrfA or PrfA* mutants with increased DNA binding affinity would be better able to stimulate the formation of active transcription complexes with RNAP.
Mutations that enhance PrfA-dependent gene expression in vitro have been isolated in multiple regions of the protein (Fig. 1). PrfA G145S and PrfA L140F map within the 
D -helix of PrfA, with G145S positioned near the centre of the helix and L140F located at one end (Eiting et al., 2005; Fig. 1). The mutations are positioned near what corresponds to the hinge region of Crp, a region believed to mediate communication between the C- and N-terminal domains of the protein (Garges & Adhya, 1985; Harman et al., 1986; Kim et al., 1992; Youn et al., 2006). Selected mutations in the Crp hinge region lead to the constitutive activation of Crp in the absence of cAMP via a change in secondary structure that enhances the solvent exposure of the DNA binding helix. Eiting et al. (2005) reported a similar structural change that occurs in PrfA G145S mutants. Based on the functional similarities of the L140F mutant with G145S, most notably a large increase in DNA binding affinity, the PrfA L140F mutation may mediate a similar structural change. While the conformational changes imparted by the L140F mutation as detected by limited proteolysis indicated that the PrfA L140F protease susceptibility was most similar to that of the wild-type protein (Fig. 3), this result may be misleading, as the L140F mutation is located near a trypsin cleavage site (K139), which could influence the efficiency of trypsin cleavage at this position.
Other prfA* mutations with the potential for distinct structural influences include the PrfA E77K, Y63C and Y154C mutations. The E77K mutation lies between β6 and β7 in a region near the central C helices (Eiting et al., 2005; Fig. 1). This mutation enhanced PrfA DNA binding to a lesser extent than that of the G145S and L140F mutations, which suggests either that E77K has a more modest effect on the repositioning of the central C helices or that E77K enhances PrfA-dependent gene expression through a different mechanism. The E77K mutation is located near a region of PrfA that corresponds to an area of Crp and CooA known to interact with RNAP (AR2) (Leduc et al., 2001; Niu et al., 1996). AR2 comprises a patch of positively charged residues that contact an acidic patch on the N-terminal domain of the -subunit of RNAP (-NTD). As the PrfA E77K substitution adds a positively charged lysine residue within a similarly located potential AR2 region, it is possible that the additional positive charge enhances PrfA interactions with RNAP.
Y154C and Y63C map within regions of PrfA (D and β5, respectively) that are associated with a structural tunnel that may serve as a binding pocket for PrfA co-factor (Eiting et al., 2005). Y154C is located at the very end of the D helix, whereas Y63C is located within the β5 domain (Fig. 1). Despite the similar chemical nature of the substitutions, these mutations have dramatically different effects on PrfA function. The Y154C mutation slightly enhanced PrfA-dependent gene expression in broth culture and exhibited a modest but reproducible increase in apparent DNA binding affinity (Fig. 5). Interestingly, this mutation impedes PrfA-dependent gene expression in cytosolic L. monocytogenes, suggesting that the Y154C mutation may interfere with the shift of PrfA to a fully activated state (Miner et al., 2008). In contrast, Y63C dramatically increased PrfA-dependent gene expression in broth culture but did not dramatically increase DNA binding affinity (Figs 2 and 5). Several possibilities exist that could account for the effects of these mutations on PrfA function. The mutations could: (1) inhibit (Y154C) or enhance (Y63C) PrfA cofactor binding; (2) stabilize the low- (Y154C) or high- (Y63C) activity form of PrfA; or (3) result in the formation of disulfide bridges that serve to lock PrfA in either a low-activity (Y154C) or a high-activity (Y63C) state. Although we cannot differentiate between these possibilities at this time, we favour the Y63C mutation enhancing cofactor binding for the simple reason that no significant increase in DNA binding was observed for this mutant in vitro, suggesting that its high activity is not due to increased accessibility of the PrfA DNA binding helix-turn domain.
The apparent inverse correlation that was found to exist between the ability of the PrfA* mutants to form dimers and their ability to activate gene expression was unexpected. Crp has long been known to form dimers as an active transcription factor, and Fnr is believed to form dimers when active and to be monomeric when inactive (Lazazzera et al., 1993). While the chemical cross-linking experiments presented here suggest that PrfA dimerization inversely correlates with DNA binding and activation of target gene expression, the cross-linking agents used were specifically reactive for amine groups and it is possible that these moieties are less available as a result of conformational changes resulting from the prfA* mutations.
Multiple prfA* mutations have been isolated in L. monocytogenes using a variety of approaches (Miner et al., 2008; Shetron-Rama et al., 2003; Vega et al., 2004), and the reconstruction of these mutations in isogenic backgrounds has been highly desirable for unambiguous comparison of the effects of the prfA* mutations on L. monocytogenes physiology and pathogenesis. While the moderately active prfA* alleles have been easily introduced into isogenic strains using allelic exchange (Miner et al., 2008; Shetron-Rama et al., 2003; Vega et al., 2004), this approach has not proven feasible for the higher-activity prfA* mutants prfA G145S and prfA L140F (Port & Freitag, 2007; Wong & Freitag, 2004; M. D. Miner, unpublished observations). The prfA G145S and prfA L140F mutations appear to confer a subtle fitness defect upon L. monocytogenes that is not evident in pure cultures of bacteria but which can be detected in mixed cultures when the mutant strains are grown in the presence of wild-type bacteria (J. Bruno and N. E. Freitag, unpublished data). A fitness defect has also been reported for high-activity crp* mutants (Youn et al., 2006). To our knowledge, until now, the prfA G145S had never been reintroduced into its correct chromosomal location by allelic exchange in any L. monocytogenes strain, including EGD and 10403S. This work therefore represents a novel method enabling the reconstruction of prfA* isogenic strains with highly active prfA* mutations without the use of plasmids and with prfA* in the proper chromosomal location.
In summary, prfA* mutations appear to activate PrfA through a variety of structural and functional modifications. In general, the prfA* mutations that most dramatically enhanced the binding of PrfA to its DNA recognition sequences resulted in the highest levels of PrfA-dependent gene expression in bacterial cultures. Surprisingly, an inverse correlation appears to exist between the level of PrfA activation conferred by a prfA* mutation and the ability of the purified mutant protein to form a dimer. Future studies focused on 3D structural analyses of the mutant proteins will help to further clarify the influences of individual prfA* mutations on PrfA activation.
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ACKNOWLEDGEMENTS |
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prfA strain, and members of the Freitag laboratory for helpful discussions. This work was supported by Public Health Service grants AI41816 (N. E. F.) from NIAID, by a NIAID Bacterial Pathogenesis training grant fellowship AI55396 (M. D. M.), a National Science Foundation Graduate Research Fellowship (NSF-GRF) (G. C. P.), and by the M. J. Murdock Trust. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of the funding sources. Edited by: H. Ingmer
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REFERENCES |
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Received 2 June 2008;
revised 21 August 2008;
accepted 23 August 2008.
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