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Journal of Bacteriology, February 2009, p. 862-872, Vol. 191, No. 3
0021-9193/09/$08.00+0     doi:10.1128/JB.01384-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

CcpC-Dependent Regulation of Citrate Synthase Gene Expression in Listeria monocytogenes

Meghna Mittal,¹ Silvia Picossi,^2, and Abraham L. Sonenshein^1,2*

Program in Molecular Microbiology, Sackler School of Graduate Biomedical Sciences,¹ Department of Molecular Biology and Microbiology, School of Medicine, Tufts University, Boston, Massachusetts 02111²

Received 2 October 2008/ Accepted 4 November 2008

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Citrate synthase, the first and rate-limiting enzyme of the tricarboxylic acid branch of the Krebs cycle, was shown to be required for de novo synthesis of glutamate and glutamine in Listeria monocytogenes. The citrate synthase (citZ) gene was found to be part of a complex operon with the upstream genes lmo1569 and lmo1568. The downstream isocitrate dehydrogenase (citC) gene appears to be part of the same operon as well. Two promoters were shown to drive citZ expression, a distal promoter located upstream of lmo1569 and a proximal promoter located upstream of the lmo1568 gene. Transcription of citZ from both promoters was regulated by CcpC by interaction with a single site; assays of transcription in vivo and assays of CcpC binding in vitro revealed that CcpC interacts with and represses the proximal promoter that drives expression of the lmo1568, citZ, and citC genes and, by binding to the same site, prevents read-through transcription from the distal, lmo1569 promoter. Expression of the lmo1568 operon was not affected by the carbon source but was repressed during growth in complex medium by addition of glutamine.

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Listeria monocytogenes is a gram-positive food-borne bacterial parasite of mammals that causes listeriosis, an infection with a 30% mortality rate in susceptible humans that is characterized by fetoplacental and central nervous system infections and gastroenteritis (38). The risk group for listeriosis includes pregnant women, neonates, the elderly, and immunocompromised adults. Listeria infection has been an important model system for the study of host-pathogen interactions and mechanisms of intracellular parasitism (38). This bacterium is an intracellular pathogen that induces its own uptake by nonphagocytic cells and spreads from cell to cell via actin-based motility (4). Even though L. monocytogenes is able to grow intracellularly in a variety of mammalian cells, it is a facultative pathogen that can adapt to saprophytic growth on decaying soil vegetation (28). It is therefore interesting to examine how this bacterium senses the environment in order to regulate expression of its virulence determinants.

Previous researchers have thoroughly investigated the molecular determinants of L. monocytogenes pathogenesis. The genes that encode all the currently known virulence factors are positively regulated by the transcriptional activator PrfA (3). These genes are repressed, however, when L. monocytogenes is grown in the presence of fermentable sugars (28). This carbon source-mediated repression of virulence genes does not involve CcpA, the global regulator of catabolite control in many gram-positive bacteria (2, 16, 36), but instead is due to effects of sugar metabolism on PrfA activity. Rapidly metabolized carbon sources alter the phosphorylation state of components of the phosphoenolpyruvate-dependent phosphotransferase system; one or more of these components appear to inhibit PrfA (17, 27). Given that there is some uncertainty about the mechanisms that couple utilization of carbon sources and expression of virulence factors in L. monocytogenes, expanding our knowledge about the regulation of central metabolism may help us gain insight into the roles played by carbon sources in virulence factor gene regulation.

One central metabolic pathway, the Krebs citric acid cycle, has the potential to generate ATP, reducing power, and key biosynthetic precursors. L. monocytogenes, however, utilizes a split Krebs cycle; it has an oxidative, tricarboxylic acid (TCA) branch (citrate synthase, aconitase, and isocitrate dehydrogenase) leading to 2-ketoglutarate synthesis and a reductive branch (37). This bacterium lacks the 2-ketoglutarate dehydrogenase system, succinyl coenzyme A (succinyl-CoA) synthetase, and succinic dehydrogenase, as well as the enzymes of the glyoxylate shunt. In L. monocytogenes, the sole metabolic role of the TCA branch enzymes seems to be to mediate the synthesis of 2-ketoglutarate. 2-Ketoglutarate is a precursor for synthesis of glutamate and glutamine, the cell's most critical nitrogen-containing metabolites.

The L. monocytogenes genome has been sequenced completely (14). Hence, it is possible to deduce the presence of some metabolic genes, as well as genes for their transcriptional regulators, by comparison with the genomes of more completely characterized relatives, such as Bacillus subtilis. Such an analysis revealed the presence of homologs of the citZ (citrate synthase), citB (aconitase), and citC (isocitrate dehydrogenase) genes. Since carbon metabolism in B. subtilis has been extensively analyzed, the information obtained in this research can be useful for gaining a better understanding of L. monocytogenes physiology.

In B. subtilis, the synthesis and activity of the three enzymes of the TCA branch of the Krebs cycle are regulated in response to the cell's requirement for ATP, reducing power, and 2-ketoglutarate as a precursor for glutamate and glutamine. The citZ, citC, and citH (malate dehydrogenase) genes form an operon (19). The citC and citH genes also have gene-specific promoters (18, 20). In the presence of a readily utilizable carbon source, such as glucose, and a source of 2-ketoglutarate, such as glutamate or glutamine, the synthesis and activities of the TCA branch enzymes are reduced (9, 11, 15), and the transcription of the citZCH operon and the citB gene is strongly repressed (20, 31). CcpC, a member of the LysR family, is a major transcriptional regulator of the citZCH operon and the citB gene (22). CcpC binds with high affinity to the citZ and citB promoter regions and is a direct repressor of transcription (22, 26, 31). In the presence of citrate, binding of CcpC to the citZ and citB promoters is altered and both of these genes are derepressed (22).

Carbon catabolite repression of the B. subtilis citZ and citB genes is also mediated by CcpA (25). When cells are grown in the presence of glucose, CcpA is activated by interaction with the phosphorylated form of either HPr or Crh proteins (13, 34, 35). In B. subtilis, activated CcpA directly represses the transcription of the citZCH operon when cells are grown in a medium containing glucose, and it plays an indirect role in citB regulation by affecting the activity of CcpC (25); that is, CcpA restricts the synthesis of citrate, keeping CcpC in its active form (25).

L. monocytogenes encodes a homolog of B. subtilis CcpC that binds tightly to the L. monocytogenes citB promoter region in vitro and represses citB transcription in vivo (24). Citrate inhibits the interaction of CcpC with the citB promoter region. We report here that the L. monocytogenes CcpC protein also represses transcription of citZ. Transcription of citZ was shown to initiate at promoters for two different genes upstream of citZ. CcpC was shown to bind to a dyad symmetry element that is very similar to dyad elements found in other CcpC-binding sites and that overlaps the proximal promoter driving citZ expression. In addition to this direct mode of regulation, CcpC also repressed read-through transcription from the more distantly located promoter.

The L. monocytogenes homolog of B. subtilis CcpA controls the utilization of certain carbon sources (2), but it was found to play no apparent role in regulation of the citZ gene.

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Bacterial strains and growth conditions. Bacterial strains used in this study are listed in Table 1. Preparation of electrocompetent Escherichia coli cells and transformation by electroporation using a TransPoratorPlus (BTX) were performed as described by other workers (7). A previously described method was used for transformation of B. subtilis with chromosomal or plasmid DNA (8). Preparation of electrocompetent L. monocytogenes cells with penicillin pretreatment and transformation by electroporation were performed as described previously (29).

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TABLE 1. Bacterial strains used

E. coli strains were grown at 37°C in Luria broth or on Luria agar plates with appropriate antibiotics. Both kanamycin (50 µg/ml) and ampicillin (100 µg/ml) were added to cultures of E. coli strains carrying pHK77 and its derivatives to prevent loss of the plasmids. B. subtilis strains were grown at 37°C in TSS minimal medium (12) supplemented with 0.2% (wt/vol) glutamine and 0.5% (wt/vol) glucose. For measurement of β-galactosidase activity in B. subtilis, 1-ml culture samples were harvested and assayed as described previously (25). L. monocytogenes was grown at 37 or 30°C in brain heart infusion (BHI) medium (Difco) or IMM (see below), as indicated. The following antibiotics were added when necessary: chloramphenicol (2.5 µg/ml), phleomycin (0.25 µg/ml), erythromycin (1 µg/ml), spectinomycin (50 µg/ml), and neomycin (5 µg/ml) for B. subtilis and chloramphenicol (10 µg/ml), spectinomycin (100 µg/ml), and neomycin (10 µg/ml) for L. monocytogenes.

IMM medium (30) was used to analyze the effects of different carbon sources on citZ-lacZ expression. L. monocytogenes strains were grown overnight in BHI medium, and an overnight culture was used as the inoculum for subsequent growth in IMM medium with 0.5% glucose as the carbon source. When they reached stationary phase, the cultures were diluted in IMM medium with different carbon sources (0.5%) to obtain an initial optical density at 600 nm (OD₆₀₀) of 0.05. One-milliliter samples were collected in the exponential (OD₆₀₀, 0.5) and stationary (OD₆₀₀, 1) phases of growth for β-galactosidase assays.

DNA manipulation. Restriction digestion, DNA ligation, and PCR were performed according to the manufacturer's instructions. E. coli plasmids were isolated using a Qiagen miniprep kit (Qiagen Inc.). Agarose gel electrophoresis and polyacrylamide gel electrophoresis were carried out as described by Sambrook et al. (32). Genomic DNA from B. subtilis was isolated as described by Fouet and Sonenshein (12) using 1 mg/ml lysozyme. L. monocytogenes chromosomal DNA was isolated using mutanolysin and the protocol of Fliss et al. (10).

Construction of B. subtilis strains carrying L. monocytogenes citZ-lacZ fusions. To construct the lacZ fusion plasmids pEMM10, pEMM11, and pEMM13, various regions upstream of the L. monocytogenes citZ gene were amplified by PCR from chromosomal DNA of L. monocytogenes strain EGD-e using forward primers OMM014, OMM015, and OMM016, respectively, and reverse primer OMM017. These primers and all other primers used in this work are listed in Table 2. The PCR products were digested with EcoRI and HindIII, and the products were cloned independently in similarly digested vector pHK23 (H. J. Kim, personal communication). Transformation of B. subtilis strain BB1888 with pEMM10, pEMM11, and pEMM13 resulted in erythromycin-resistant transformants BMM6, BMM7, and BMM8, respectively, in which citZ-lacZ fusions were integrated at the chromosomal amyE locus by double-crossover recombination. The BMM6, BMM7, and BMM8 strains contain 1,180, 689, and 484 bp, respectively, of sequence upstream of the citZ start codon fused to the lacZ reporter (Fig. 1).

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TABLE 2. Oligonucleotides used in this work

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FIG. 1. Organization of the citZ locus in L. monocytogenes and genetic content of lacZ fusions used in B. subtilis. L. monocytogenes genes are indicated by designations used at the ListiList database site (http://genolist.pasteur.fr/ListiList/). The arrows indicate genes and their orientations, and the balloons indicate the locations of putative transcription termination sites. A putative CcpC-binding site is indicated by a box in the region upstream of lmo1568. The regions fused to lacZ and used for assays of gene expression in B. subtilis are indicated. The arrowheads indicate the putative positions of transcription start sites. The figure is not drawn to scale.

Strains BMM1, BMM3, and BMM4 were constructed by transforming strains BMM6, BMM7, and BMM8 with chromosomal DNA from LS1002 (ccpC::spc) and selecting for Spc^r transformants. To create strains BMM12 and BMM13, BMM6 and BMM7 were transformed with chromosomal DNA of strain HKB5 (ccpA::ble) and phleomycin-resistant transformants were selected.

Construction of a citZ null mutant strain of L. monocytogenes. An L. monocytogenes citZ null mutant strain (LMM33) was created using the overlap PCR technique (39). First, a 706-bp PCR product corresponding to the 5' end of the citZ gene (the first 100 codons) and upstream DNA including part of the lmo1568 gene was generated by PCR using L. monocytogenes chromosomal DNA as the template and primers OMM099 and OMM100. A second PCR product (688 bp) corresponding to the 3' end of citZ and a downstream region including part of the citC gene was generated using primers OMM101 and OMM102. The ends of primers OMM100 and OMM101 have extensions that are complementary to each other. The two PCR products were purified, mixed together, and used in a third PCR with primers OMM099 and OMM102 to generate a 1.4-kb product that corresponded to the citZ gene with an internal, in-frame deletion of 171 codons and about 827 bp of flanking DNA upstream and downstream. This PCR product was purified, digested with BamHI and NcoI, and ligated to a similarly digested vector, pMAD (1), to create pEMM47. The latter plasmid was used to transform electrocompetent cells of L. monocytogenes strain HKB214 to erythromycin resistance. The initial transformants arose by single crossover, integrating the entire pEMM47 plasmid. To isolate a clone that had undergone a second crossover event and lost the pMAD-1 sequence, we subcultured an initial transformant three times overnight at 30°C. The presence of the citZ null mutation in the resultant strain, LMM33, was confirmed by PCR and by the absence of citrate synthase enzyme activity.

Construction of strains of L. monocytogenes carrying lacZ fusions. To construct fusions to lacZ of various regions from the L. monocytogenes citZ locus, the regions between pykA and lmo1569, between lmo1569 and lmo1568, and between lmo1568 and citZ were amplified by PCR using primer pairs OMM048/OMM049, OMM050/OMM051, and OMM016/OMM017, respectively, and chromosomal DNA of strain EGD-e as the template. To create a lacZ fusion to the region between lmo1569 and citZ, primer pair OMM015/OMM017 was used. The PCR products were digested with EcoRI and KpnI and ligated independently to pHK77 (24), which had been similarly digested. Plasmid pHK77 does not replicate in L. monocytogenes at elevated temperatures but integrates by homologous recombination at the nonessential int'-'comK locus (24). After transformation of E. coli and isolation of plasmid DNA, the insert was sequenced to confirm that it did not have PCR-generated mutations. Five micrograms of each pHK77 derivative was mixed with electrocompetent cells of strain EGD-e. After electroporation (2 kV, 400 , 25 µF, 5 ms), cells were diluted with 1 ml BHI medium containing 0.5 M sucrose and incubated without shaking at 30°C for 60 min. Transformants were selected at 30°C on BHI medium plates containing neomycin and then grown at 30°C in the presence of neomycin overnight three times to increase the copy number of the plasmid. After subsequent overnight growth at the nonpermissive temperature (41°C), neomycin-resistant clones were tested individually by performing PCR to determine integration into the chromosome. In the last step, strains were grown at 30°C in the absence of neomycin to permit growth of segregants that had lost the plasmid due to excision. A series of PCRs confirmed that the lacZ fusions were inserted by double-crossover recombination at the int-comK locus and that the strains did not carry any free plasmid.

To introduce a ccpC::spc mutation into the strains carrying lacZ fusions, pHK95 (24) was transferred by electroporation into electrocompetent cells. After incubation at 41°C in the presence of spectinomycin to select for integration of pHK95 into the chromosome, cells were grown at 30°C to allow plasmid excision, and null mutants were purified based on their spectinomycin resistance and chloramphenicol sensitivity. A series of PCRs confirmed that there was allelic replacement of ccpC⁺ by ccpC::spc.

ccpA null mutant strains SP121, SP122, and SP124 were created using overlap PCR (39). First, a 617-bp PCR product including the 5' region of the ccpA gene was generated by PCR using L. monocytogenes chromosomal DNA as the template and primers OSP172 (with an appended SmaI site at the 5' end) and OSP158 (with an internal BamHI site). A second PCR product (614 bp) including the 3' end of ccpA was generated using primers OSP159 (with an internal NcoI site) and OSP174 (with an appended SmaI site at the 5' end). Primers OSP158 and OSP159 have extensions that are complementary to each other. The two PCR products were purified, mixed together, and used in a third PCR with primers OSP172 and OSP174. The PCR product was purified, digested with BamHI and NcoI, ligated to the BamHI-XhoI DNA fragment (1.8 kb) from plasmid pEMM20 (M. Mittal, unpublished) corresponding to the tetracycline resistance cassette, digested again with SmaI, and ligated to SmaI-digested pMAD to create pSP65.

The latter plasmid was used to transform electrocompetent cells of L. monocytogenes strains LMM18, LMM19, and LMM24 to erythromycin and tetracycline resistance. The initial transformants arose by a single crossover event, and the entire pSP65 plasmid was integrated. To isolate a clone that had undergone a second crossover event and lost the pMAD sequence, we incubated a culture of an initial transformant overnight three or four times at 30°C in BHI medium to allow plasmid excision and create ccpA null strains SP121, SP122, and SP124, respectively. The overnight cultures and plates contained BHI medium with tetracycline, as well as 1.6 mg of glutamate per ml and 1.6 mg of glutamine per ml. The presence of the ccpA null mutation was confirmed by PCR.

Citrate synthase enzyme assay. Cells from a 25-ml stationary-phase culture in BHI medium were collected by centrifugation at 10,000 x g for 20 min using a refrigerated centrifuge. The pellet was resuspended in 2 ml of 0.05 M Tris-Cl buffer (pH 7.5), and the cells were broken by sonication with a Branson Sonifier cell disrupter (model 200) using five 30-s cycles with 30-s rest periods between the cycles on ice. Cell debris was removed by centrifugation at 5,000 rpm with a refrigerated tabletop centrifuge for 10 min. The total protein concentration in the supernatant was determined using the Bio-Rad protein assay reagent. The activity of citrate synthase was determined by measuring the splitting of 5,5'-dithio-bis(2-nitrobenzoic acid) by CoA liberated from acetyl-CoA using a reaction mixture containing 20 mM Tris-Cl (pH 7.4), 10 mM sodium oxaloacetate, 0.5 mM acetyl-CoA, and 1 mM 5,5'-dithio-bis(2-nitrobenzoic acid). The reaction was monitored at 412 nm and was calibrated with CoA. One unit of enzyme activity was defined as the formation of 1 µmol of product per min. Specific activity was expressed in units per milligram of protein.

Gel mobility shift assays and DNase I footprinting experiments. L. monocytogenes CcpC-His₆ was purified as previously described (24). For gel shift assays, a 237-bp DNA fragment corresponding to the lmo1568 promoter region was amplified by PCR using ³²P-labeled OMM050 and OMM064 as the forward and reverse primers and genomic DNA from L. monocytogenes EGD-e as the template. A 191-bp DNA fragment corresponding to the lmo1569 promoter region was amplified by PCR using ³²P-labeled OMM048 and OMM049 as the forward and reverse primers. The primers (50 pmol) were end labeled by incubation with 150 µCi of [-³²P]ATP (6,000 Ci mmol^–1; NEN) and T4 polynucleotide kinase (Invitrogen). The labeled primers were purified using a nucleotide removal kit (Qiagen Inc.). The labeled PCR products (1,000 cpm) and various amounts of CcpC-His₆ were incubated in buffer B for 30 min at room temperature and electrophoresed on nondenaturing polyacrylamide gels as previously described (24).

For DNase I footprinting experiments, primers were labeled as described above. OMM074 and ³²P-labeled OMM064 were the primers used to amplify a 548-bp fragment covering the lmo1568 promoter region along with genomic DNA from L. monocytogenes strain EGD-e as the template. The ³²P-labeled DNA product (20,000 cpm) and various amounts of CcpC-His₆ were incubated in 20-µl reaction mixtures containing buffer B as described above. After 30 min of incubation at room temperature, MgCl₂ and CaCl₂ were each added to a final concentration of 6 mM, and the reaction mixtures were treated with 0.5 U of RQ1 DNase I (Promega) for 1 min at room temperature. For CcpC-His₆ concentrations of 62 nM, the DNase I reaction time was increased to 2 to 3 min. To create a DNA sequencing ladder, plasmid pEMM40 was subjected to the dideoxy chain termination protocol (33) using a Sequenase reagent kit, -³⁵S-dATP, and primer OMM064.

RNA isolation. L. monocytogenes strains grown overnight in BHI medium were diluted to obtain an initial OD₆₀₀ of 0.05 in fresh BHI medium, and the cultures were incubated at 37°C with shaking. Samples used for RNA isolation were collected in mid-exponential phase (OD₆₀₀, 0.3) and stationary phase (OD₆₀₀, 1.0), resuspended in 1 ml of ice-cold 50% acetone-50% ethanol, and stored at –80°C. The cells were centrifuged at 4°C, washed twice with 500 µl of TE buffer (10 mM Tris-Cl, 1 mM EDTA; pH 8.0), and resuspended in 1 ml of buffer RLT from a Qiagen RNeasy kit. Silica glass beads (diameter, 0.1 mm) were added, and the cells were disrupted using a Mini-BeadBeater (BioSpec Products Inc.). The silica beads were removed by centrifugation, and total RNA was isolated from the supernatant fluid using a Qiagen RNeasy kit (Qiagen Inc.) according to the manufacturer's instructions.

RT-PCR experiments. To remove contaminating DNA, the RNA samples described above were treated with Turbo DNase I from an Ambion DNA-free kit. Reverse transcription (RT) was performed by using the manufacturer's protocol, SuperScript II reverse transcriptase (Invitrogen) with total RNA (1,000 ng for the P1 read-through transcript, 250 and 500 ng for citZ and citC, or 50 ng for rRNA), and 2 pmol of a gene-specific primer (OMM086 for rRNA, OMM064 for the P1 read-through transcript, OMM098 for citC and the citZ-citC read-through transcript, or OMM088 for citZ) (Fig. 2) in a 20-µl (total volume) mixture. Two-microliter samples of the RT reaction mixtures were used as templates for PCR with gene-specific primers OMM086 and OMM085 for 16S rRNA, OMM064 and OMM050 for the P1 read-through transcript, OMM087 and OMM088 for citZ, OMM097 and OMM098 for citC, and OMM140 and OMM098 for the citZ-citC read-through transcript. The PCRs were performed using 12 to 27 cycles of denaturation (94°C, 30 s), annealing (45°C, 1 min), and extension (72°C, 30 s). For each gene, the PCR conditions were tested to be sure that the number of PCR cycles and the amount of RNA used were below the saturation limits. To check for chromosomal DNA contamination, 1-µl samples of RNA were used directly for PCRs after Turbo DNase treatment.

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FIG. 2. Genetic content of lacZ fusions integrated at the L. monocytogenes int'-'comK locus. The regions fused to lacZ and used for assays of gene expression in L. monocytogenes are indicated below the gene map. The arrows above the gene map indicate the annealing sites and orientations of primers used for RT-PCR experiments. See the legend to Fig. 1 for additional details.

To quantify the amount of product in RT-PCR experiments, ImageQuant software was used to capture gel bands, calculate the band intensities (expressed in pixels), subtract the local background, and normalize the data to the areas of the bands. For all experiments, the amount in the experimental sample was normalized to the amount of an RT-PCR product of 16S rRNA. The values obtained for RNA samples from mutant cells were compared to the values obtained for RNA samples from wild-type cells. The values reported below are the averages of two or three biological replicates.

Primer extension experiments. Primer extension experiments were performed with the following oligonucleotides: primer OMM047 for lmo1569 and primers OMM070, OMM071, and OBB102 for lmo1568-lacZ fusion mRNA. The latter primers anneal within the lacZ gene or the spoVG ribosome-binding site of the fusion construct (24). Primers were labeled as described above. A total of 10⁶ cpm of primer and 20 µg of total RNA were lyophilized together, resuspended in 20 µl of hybridization buffer containing 80% formamide, 0.5 M NaCl, 1 mM EDTA, and 40 mM piperizine-N,N'-bis(2-ethanesulfonic acid) (PIPES) (pH 6.8), and incubated at 80°C for 10 min. After slow cooling to 30°C, the primer-RNA mixture was precipitated with ammonium acetate and ethanol and used for an RT reaction catalyzed by SuperScript II (Gibco BRL, Life Technologies). After incubation at 42°C for 50 min, the enzyme was inactivated by heating the mixture at 70°C for 15 min, and then the reaction mixture was extracted with phenol-chloroform, precipitated, and dissolved in 10 µl of formamide-containing loading dye. Extension products were analyzed on 6% polyacrylamide-8 M urea sequencing gels.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Identification of the L. monocytogenes citZ gene. In B. subtilis, citZ is the first gene of the citZCH operon (20). The contiguous L. monocytogenes citZ and citC genes (Fig. 1) were located on the basis of homology with their B. subtilis counterparts using the ListiList database (http://genolist.pasteur.fr/ListiList/) that includes the genome sequence data of Glaser et al. (14). In B. subtilis, a second citrate synthase is encoded by the citA gene (19), but our search of the L. monocytogenes genome revealed only a single coding sequence with significant homology to B. subtilis citZ. Based on the locations of apparent transcription terminators, the L. monocytogenes citZ and citC genes seemed to be organized in an operon with the upstream genes lmo1569 and lmo1568, which encode proteins having unknown functions (Fig. 1). No gene similar to citH was detected downstream of citC. We created a citZ in-frame deletion mutant (see Materials and Methods) and confirmed that the citZ gene encodes citrate synthase enzyme activity. An extract of the wild-type strain had a citrate synthase specific activity of 0.5 U per mg protein, while the citZ mutant had a specific activity of 0.02 U per mg protein.

Use of lacZ fusions in B. subtilis to locate citZ promoters. To study the regulation of L. monocytogenes citZ, we first mapped the promoter(s) that drives citZ expression using B. subtilis as a surrogate host. We created lacZ transcriptional fusions with DNA segments corresponding to various lengths of DNA upstream of the citZ coding sequence and integrated the fusions in the B. subtilis chromosome at the nonessential amyE locus (Fig. 1). When lacZ expression was measured for the long, medium, and short fusions in a wild-type B. subtilis strain, only the long fusion expressed significant β-galactosidase activity (Table 3).

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TABLE 3. Effects of ccpC or ccpA mutations on expression of L. monocytogenes citZ in B. subtilis

To analyze the potential roles of CcpC and CcpA transcription factors in the regulation of L. monocytogenes citZ expression, we tested the three fusions in B. subtilis ccpC and ccpA mutant strains. A null mutation in ccpC caused expression of the long fusion to be derepressed sixfold compared to the expression in the wild-type strain (Table 3). While the medium-length fusion showed a very low level of citZ-lacZ expression in a ccpC⁺ background, 38-fold derepression was observed in the ccpC null mutant. This result indicated that the medium-length fusion contained a promoter (which we designated P2) that was repressed by B. subtilis CcpC. Since with the long fusion there was a high level of β-galactosidase activity both in the presence and in the absence of active CcpC, we deduced that there are probably at least two promoters that drive citZ-lacZ expression, at least one of which (P2) is regulated by B. subtilis CcpC (see below). The short fusion did not result in significant β-galactosidase activity in either the cppC⁺ or ccpC mutant strain (Table 3), indicating that no promoter active under such conditions is located between lmo1568 and citZ.

In a B. subtilis ccpA null mutant, both the medium-length and long fusions had the same β-galactosidase activities that they had in the wild-type strain, indicating that B. subtilis CcpA does not regulate L. monocytogenes citZ transcription (Table 3).

Expression of lacZ fusions in L. monocytogenes. To confirm the CcpC-mediated regulation of citZ expression, we cloned four regions (designated fusions A to D [Fig. 2 and Table 4]) upstream of the citZ gene in plasmid pHK77 (24) and in this way created lacZ transcriptional fusions integrated at the ectopic int'-'comK locus on the L. monocytogenes chromosome. We compared the promoter activities of these regions by performing β-galactosidase assays with cells grown in BHI medium (Table 4). A strain carrying fusion D had a level of β-galactosidase activity similar to the background level, indicating that the region immediately upstream of the citZ gene does not have a promoter that is active under the conditions tested. Fusion A, corresponding to the region upstream of lmo1569 fused to lacZ, had a relatively high level of β-galactosidase activity, indicating the presence of a promoter (designated P1) in the region upstream of lmo1569. Fusions B and C were expressed at lower but still significant levels, consistent with the idea that there is a promoter (P2) in the region upstream of lmo1568. These results are also consistent with the results obtained for B. subtilis (Table 3).

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TABLE 4. Effects of ccpC, ccpA, and ccpA ccpC mutations on citZ expression in L. monocytogenes

Repression of citZ expression by CcpC in L. monocytogenes. To investigate the role of L. monocytogenes CcpC in regulation of the citZ gene, we introduced a null mutation in the ccpC gene (24) into strains carrying fusions A to D (Fig. 2). The β-galactosidase activities in the resultant strains were compared to those in the corresponding wild-type strains (Table 4). The ccpC null versions of strains carrying fusions B and C showed approximately four- to eightfold derepression compared to their ccpC⁺ counterparts, confirming that P2, the promoter upstream of lmo1568, is regulated by CcpC. On the other hand, the ccpC null strain carrying the lmo1569-lacZ fusion (fusion A) had the same level of lacZ expression as its ccpC⁺ counterpart. These results indicate that the P1 promoter upstream of lmo1569 is CcpC independent (Table 4). A strain carrying fusion D had the same low level of β-galactosidase activity whether the ccpC gene was the wild-type gene or a mutant gene, indicating that the region between lmo1568 and citZ does not have promoter activity in either B. subtilis or L. monocytogenes.

Repression of citZ expression by CcpA in L. monocytogenes. To examine the potential role of CcpA in the regulation of citZ in L. monocytogenes, we created ccpA single mutants and ccpA ccpC double mutants of the strains carrying fusions C and D. The ccpA single mutants had levels of lmo1568-citZ-lacZ and citZ-lacZ expression similar to those of the corresponding wild-type strains with these fusions (Table 4). Fusion D did not result in significant β-galactosidase activity even in a ccpA ccpC double mutant, reinforcing the idea that the region between lmo1568 and citZ does not contain a promoter (Table 4).

To confirm that CcpA does not regulate citZ expression in L. monocytogenes, we introduced the ccpA mutation into wild-type (LMM19) and ccpC null (LMM24) strains and assayed citZ transcription by RT-PCR (Fig. 3). By quantifying the results shown in Fig. 3 and the results of two additional experiments, we determined that the citZ transcript is present at the same level in the wild-type and ccpA mutant strains. The ccpC and ccpC ccpA mutants had similar 9- to 10-fold-higher levels of the citZ transcript than the wild-type strain (Fig. 3). Thus, CcpA does not regulate citZ expression in L. monocytogenes.

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FIG. 3. RT-PCR analysis of the effect of ccpA mutation on citZ expression. RNA was extracted from wild-type, ccpC mutant, ccpA mutant, or ccpC ccpA double-mutant cells growing exponentially in BHI medium. Primer OMM88 was used for RT, and OMM88 and OMM87 were the primers used in PCR with the cDNA. The upper panels show the results for the citZ transcript (two different amounts of RNA and two different numbers of PCR cycles). The lower panels show the results for two different amounts of rRNA used as a control. The intensities of the bands were quantified using ImageQuant software. In two independent experiments, the average values for the citZ transcript relative to the wild type, normalized to rRNA, were 0.9 ± 0.1 for the ccpA mutant, 10 ± 2 for the ccpC mutant, and 9 ± 3 for the ccpA ccpC mutant.

Mapping transcription start sites. To map the start sites of L. monocytogenes citZ transcripts, we extracted RNA from wild-type and ccpC null strains (grown in BHI medium) and performed a primer extension analysis. We mapped the apparent transcription start point (tsp) of lmo1569 to a position located 21 bp upstream of the start codon (data not shown). The abundance of this primer extension product was not affected by a ccpC mutation (data not shown). The putative P1 tsp is preceded by sequences that resemble the –10 and –35 recognition sites for the sigma A form of RNA polymerase. We tried to map the tsp of lmo1568 in the same way, but, due to the presence of multiple RT products, the tsp could not be determined conclusively even when we tested several different primers (data not shown). Nonetheless, a comparison of the levels of primer extension products obtained for the wild-type and ccpC null strains showed that some products were more abundant when RNA from the ccpC mutant strain was used (data not shown).

Since the complexity of the primer extension products observed for lmo1568 might have been due to transcripts originating upstream of P2, we tried to simplify the analysis by using strains in which the region containing the P2 promoter is upstream of the lacZ gene at the ectopic int-comK locus. In such strains, lacZ expression is not influenced by read-through transcription from any upstream promoter. We designed primers that anneal within the lacZ part of the fusion and used them to map the P2 tsp. Using three different primers, we found that the apparent tsp is located 42 bp upstream of the start codon of lmo1568 and that transcripts corresponding to this site are derepressed in a ccpC mutant strain (Fig. 4).

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FIG. 4. Primer extension analysis to map the tsp for the lmo1568 gene. Three oligonucleotide primers that were labeled with 32P at the 5' end and were complementary to lacZ mRNA (see Materials and Methods) were annealed to total cellular RNA that was extracted from wild-type strain EGD-e and ccpC null mutant cells carrying a lmo1568-lacZ fusion and growing exponentially in BHI medium and were extended by using deoxynucleoside triphosphates and reverse transcriptase. The DNA products were separated by electrophoresis on a 6% polyacrylamide-7 M urea gel, and their mobilities were compared with the mobilities of dideoxynucleotide sequencing ladders (lanes A, C, G, and T) generated using two of the same primers used for primer extension and the relevant cloned DNA as the template. (The length of the third reverse transcript was extrapolated from its mobility relative to the mobilities of DNA sequences having known lengths generated with the other primers.) The left and right lanes in each set show products obtained with RNA from the wild-type (WT) and ccpC mutant strains, respectively. The following primers were used: for the left panel, OMM070; for the middle panel, OMM071; and for the right panel, OBB102. The arrows indicate the positions of the primer extension products. The sequence of the lmo1568 promoter region is shown below the gels. The consensus recognition sequences for CcpC are indicated by uppercase letters. The putative –35 and –10 regions and transcription start site (+1) of the deduced P2 promoter are also indicated. The start codon (ATG) is italicized.

Read-through transcription from P1 into lmo1568. To test directly whether there is read-through transcription from the P1 promoter into the downstream genes, we carried out a semiquantitative RT-PCR analysis using primers that flank the P2 promoter (Fig. 5). The presence of a PCR product would have indicated that transcription that initiates upstream of the P2 tsp can extend beyond the P2 promoter. By quantifying the amount of this transcript in wild-type and ccpC null strains (see Materials and Methods), we found that read-through transcription was derepressed 3.5-fold by a ccpC mutation in cells grown in BHI medium.

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FIG. 5. RT-PCR analysis of read-through transcription from the P1 promoter into lmo1568. RNA was extracted from cells growing exponentially in BHI medium. Lanes 1, 3, 5, 7, 9, and 11 contained RT-PCR products from the wild-type strain, and lanes 2, 4, 6, 8, 10, and 12 contained RT-PCR products from the ccpC null strain. For analysis of read-through transcription from P1, the primer used for RT was OMM64; the PCR was primed with OMM50 and OMM64 (Fig. 2). One microgram of RNA was used for RT, and 10% of the product was used as a template for the PCR. To assay for rRNA as a normalization factor, 50 ng of RNA was used for cDNA synthesis and 10% of the product was used as a template for a PCR. The numbers of PCR cycles are indicated below the gels. The band intensities for the ccpC mutant strain RNA normalized to rRNA and relative to the wild-type intensities (wt) for the experiment shown were determined by using densitometry and ImageQuant software and are also indicated below the gels. In three independent experiments, the average value for the read-through transcript normalized to rRNA and relative to the wild-type value was 3.5 ± 0.5 for the ccpC mutant.

Read-through transcription into the citC gene and regulation of citC by CcpC. In B. subtilis, citC, the gene downstream of citZ in the operon, encodes isocitrate dehydrogenase and is coregulated with citZ. About 50% of the citC transcripts arise from the citZ promoter and are regulated by CcpC (20). A secondary citC promoter is located immediately upstream of the citC gene but is not subject to carbon source-mediated regulation. To find out if citC is coregulated with citZ in L. monocytogenes, we carried out a semiquantitative RT-PCR analysis using primers within the citZ and citC genes. We detected a transcript, and it was repressed 2.5- ± 0.2-fold by CcpC in cells grown in BHI medium (Fig. 6A). We also tested the expression of the citC gene itself using primers that are within the citC gene. The ccpC null strain had a 3- ± 0.2-fold-higher level of the citC transcript than the wild-type strain (Fig. 6B).

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FIG. 6. RT-PCR analysis of read-through transcription from citZ into the citC gene and regulation of citC by CcpC. RNA was extracted from wild-type and ccpC mutant cells growing exponentially in BHI medium. To assay for citZ-citC read-through transcription or for citC transcription, 500 ng of RNA was used for RT and 10% of the product was used as a template for a PCR. To assay for rRNA as a normalization factor, 50 ng of RNA was used for cDNA synthesis and 10% of the product was used as a template for PCR. Data from two independent experiments (experiments 1 and 2) are shown. Each pair of lanes shows the results obtained with RNA from wild-type cells (lanes a) and from ccpC mutant cells (lanes b). The numbers of PCR cycles are indicated below the gels. Band intensities were quantified by using densitometry and ImageQuant software. (A) Read-through transcription from citZ into the downstream citC gene. The primers used were OMM140 and OMM98 (Fig. 2). In two independent experiments, the average amount of read-through transcript (normalized to the amount of rRNA) was 2.5- ± 0.5-fold larger in the ccpC mutant than in the wild-type strain. (B) Transcripts of the citC gene. The primers used were OMM97 and OMM98 (Fig. 2). In two independent experiments, the amount of citC transcript (normalized to the amount of rRNA) was 3- ± 0.2-fold larger in the ccpC mutant than in the wild-type strain.

Interaction of CcpC with the P2 promoter region in vitro. To study the interaction of CcpC with the P1 and P2 promoter regions, we purified a hexahistidine-tagged version of the L. monocytogenes CcpC protein (24) and tested its binding to these regions by performing gel mobility shift assays. A 237-bp DNA fragment carrying the P2 promoter showed retarded mobility when it was incubated with CcpC-His₆. CcpC was able to shift 50% of the DNA at a protein concentration of 3.9 to 7.8 nM (Fig. 7). By contrast, CcpC did not interact with the P1 promoter region in vitro (Fig. 7). These results are consistent with the hypothesis that there is CcpC-dependent regulation of the P2 promoter and CcpC independence of transcription initiation at the P1 promoter.

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FIG. 7. Gel mobility shift assay of the interaction of L. monocytogenes CcpC with the P1 and P2 promoters. CcpC-His6 was incubated with 32P-labeled DNA fragments containing either the lmo1568 (left panel) or lmo1569 (right panel) promoter region. The concentrations of CcpC-His6 used in the reactions were as follows: for lmo1568, 0, 3.9, 7.8, 15.6, 31.3, 62.5, 125, 250, 500, and 1,000 nM (lanes 1 to 10, respectively); and for lmo1569, 0, 7.8, 15.6, 31.3, 62.5, 125, 250, 500, and 1,000 nM (lanes 1 to 9, respectively). See Materials and Methods for details.

The specific site of CcpC binding in the P2 promoter region was determined by performing a DNase I footprinting experiment. In the presence of CcpC-His₆, the region from position –9 to position –61 with respect to the tsp was protected against DNase I digestion (Fig. 8). Within this region there is a dyad symmetry element that matches elements found in the CcpC-binding site of the L. monocytogenes citB promoter and in B. subtilis binding sites for CcpC. A half-dyad element is located 17 bp from the upstream end of the dyad element. Similar orientations of full and half sites have been found in other CcpC-binding sites (23).

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FIG. 8. DNase I footprinting assay of the interaction between CcpC-His6 and the P2 promoter region. A 32P-labeled DNA fragment corresponding to the lmo1568 promoter region was incubated with various amounts of CcpC-His6 prior to DNase I digestion. The concentrations of CcpC-His6 (in nM) used in the reactions are indicated above the lanes. The vertical bar indicates the region protected from DNase I digestion by CcpC-His6. In the sequence of the lmo1568 promoter region shown on the right, the region protected by CcpC-His6 from DNase I digestion (positions –9 to –61 with respect to the transcription start site, as deduced from other experiments in which a DNA sequence ladder was generated) is indicated by bold type. The consensus recognition sequences for CcpC are indicated by uppercase letters. The transcription start site is indicated by a single uppercase T, and the –10 and –35 regions are also indicated. The start codon of lmo1568 is indicated by italics.

Regulation of citZ expression by different carbon sources. It is possible that we did not observe any promoter activity in the region immediately upstream of the citZ gene because it is masked by CcpA-independent catabolite repression. BHI medium, in which the cells were grown for these experiments, contains glucose at a concentration of 0.2% (6). To address this issue, we tested the strains carrying fusion C or D in defined medium with different carbon sources. These carbon sources included the catabolite-repressing sugars glucose and mannose, as well as glycerol and citrate, which do not cause catabolite repression (21). In defined medium containing any carbon source tested except citrate, the strains grew with similar doubling times and reached stationary phase (OD₆₀₀, 1.0). The strains grew poorly in citrate medium, and the maximum OD₆₀₀ was 0.1 to 0.2 even after prolonged incubation (48 h). The poor growth in citrate probably reflected poor uptake of citrate by the cells. Moreover, due to the split nature of the Krebs cycle in L. monocytogenes, citrate cannot be used to generate oxaloacetate, which is used in gluconeogenesis as a source of hexose and pentose as precursors of nucleic acids and cell envelope polymers. L. monocytogenes also appears to lack citrate lyase and isocitrate lyase enzymes, which could be used alternatively for synthesizing oxaloacetate. The strains containing the D fusion had the same low levels of β-galactosidase activity in all media tested and in both null backgrounds, reinforcing the idea that the region immediately upstream of the citZ gene does not contain a promoter that is masked by the repressive effect of glucose (Table 5). In addition, we found that CcpC-mediated repression of the C fusion occurred at the same level under all the conditions tested, suggesting that no other carbon source-responsive regulator acts along with CcpC to regulate citZ expression.

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TABLE 5. Effects of different carbon sources on citZ expression in L. monocytogenes

Regulation of citZ expression by glutamine. We measured the expression of the lmo1568-lacZ fusion in strain LMM16 grown at 37°C in BHI medium supplemented with glutamine, glucose, or citrate or combinations of nutrients. Neither glucose (which is present at a concentration of 0.2% in unsupplemented BHI medium [6]) nor citrate altered the expression of β-galactosidase (Fig. 9). When BHI medium was supplemented with glutamine or combinations of glutamine and glucose or citrate, lacZ expression was repressed about fivefold (Fig. 9). Repression in glutamine-containing medium was overcome by a ccpC mutation (Fig. 9).

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FIG. 9. Effect of medium composition on expression of a lacZ fusion in L. monocytogenes. Strains LMM16 (ccpC+) (gray bars) and LMM21 (ccpC::spc) (black bars), both of which carried an lmo1568-lacZ fusion, were grown at 37°C in BHI medium supplemented with 0.5% glucose, 1.6 mg glutamine per ml, or 0.5% citrate or with combinations of these supplements. β-Galactosidase activity was measured using samples taken at various points during the exponential phase of growth. The values are the averages of two experiments.

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Our work showed that the L. monocytogenes citZ gene is part of a complex operon along with the upstream genes lmo1569 and lmo1568 and the downstream gene citC. While the RT-PCR data indicate that citZ and citC are cotranscribed, it was not possible to deduce from our results whether citC also has an independent promoter and whether, if there is a promoter, it is a target of CcpC. (The fact that a ccpC mutation derepressed citZ transcription ninefold but derepressed citC transcription only threefold suggests that the citC gene may have a second, CcpC-independent promoter.) lmo1569 and lmo1568 are annotated as genes with unknown functions (14). A homology search revealed that they are similar only to other genes with unknown functions. Why these genes are coregulated with citZ remains unknown, but it is interesting that ytwI, the gene immediately upstream of citZ in B. subtilis, is a close homolog (57% identity) of lmo1568 (Fig. 10). The closest homolog of lmo1569, ytzA, is the third gene upstream of citZ in B. subtilis (Fig. 10). The possibility of cotranscription of ytzA or ytwI with citZ has not been addressed, but the region between ytwI and citZ contains an apparent transcription terminator (Fig. 10).

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FIG. 10. Synteny of the citZ loci in L. monocytogenes and B. subtilis. The designations of the L. monocytogenes and B. subtilis genes are the designations used in the ListiList (http://genolist.pasteur.fr/ListiList/) and SubtiList (http://genolist.pasteur.fr/SubtiList/) databases. The arrows indicate genes and their orientations, and the balloons indicate the locations of putative transcription termination sites. Dotted lines connect genes whose products share sequence similarity.

CcpC binds to a single site but acts as a direct repressor of the lmo1568, citZ, and citC genes in two different ways. Two promoters drive the expression of these genes; CcpC represses P2, presumably by competition with RNA polymerase for access to the promoter, and aborts transcripts from P1 by acting as a roadblock to RNA polymerase molecules that initiate transcription at P1. By performing a footprinting analysis we located the CcpC-binding site, which includes a dyad symmetry element (ATAAN₇TTAT) and a half-copy of the dyad symmetry element (TTAT). This pattern of dyad and half-dyad elements is typical of CcpC-binding sites in B. subtilis and L. monocytogenes (23, 24).

Transcription of citZ is partially constitutive during growth in BHI medium, because even when CcpC is active enough to repress P2, some transcripts from P1 pass through the CcpC roadblock. As a result, the cell is primed to synthesize some citrate and to fully derepress the TCA branch when the substrates for citrate synthase, oxaloacetate, and acetyl-CoA become available.

Is there a second mechanism that couples TCA branch expression to the physiological state of the cell? In B. subtilis, expression of citZ is regulated by the carbon source through the activity of CcpA (25). We found no effect of the carbon source on citZ expression, no role for CcpA in L. monocytogenes citZ regulation, and no sequence similar to the consensus CcpA-binding site in the region upstream of or within the L. monocytogenes citZ coding sequence.

Instead, L. monocytogenes citZ expression is repressed in a CcpC-dependent manner in BHI medium containing glutamine. Given the split Krebs cycle in this bacterium, 2-ketoglutarate is the only end product of the TCA branch, and the principal role of 2-ketoglutarate is to provide the carbon skeleton for glutamate and glutamine. Hence, repression of the lmo1569-lmo1568-citZ operon by CcpC in the presence of excess glutamine (which would presumably be reflected in high levels of 2-ketoglutarate) would allow efficient use of these enzymes. In B. subtilis, citrate synthase enzyme activity is feedback inhibited by 2-ketoglutarate (H. J. Kim, unpublished). If 2-ketoglutarate has the same effect on L. monocytogenes citrate synthase, its accumulation would affect the activity of CcpC indirectly by reducing the synthesis of citrate.

The dual regulation of citZ expression by CcpC presumably helps L. monocytogenes fine-tune the expression of TCA branch enzymes according to the metabolic state of the cell. Citrate synthase, the first enzyme in this pathway, controls the downstream enzymes aconitase and isocitrate dehydrogenase in two ways. First, citrate synthase produces citrate, the substrate for aconitase. Second, citrate antagonizes the interaction of CcpC with the citB promoter, thus inducing citB transcription (24). If it is cotranscribed with citZ, citC would also be under CcpC control. Hence, CcpC is likely to coordinate the regulation of all three enzymes of the oxidative branch of the Krebs cycle in response to the intracellular concentration of citrate.

 
We thank H. J. Kim for providing plasmid pHK23 and B. R. Belitsky for helpful discussions and a critical review of the manuscript.

This work was supported by research grant GM036718 from the Public Health Service.

 
* Corresponding author. Mailing address: Department of Molecular Biology and Microbiology, Tufts University School of Medicine, 136 Harrison Avenue, Boston, MA 02111. Phone: (617) 636-6761. Fax: (617) 636-0337. E-mail: linc.sonenshein{at}tufts.edu

Published ahead of print on 14 November 2008.

Present address: Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and Universidad de Sevilla, E-41092 Seville, Spain.

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Journal of Bacteriology, February 2009, p. 862-872, Vol. 191, No. 3
0021-9193/09/$08.00+0     doi:10.1128/JB.01384-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

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