1 Department of Genetics, University of Georgia, Athens, GA 30602, USA
2 Seattle Biomedical Research Institute, 4 Nickerson St, Seattle, WA 98109, USA
3 Department of Pathobiology and Department of Microbiology, University of Washington, Seattle, WA 98195, USA
Correspondence
Nancy E. Freitag
nancy.freitag{at}sbri.org
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ABSTRACT |
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Present address: University of Wisconsin Rock County, 2909 Kellogg Ave, Janesville, WI 53546, USA.
Present address: Elitra Pharmaceuticals, 3510 Dunhill St, San Diego, CA 92121, USA.
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INTRODUCTION |
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PrfA is the only definitive regulator of virulence gene expression identified thus far in L. monocytogenes. PrfA-mediated activation requires binding of the PrfA protein at conserved 14 bp sequences of dyad symmetry found in target promoters (Chakraborty et al., 1992; Freitag et al., 1992
, 1993
; Leimeister-Wachter et al., 1990
; Mengaud et al., 1991
), a characteristic shared by the cyclic AMP (cAMP) receptor protein (CRP)-recognition elements (Ripio et al., 1997b
). The primary structure of PrfA has limited but significant similarities (approx. 20 % amino acid identity, 30 % similarity) to CRP and other members of the CRP-FNR family of transcription factors (Kreft et al., 1995
; Lampidis et al., 1994
). Fundamental functional similarity of the two proteins is most strongly indicated by the properties of a specific PrfA mutant resulting in a glycine to serine substitution at amino acid position 145. PrfA-dependent genes are constitutively overexpressed in this mutant even under environmental conditions that normally down-regulate expression of virulence genes (Behari & Youngman, 1998a
; Ripio et al., 1996
, 1997a
, b
). An analogous mutation in CRP leads to a cAMP-independent, constitutively active CRP protein. Thus, it has been proposed that PrfA may require a similar co-factor or some form of post-translational modification for efficient binding and activation of its target promoters (Ripio et al., 1997b
; Vega et al., 1998
).
Several recent studies have contributed to a better understanding of the mechanisms that mediate regulation of genes under PrfA control (Bockmann et al., 2000; Dickneite et al., 1998
; Herler et al., 2001
; Lalic-Multhaler et al., 2001
; Renzoni et al., 1997
, 1999
; Shetron-Rama et al., 2002
; Williams et al., 2000
). Park & Kroll (1993)
originally reported that the disaccharide cellobiose is the only one of many carbohydrates tested that has a repressive effect on the expression of two virulence genes, hly and plcA, in L. monocytogenes strain NCTC 7873. These results led to the proposal that cellobiose functions as a specific signature molecule providing L. monocytogenes bacteria with a mechanism for sensing their environment (Park & Kroll, 1993
). However, it was subsequently demonstrated in experiments utilizing three other L. monocytogenes wild-type isolates that cellobiose is not unique in its repressive effect (Milenbachs et al., 1997
). In these isolates, growth in the presence of several other readily metabolized sugars was found to significantly down-regulate virulence gene expression. PrfA protein levels were unaffected by growth in the presence of these sugars, suggesting either that the activity of PrfA may be subject to regulatory modulation (e.g. via a covalent modification or allosteric interaction) or that some additional unidentified regulatory factor may be involved (Milenbachs et al., 1997
; Renzoni et al., 1997
).
Recently, gene products encoded by the bvrABC locus have been implicated in the repression of L. monocytogenes virulence gene expression in response to the -glucosides cellobiose and salicin (Brehm et al., 1999
). The bvrABC locus encodes an anti-terminator of the BglG family (bvrA), a
-glucoside-specific enzyme II permease component of the phosphoenolpyruvate-sugar phosphotransferase system (PTS) (bvrB) and a putative ADP-ribosylglycohydrolase (bvrC). The mechanisms mediating BvrABC-dependent repression of virulence gene expression in the presence of cellobiose have not been elucidated, but mutations within this locus do not prevent utilization of the sugar by L. monocytogenes. A second locus has also been implicated in cellobiose-mediated repression of hly expression, but the gene products that mediate repression were not well defined (Huillet et al., 1999
).
In this work, we identify a novel locus involved in cellobiose-dependent repression of virulence gene expression in L. monocytogenes and also provide genetic evidence for regulatory factors in L. monocytogenes other than PrfA that are involved in virulence gene regulation in response to diverse environmental stimuli. We have characterized two independent ethyl methanesulfonate (EMS)-generated mutations, both unlinked to the prfA gene, which result in deregulation of virulence gene expression. One mutation deregulates hly expression in the presence of several repressing sugars and under environmental conditions that normally down-regulate hly, including low temperature. The second mutation alleviates repression by cellobiose via the truncation of a putative multi-domain regulatory protein with homology to Bacillus subtilis LevR, which regulates bacterial utilization of levans (polymers of fructose) (Debarbouille et al., 1991). Complete relief of cellobiose repression in L. monocytogenes requires both mutations. However, even in the double mutant, regulation of the catabolite control protein (CcpA)-controlled enzyme
-glucosidase remains intact. These results are consistent with a model in which cellobiose acts through at least two semi-independent pathways to repress virulence gene expression. The results also support the earlier proposal by Park & Kroll (1993)
that cellobiose may play a role in virulence gene regulation distinct from that of other readily metabolized sugars. Either or both of these hypothesized pathways could act through a co-factor that modulates PrfA activity.
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METHODS |
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Construction of pCON1-HGNG and hlygusA transcriptional fusions.
The E. coliL. monocytogenes shuttle vector pCON1-HGNH, used to create a stable hlygus transcriptional fusion in L. monocytogenes 10403S, was constructed as follows. An internal fragment of the hly gene was PCR-amplified from the 10403S chromosome with the oligonucleotides AAMhly3250R (5'-CAGTGGATCCCAATTAATTGCGAAATTTGG-3') and AAMhly3762L (5'-ACTCCTGGTGTTTGTCGACTAAAAGTAGCG-3'). Each primer contains two mismatches, indicated by bold type, which create BamHI and SalI sites (underlined), respectively. This 512 bp fragment was digested with BamHI and SalI and ligated into the vector pMLK117 (Karow & Piggot, 1995) which contains a promoterless copy of the gusA gene (encoding
-glucuronidase) from E. coli and a neomycin-resistance cassette, creating the vector pMLK117-hly. This vector was digested with BamHI and HindIII and the resulting fragment (containing the hly fragment, the promoterless gus gene and the neomycin cassette) was ligated into BamHI-/HindIII-digested pCON1 (Freitag, 2000
), creating pCON1-HGN. Propagation of pCON-1 in Gram-positive bacteria depends upon a temperature-sensitive origin of replication derived from pE194ts, and it carries a chloramphenicol-resistance (Cmr) marker for selection in Gram-positive bacteria. A fragment spanning the intergenic region between the hly and mpl gene was PCR-amplified from the 10403S chromosome with primers AAMhly4143R (5'-CCATCTGGGGCACCACGCTTTATCC-3') and AAMmpl4670L (5'-TTAAATCAGCAGGCGCCTTTTTGGC-3'). AAMmpl4670L contains two mismatches in its sequence, indicated in bold type, which create a NarI site (underlined). This 527 bp fragment was digested with HindIII and NarI and ligated into HindIII-/-NarI-digested pCON1-HGN, creating pCON1-HGNH. pCON1-HGNH was transformed into the E. coli donor strain S-17-1 and conjugated into L. monocytogenes 10403S as described previously (Behari & Youngman, 1998a
). 10403S(pCON1-HGNH) was initially grown at 30 °C until mid-exponential phase and then shifted to the non-permissive temperature (42 °C) with selection for Cmr. These growth conditions selected for the homologous recombination of pCON1-HGNH into the 10403S chromosome at the hly locus. Resultant clones were Cmr, Nmr and non-haemolytic on blood agar plates (Hly-). To select for cells where spontaneous excision of the undesired pCON1 sequences had occurred, several isolates were diluted 1 : 1000 from overnight cultures into BHI medium containing 5 µg neomycin ml-1 and grown at the permissive temperature (30 °C) for 24 h. For subsequent curing of the vector, stationary-phase cultures were diluted 1 : 100 into pre-warmed BHI medium containing 5 µg neomycin ml-1 and grown at 42 °C for 24 h. Ten-fold serial dilutions of these cultures were plated onto pre-warmed BHI agar containing neomycin and incubated at 42 °C for 48 h. Colonies were tested for vector integration on neomycin/chloramphenicol/blood agar plates. Desired colonies, in which the second recombination event at the intergenic region between hly and mpl occurred, were neomycin-resistant (Nmr), Cms and Hly-, and were blue when tested for
-glucuronidase activity on plates containing the substrate 5-bromo-4-chloro-3-indoyl
-D-glucuronide (XG) (US Biologicals). This strain was named AML73. Constructs were confirmed by PCR amplification and Southern blot hybridization.
EMS mutagenesis of AML73.
Bacteria harvested from mid-exponential-phase cultures (OD595 0·5) of AML73 grown in 50 ml BHI medium were washed twice in 1xPBS (137 mM NaCl, 2·7 mM KCl, 4·3 mM Na2HPO4.7H2O, 1·4 mM KH2PO4, pH 7·3) and resuspended in 25 ml PBS. An aliquot (2·5 ml) of a pre-warmed EMS (Sigma) solution (1 mM EMS/24 ml PBS) was mixed with an equal volume of resuspended cells and incubated at 37 °C for various lengths of time spanning 075 min. Each sample was washed twice and resuspended in 2·5 ml PBS. Colony titres were determined and samples exhibiting 9095 % killing were used for mutational analysis. Three independent EMS libraries were generated, and a total of 1x103, 3·75x104 and 6·25x104 colonies were screened from these libraries.
Generalized transduction.
Generalized transduction for linkage analysis and transposon tagging procedures were carried out as described by Hodgson (2000). To generate phage lysates, 100 µl of bacteriophage U153 dilutions made from a high-titre stock (108 p.f.u.) was mixed with 100 µl of mid-exponential-phase BHI culture of the L. monocytogenes donor strain grown at 30 °C and incubated for 40 min at room temperature. Three millilitres of molten LB agar plus 10 mM CaCl2 and 10 mM MgSO4 were added to the mix and poured onto LB plates containing 10 mM CaCl2 and 10 mM MgSO4. Plates were incubated overnight at room temperature. Lysates were harvested from just-confluent plates by adding 5 ml sterile TM buffer (8·0 mM MgSO4, 10 mM Tris/HCl, pH 8·0) and the recovered lysate was filter-sterilized to remove bacteria. The bacteriophage titres were determined as p.f.u. ml-1. To transduce L. monocytogenes, 107 p.f.u. of the bacteriophage grown on the appropriate donor strain was mixed with 108 mid-exponential-phase recipient cells, and the mixture was incubated at room temperature for 40 min. To select for Nmr transductants, the mixture was plated directly onto BHI agar containing 10 mM sodium citrate (pH 7·5) and 5 µg neomycin ml-1. To select for erythromycin-resistant (Emr) transductants, 2·5 ml BHI molten top agar containing 10 mM sodium citrate (pH 7·5) to which 100 µl of 10 µg ml-1 erythromycin had been previously added was mixed with the cells and bacteriophages, and the mixture was poured onto BHI agar containing 10 mM sodium citrate (pH 7·5). The plates were incubated for 2 h at 37 °C for induction of erm expression before another 2·5 ml BHI top agar containing 10 mM sodium citrate, 40 µl erythromycin (1 µg ml-1) and 40 µl lincomycin (25 µg ml-1) was added (lincomycin prevents the growth of spontaneous Emr colonies). Plates were incubated for 48 h at 37 °C.
Preparation of cell lysates.
Samples (10 ml) from mid-exponential-phase cultures were collected and washed once in an equal volume of 50 mM potassium phosphate buffer. The cells were resuspended in 23 ml of the same buffer and lysed three times by sonication (on ice) for 30 s each. Debris was cleared by centrifugation and 0·05 ml of the clarified supernatant was used for assay of -glucosidase activity [units of which are nmol substrate hydrolysed (mg protein)-1 min-1]. Protein concentrations in cell lysates were determined by the method of Bradford (1976)
using a Bio-Rad protein assay, with BSA as the standard.
Enzyme assays.
For plate assays, -glucuronidase activity was estimated by intensity of blue colour of bacteria spotted onto buffered LB plates containing 50 µg XG ml-1, with or without cellobiose or glucose each at a concentration of 25 mM. For liquid assays,
-glucuronidase activity was measured from late-exponential-phase cultures using 4-methylumbelliferyl-
-D-glucuronide trihydrate (US Biologicals) as a substrate. The activity was determined essentially by the fluorescence assay of Youngman (1987)
, except that 0·1 % Triton X-100 was added to the assay buffer to enhance bacterial permeability to the substrate.
-Glucosidase-specific activity, using p-nitrophenyl
-D-glucopyranoside (Sigma) as a substrate, was measured from mid-exponential-phase cultures and was assayed as described previously (Behari & Youngman, 1998b
), except that the increase in absorbance was monitored at 405 nm on a Shimadzu UV-1201 spectrophotometer. Lecithinase activity of the plcB gene product was measured on egg yolk agar plates. Five microlitres of mid-exponential-phase cultures were spotted onto LB plates (±sugars) topped with 3 ml LB agar containing 5·0 % of an equal volume egg yolk/1xPBS solution. Following incubation at 37 °C, plates were examined for precipitation of degraded egg yolk. The degree of lecithinase activity was estimated from the size of the zone (in mm) of degraded egg yolk precipitate surrounding the spots.
Mapping of csr mutation and construction of an isogenic csr mutant strain.
The csr mutation was tagged with linked transoposon insertions as described by Kaiser (1984). A U153 lysate was prepared from a population of wild-type bacteria containing a library of random Tn917 insertions and this lysate was used to transduce AML1142 to Emr. Transductants were isolated and tested by restoration of wild-type regulation by sugars using buffered LB plates containing erythromycin, 50 µg XG ml-1 and either 25 mM cellobiose or 25 mM glucose. Among 2000 AML1142 transductants tested, one was found that was no longer deregulated in the presence of cellobiose. Repeated efforts to tag the gcr mutation using Tn917 were unsuccessful. Following Tn917-tagging of the csr mutation, the exact site of Tn917 insertion was determined by DNA sequencing of the transposonchromosome junction site which had been cloned as described previously (Camilli et al., 1990
). Dideoxy sequencing of double-stranded plasmid DNA was performed by the SBRI Genome Center using an oligonucleotide primer complementary to a sequence 83 bp from the lacZ-proximal end of Tn917. Based on sequence comparisons with the L. monocytogenes EGDe genome (Glaser et al., 2001
), Tn917 was found to have inserted within the open reading frame (ORF) designated lmo1716. Based on the estimates obtained from transduction experiments regarding the linkage of csr to the Tn917-encoded erythromycin-resistance gene, chromosomal regions within 57 kb of the transposon insertion were PCR-amplified from wild-type and csr mutant strains and sequenced to identify the csr mutation. A substitution of a T for a C was found in the csr mutant strain at position 1687 of a 2679 bp ORF designated lmo1721. This mutation was not present in PCR products derived from the wild-type strain, and it was confirmed by DNA sequencing of two independent PCR products.
Reconstruction of the csr mutation in AML73.
Primers 1721-A (5'-GGGGGATCCGCAGATCAATTAATGAAAG-3') and 1721-B (5'-GGGGAATTCAGAACTTACTGCTTGTAA-3') were used to amplify a 1·2 kb fragment containing the csr mutation as well as 600 bp of flanking DNA on each side of the mutation and to introduce BamHI and EcoRI restriction sites (underlined) for subsequent cloning. The resulting PCR product was digested with BamHI and EcoRI and ligated into appropriately digested pKSV7 (Smith & Youngman, 1992), then transformed into competent E. coli DH5
. The resulting plasmid, pNF1024, was introduced into AML73 by electroporation and the csr mutation was introduced into the chromosome by allelic exchange as described previously (Camilli et al., 1993
). The presence of the csr mutation within the chromosome in the correct location was confirmed by PCR amplification of genomic DNA and digestion of the PCR product with RsaI, a restriction site created by the point mutation, and by sequencing of the PCR products.
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RESULTS |
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To evaluate the phenotypes associated with csr and gcr mutations quantitatively, single and double mutants were assayed for -glucuronidase activity during growth in buffered LB broth in the presence or absence of glucose or cellobiose. The results confirmed that mutations in both csr and gcr are required for full derepression in the presence of cellobiose, but that a mutation in gcr alone is sufficient for derepression in the presence of glucose (Fig. 4
).
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Not all glucose-repressed genes are deregulated in AML1142
Because mutations in AML1142 resulted in deregulation of hly expression in the presence of several repressing sugars, it was possible that these mutations might also affect general mechanisms of catabolite control in L. monocytogenes. Although little is known about catabolite control mechanisms in L. monocytogenes, a ccpA homologue has been identified and shown to mediate at least some aspects of catabolite control, including glucose repression of -glucosidase activity (Behari & Youngman, 1998b
). To determine whether the mutations in AML1142 affected the CcpA-mediated control pathway, we assayed
-glucosidase levels from wild-type and mutant bacteria grown in LB medium plus 25 mM maltose (for the induction of
-glucosidase) with or without the addition of cellobiose or glucose (Fig. 5
). We found that the level of
-glucosidase activity in AML1142 was slightly higher than wild-type levels in the absence of cellobiose or glucose, but did not significantly differ from that of wild-type in the presence of either sugar. This indicates that while the mutations in AML1142 significantly affected carbon-source regulation of hlygus expression in L. monocytogenes, they had little effect on the regulation of a known member of the CcpA-controlled regulon, nor presumably on CcpA-mediated catabolite regulation in general. However, since mechanisms of catabolite repression are still poorly understood in L. monocytogenes, the possibility remains that virulence gene regulation occurs through some unknown general mechanism of catabolite control.
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To confirm that the mutation within csrA was sufficient to confer the deregulated hly expression phenotype in response to cellobiose, the C to T mutation that truncates CsrA at amino acid 562 was introduced into the AML73 parent strain and the resulting mutant AML934 was assayed for hly-dependent -glucuronidase expression in the presence and absence of cellobiose. In the AML73 parent strain, the expression of hlygus in the presence of cellobiose was reduced to only 6 % (±1·4 % SE) of the levels observed in the absence of cellobiose. AML134, containing the original csr mutation, retained expression levels that were 21 % (±1·0 % SE) of those observed in the absence of cellobiose, and the introduction of the csrA mutation into AML73 was sufficient to increase the levels of hly expression in the presence of cellobiose (20 %±2·4 % SE) to those observed for the original AML134 mutant strain. These experiments confirm the role of this mutation in the partial alleviation of repression of hly expression in response to cellobiose.
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DISCUSSION |
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The introduction of a stop codon mutation at position 563 within the csrA coding sequences was sufficient to confer the partial relief of cellobiose repression observed in the AML134 mutant strain. The csrA gene product shares homology with a family of multi-domain transcriptional regulatory proteins, including LevR of B. subtilis, which controls the expression of a fructose-specific PTS and an extracellular levanase, which hydrolyses fructose polymers and sucrose (Martin et al., 1987; Martin-Verstraete et al., 1990
). LevR is a multi-domain protein, with its N-terminal domain similar to the NifA/NtrC transcriptional activator family and a C-terminal domain similar to the regulatory part of bacterial anti-terminators, such as BglG and LicT (Martin-Verstraete et al., 1998
). Based on the EGDe L. monocytogenes genome sequence (Glaser et al., 2001
), the csrA gene is followed by a transcriptional terminator. Downstream of csrA is a series of ORFs whose predicted gene products share homology with a PTS lichenan-specific enzyme IIB component (lmo1720) and a PTS lichenan-specific enzyme IIA component (lmo1719). Lichenan, like cellobiose, is a
-glucoside, and it is possible that this putative transport system responds to the presence of lichenan and/or cellobiose, as does the lic operon of B. subtilis (Tobisch et al., 1997
). The csrA mutation may therefore prevent the expression of the associated PTS gene products and thus eliminate the expression of a
-glucoside-specific sensor that mediates virulence gene repression in response to cellobiose. Such an effect would be similar to that proposed for the disruption of the bvr locus of L. monocytogenes, also reported to contribute to the repression of virulence gene expression by
-glucosides (Brehm et al., 1999
).
The results presented here imply that cellobiose can influence the expression of virulence genes by at least two separate, semi-independent pathways. One of these pathways (the one affected by gcr mutations) mediates the repressive effects of all readily metabolized sugars and probably represents a global pathway for catabolite repression. If so, however, our results and those of Behari & Youngman (1998b) indicate that the L. monocytogenes CcpA protein is probably not involved. The other pathway is cellobiose (or
-glucoside) specific and is influenced by both csr- and bvr-encoded gene products. Nevertheless, the csr pathway (and perhaps bvr) is not completely independent of the gcr pathway, since mutations in csr are necessary but not sufficient for full relief from cellobiose repression. Although we cannot provide a mechanistic description of these pathways, earlier work suggests that they both act through PrfA, as indicated schematically in Fig. 7
. Moreover, since previous work has demonstrated that levels of PrfA protein do not change in the presence of repressing carbon sources (Milenbachs et al., 1997
; Renzoni et al., 1997
), it is likely that both pathways converge upon PrfA either through a covalent modification of the protein or through the synthesis of a co-inducer molecule that modifies the DNA-binding activity of PrfA.
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ACKNOWLEDGEMENTS |
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Received 18 August 2003;
revised 19 November 2003;
accepted 20 November 2003.
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