AAC
Home Help [Feedback] [For Subscribers] [Archive] [Search] [Contents]
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Seidl, K.
Right arrow Articles by Bischoff, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Seidl, K.
Right arrow Articles by Bischoff, M.

 Previous Article  |  Next Article 

Antimicrobial Agents and Chemotherapy, April 2006, p. 1183-1194, Vol. 50, No. 4
0066-4804/06/$08.00+0     doi:10.1128/AAC.50.4.1183-1194.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Staphylococcus aureus CcpA Affects Virulence Determinant Production and Antibiotic Resistance

Kati Seidl,1,{dagger} Martin Stucki,1,{dagger} Martin Ruegg,1 Christiane Goerke,2 Christiane Wolz,2 Llinos Harris,3 Brigitte Berger-Bächi,1 and Markus Bischoff1*

Department of Medical Microbiology, University of Zürich, Zürich, Switzerland,1 Institute for Medical Microbiology and Hygiene, University Hospital Tübingen, Tübingen, Germany,2 AO Research Institute, Davos, Switzerland3

Received 27 October 2005/ Returned for modification 11 January 2006/ Accepted 3 February 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results and Discussion
 References
 
Carbon catabolite protein A (CcpA) is known to function as a major regulator of gene expression in different gram-positive organisms. Deletion of the ccpA homologue (saCOL1786) in Staphylococcus aureus was found to affect growth, glucose metabolization, and transcription of selected virulence determinants. In liquid culture, deletion of CcpA decreased the growth rate and yield; however, the effect was only transient during the exponential-growth phase as long as glucose was present in the medium. Depletion of glucose and production of lactate was delayed, while the level of excretion of acetate was less affected and was even higher in the mutant culture. On solid medium, in contrast, growth of the {Delta}ccpA mutant resulted in smaller colonies containing a lower number of CFU per colony. Deletion of CcpA had an effect on the expression of important virulence factors of S. aureus by down-regulating RNAIII, the effector molecule of the agr locus, and altering the transcription patterns of hla, encoding {alpha}-hemolysin, and spa, encoding protein A. CcpA inactivation markedly reduced the oxacillin resistance levels in the highly methicillin-resistant S. aureus strain COLn and the teicoplanin resistance level in a glycopeptide-intermediate-resistant S. aureus strain. The presence of CcpA in the capsular polysaccharide serotype 5 (CP5)-producing strain Newman abolished capsule formation and decreased cap operon transcription in the presence of glucose. The staphylococcal CcpA thus not only is involved in the regulation of carbon metabolism but seems to function as a modulator of virulence gene expression as well.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results and Discussion
 References
 
Carbon catabolite repression (CCR) in bacteria is a widespread, global regulatory phenomenon that allows modulation of the expression of genes and operons involved in carbon utilization and metabolization in the presence of preferred carbon source(s). In CCR, the presence of a preferred carbon source represses the expression of genes and operons whose products are involved in the metabolism of alternative, less-preferred carbon sources. In low-GC gram-positive bacteria, CCR is achieved via transcriptional control, inducer exclusion, and induction prevention (reviewed in references 55 and 60). In this group of bacteria, a common mechanism for transcriptional control has evolved that is mediated via the proteins phosphotransferase HPr, the bifunctional HPr kinase-phosphatase (HPrK/P), and the pleiotropic regulator CcpA (catabolite control protein A). CCR in Bacillus subtilis has been studied extensively and is thought to serve as the prototype of CCR-regulated gene expression in gram-positive organisms (reviewed in reference 52). In B. subtilis, regulation of transcription of catabolite-repressive genes is exerted mainly through the binding of CcpA to specific cis-acting DNA sequences called catabolite-responsive elements (CREs). The DNA-binding activity of CcpA itself is triggered by HPr or its regulatory paralog Crh, which, in the presence of glucose, are phosphorylated by HPrK/P on regulatory seryl residues, in which state they act as cofactors for CcpA. Depending on the localization of the CRE, CcpA may function either as an activator or as a repressor of gene expression. Whole-transcriptome analyses suggest that 10% of all genes in B. subtilis are affected in their regulation by glucose by a factor of more than 3, with repressed genes outnumbering activated genes by three to one (3, 39). The majority (80%) of these genes depend on CcpA for regulation, and a recent study indicated that CcpA required interaction with RNA polymerase to inhibit transcription (31).

Although CCR by the catabolite control protein CcpA has been demonstrated in Staphylococcus xylosus (reviewed in reference 26), only a little is known about this element in the closely related, pathogenic Staphylococcus aureus. However, indications that glucose affects gene expression in S. aureus (12, 24, 27, 44, 45, 49), the identification of a potential CRE in the promoter region of the glucose-repressible pckA (49) that is highly homologous to the CRE consensus of B. subtilis (38), and the presence of HPr (SaCOL1091), HPrK (SaCOL0825), and CcpA (SaCOL1786) homologues in S. aureus suggest that a similar mechanism might be present in this pathogen.

Site-directed inactivation of ccpA showed here that the lack of CcpA, although causing only minor effects on growth of S. aureus, affected oxacillin and glycopeptide resistance and had a significant impact on the ability of S. aureus to express virulence factors such as RNAIII, hla, and spa in either the presence or absence of glucose.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results and Discussion
 References
 
Bacterial strains and culture conditions. The bacterial strains and relevant phenotypes are listed in Table 1. When not otherwise specified, bacteria were grown in Luria-Bertani medium (Difco Laboratories, Detroit, Mich.) buffered using 50 mM HEPES (4-2-hydroxyethyl-1-piperazineethanesulfonic acid [pH 7.5]) and a flask volume/culture volume ratio of 5:1 at 200 rpm and 37°C. Where indicated, mutant strains were grown on antibiotic-supplemented media containing 100 µg of ampicillin, 10 µg of erythromycin, or 10 µg of tetracycline per ml.


View this table:
[in this window]
[in a new window]
  		TABLE 1. Strains and plasmids used in this study

 
DNA manipulations. DNA sequencing, PCR, and plasmid isolation were performed using standard procedures (1) or according to manufacturers' instructions.

Construction of S. aureus {Delta}ccpA. A 2.7-kp fragment containing the ccpA gene and its flanking regions was amplified by PCR from chromosomal DNA of S. aureus COL by use of primer pair ccpABamHI-F/ccpAEcoRI-R (Table 2), digested, and cloned into the BamHI/EcoRI-digested vector pEC1 (4) to generate plasmid pMR1 (Fig. 1). The plasmid was used in a second step to amplify a 5.8-kb fragment containing the pEC1 backbone and the regions flanking ccpA by use of the primer pair ccpAko-F/ccpAko-R. The 5.8-kb fragment was ligated to a PCR-amplified and phosphorylated 1.7-kb tet(L) cassette obtained from plasmid pBT (17) by use of primer pair tetL-F/tetL-R to generate the suicide vector pMR2, and the plasmid was subsequently electroporated into S. aureus RN4220. Mutants with the allelic replacement were selected for tetracycline resistance and screened for loss of erythromycin resistance, yielding MST04 (RN4220 ccpA::tet[L]), which was subsequently used as a donor for transducing the ccpA deletion into other S. aureus strains.


View this table:
[in this window]
[in a new window]
  		TABLE 2. Primers used in this study

 

Figure 1
View larger version (16K):
[in this window]
[in a new window]
  		FIG. 1. Schematic representation of the ccpA region of S. aureus and of the strategy used to obtain MST23 (COLn {Delta}ccpA). The genetic organization of the S. aureus COL ccpA region (A), pMR1 (B), pMR2 (C), and MST23 (D) is shown. Open reading frame notations and nucleotide numbers correspond to those of the respective genomic regions of strain COL (GenBank accession no. NC_002951). Primers used to amplify the ccpA region (A), the ccpA-flanking regions including the backbone of pEC1 (B), and the respective PCR products (dotted lines) are indicated.

 
Construction of plasmid pMST1. A 1.76-kp fragment, covering the ccpA gene and 770 bp of its upstream region, was amplified by PCR from chromosomal DNA of S. aureus COL by use of primer pair MST12/MST14 (Table 2), digested, and cloned into the BamHI/XbaI site of vector pAW17 (46) to generate plasmid pMST1. The plasmid was first electroporated into S. aureus RN4220 and then transduced into the various ccpA mutants.

Determination of acetate, glucose, and lactate levels. Aliquots (2 ml) of bacterial cultures were harvested at the indicated time points and centrifuged for 2 min at 16,000 x g. The supernatants were incubated at 80°C for 15 min and stored at –20°C until use. Acetate, glucose, and lactate levels were determined with kits from R-Biopharm (Darmstadt, Germany) according to the manufacturer's directions.

Adherence studies. S. aureus strains were grown in brain heart infusion (BHI) at 37°C in a shaking water bath at 250 rpm for 3 h to midexponential phase (A600 of 1) and used to inoculate 1 ml prewarmed BHI in 24-well plates containing presterilized polyethylene terephthalate (Thermanox) 13-mm disks (Life Technologies, Basel, Switzerland) to a starting A600 of 0.05 and incubated without shaking at 37°C for 15 h before fixation for scanning electron microscopy. Fixation and electron microscopy were carried out as described earlier (21).

Susceptibility testing. For testing antibiotic resistance on gradient plates, plates were prepared by pouring 35 ml of LB agar containing 1,000 µg ml–1 oxacillin into a rectangular inoculum dish (Dynatech, Dübendorf, Switzerland) that was reposed on one side to allow the solidifying LB agar to form a wedge. In a second step, the solidified LB agar plate was placed horizontally and 35 ml of LB agar lacking the antibiotic was poured onto the first layer and allowed to solidify for 3 h, thereby allowing the antibiotic of the lower layer to diffuse into the upper layer to form a gradient. Cells of the strains to be tested were resuspended in physiological NaCl solution to a density of 0.5 McFarland (McF 0.5) and swabbed onto the plate along the gradient. Growth was read after 24 h and 48 h of incubation at 35°C. For E-tests, bacterial suspensions (McF 0.5 for oxacillin and McF 2 for teicoplanin) were swabbed onto the surface of either Mueller-Hinton agar plates supplemented with 2% NaCl (oxacillin) or BHI agar plates (teicoplanin) according to the manufacturer's instructions (AB-Biodisk, Solna, Sweden). Determinations of MICs by broth microdilution were performed as recommended by the CLSI (formerly NCCLS) (9). For population analysis profiles, appropriate dilutions of an overnight culture were plated on LB agar plates containing increasing concentrations of oxacillin (0 to 2,048 µg ml–1) and the numbers of CFU were determined after 48 h of incubation at 35°C.

Northern blot analyses. For in vitro growth studies, overnight cultures of S. aureus were diluted 1:100 into fresh prewarmed, HEPES-buffered LB medium (pH 7.5). Cells were grown either with or without 10 mM glucose, samples were removed from the cultures after 1, 3, 5, and 8 h of growth and centrifuged at 13,000 x g and 4°C for 5 min, and the cell sediments were snap-frozen in liquid nitrogen. In a second approach, cells were grown in HEPES-buffered LB to an A600 of 1, the cultures were split in two, and 10 mM glucose added to one half. Aliquots were sampled at 0, 10, 20, and 30 min and harvested at 16,000 x g at room temperature for 1 min, and the cell sediments were snap-frozen in liquid nitrogen. Total RNAs were isolated according to the method of Cheung et al. (6). Blotting, hybridization and labeling were performed as previously described (17). The intensities of the 23S and 16S rRNA bands stained with ethidium bromide were verified to be equivalent in all the samples before transfer. Primer pairs ccpA-F/ccpA-R, hla-F/hla-R, pckA-F/pckA-R, RNAIII-F/RNAIII-R, and spa-F/spa-R (Table 2) were used, respectively, to generate digoxigenin-labeled ccpA-, hla-, pckA-, RNAIII-, and spa-specific probes by PCR labeling. Data shown were confirmed in at least two independent experiments.

RNA quantification by LightCycler RT-PCR. For quantification of transcripts by LightCycler reverse transcription-PCR (RT-PCR), RNA preparations were performed as described earlier (19). Briefly, approximately 109 S. aureus cells were lysed in 1 ml TRIzol reagent (Invitrogen Life Technologies, Karlsruhe, Germany) with 0.5 ml of zirconia-silica beads (BioSpec Products, Bartlesville, OK) (0.1 mm diameter) in a high-speed homogenizer (Savant Instruments, Farmingdale, N.Y.). RNA was isolated as described in the instructions provided by the manufacturer of TRIzol. Contaminating DNA was degraded by digesting RNA samples with DNase as described before (19). Sequence-specific RNA standards for quantitative RT-PCR were engineered as described previously (20) using primer pair T7-gyr/gyr864 for gyrB and primer pair T7-capA59/capA710 for capA (Table 2).

LightCycler RT-PCR was carried out using a LightCycler RNA amplification kit for hybridization probes or with a LightCycler RNA amplification kit and SYBR green I (Roche Biochemicals). Master mixes were prepared following the manufacturer's instructions using the oligonucleotides gyr297/gyr574 for gyrB (20) and primer pair capA167/capA501 for capA (Table 2). Specific primers were selected in such a way that they bound to an internal part of the respective RNA standard. Standard curves were generated using 10-fold serial dilutions (104 to 108 copies/µl) of the specific RNA standards. The number of copies of each sample transcript was then determined with the aid of the LightCycler software. At least two independent RT-PCR runs were performed for each sample. The specificity of the PCR was verified by ethidium bromide staining on 3% agarose gels. To check for DNA contamination each sample and RNA standard was subjected to PCR using a LightCycler DNA amplification kit and SYBR green I (Roche Biochemicals). No amplification product was detectable in any of the cases.

Capsular polysaccharide serotype 5 (CP5) detection by indirect immunofluorescence. CP5 production was determined from cultures grown for 24 h in LB medium. Slides with heat-fixed bacteria were washed three times with phosphate-buffered saline (PBS)-0.05% Tween 20 and incubated with 0.2 mg ml–1 human immunoglobulin (IgG) (Sigma, Deisenhofen, Germany) diluted in PBS-0.05% Tween 20 for 30 min to prevent unspecific binding of IgG by cell-wall-associated protein A. The slides were incubated with mouse IgM monoclonal antibodies to CP5 (22) diluted 1:50 in PBS-0.05% Tween 20 for 1 h followed by incubation with CY3-conjugated anti-mouse F(ab)2 fragment (Dianova, Hamburg, Germany) diluted 1:500 in PBS-0.05% Tween 20 for 1 h. Bacteria were stained with 4',6'-diamidino-2-phenylindole (DAPI) (2 µg ml–1) for 5 min, washed three times with water, and air dried. The slides were then mounted with fluorescent mounting medium (DakoCytomation, Hamburg, Germany), and positively stained bacteria were detected using fluorescence microscopy.


   RESULTS AND DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussion
References
 
Growth and carbohydrate utilization of S. aureus {Delta}ccpA mutants. The ccpA gene was deleted in strain RN4220 by allelic replacement (Fig. 1) and transduced therefrom into the capsular polysaccharide serotype 5 (CP5)-producing strain Newman and the methicillin-resistant strain COLn, yielding strains MST14 and MST23, respectively. Analysis of the transcript sizes of the genes surrounding ccpA, i.e., acuAC, encoding acetoin utilization protein A and C, and saCOL1787, thought to encode a chorismate mutase-phospho-2-dehydro-3-deoxyheptonate aldolase, yielded identical patterns for Newman and its {Delta}ccpA mutant MST23, indicating that the genetic manipulations leading to the deletion of ccpA did not affect the integrity of the adjacent genes (data not shown).

The growth characteristics and metabolite production of strain COLn and its {Delta}ccpA mutant MST23 were monitored in HEPES-buffered LB supplemented with 10 mM glucose. Comparison of the wild-type and mutant cultures revealed a clear difference only for the mid- to late-exponential-growth phases (i.e., hours 3 to 6). During these growth periods, the mutant displayed slower growth, yielding cell densities lagging approximately 1 h behind the wild-type culture densities. However, the mutant culture reached almost the same A600 values after 10 h of growth (Fig. 2A), and no differences in A600 values were observable between the wild type and the mutant after 16 and 24 h of growth (data not shown). CFU determinations of the growing cultures showed the same tendency, being different only during the mid- to late-exponential-growth phase, and again, no differences in growth yield were observed after an incubation period of 24 h (data not shown). During the exponential-growth phase, differences in growth rate were also visible with respect to doubling times; the wild type yielded a significantly lower doubling time (44.34 ± 0.65 min) than MST23 (51.67 ± 0.75 min; P < 0.01). A clearer difference was observed between the wild type and the mutant on solid media. MST23 produced significantly smaller colonies on sheep blood agar or on Muller-Hinton plates, with lower CFU numbers per colony than COLn after 48 h of incubation (Fig. 2B). No differences were observed when COLn and MST23 were checked for cell size and adherence properties (Fig. 2D), signaling that the differences in growth on solid media were likely to be due to a reduced growth rate of the mutant.


Figure 2
View larger version (29K):
[in this window]
[in a new window]
  		FIG. 2. Growth characteristics of COLn (squares) and its {Delta}ccpA mutant MST23 (triangles) in LB supplemented with 10 mM glucose. (A) Absorbances at 600 nm (A600) and CFU over the growth cycle. (B) Cfu per colony of COLn and MST23 grown on sheep blood agar for 48 h at 35°C. (C) Glucose, acetate, and lactate concentrations in the culture supernatants corresponding to panel A. The data presented are mean values of three independent experiments. (D) Scanning electron microscopy images of COLn and MST23 adhering to polyethylene terephthalate (Thermanox) disks.

 
In addition to their growth kinetics, growing COLn and MST23 cultures were further analyzed for glucose metabolism and breakdown. The glucose level of the wild-type culture visibly decreased from hour 3 on and glucose was depleted after hour 7. The glucose level of the MST23 culture started to decrease from hour 5 and was depleted only after 9 h of growth (Fig. 2C). Simultaneously with glucose degradation, the lactate level of the wild-type culture started to increase from hour 3 on, reached its maximum around hour 6, and significantly decreased thereafter. In contrast, the lactate level of the mutant culture started to increase only after 7 h of growth, representing a delay of more than 2 h compared with glucose consumption, and reached its maximum after 10 h. Interestingly, only slight differences were observable between the wild type and the mutant with respect to acetate accumulation; the acetate level seen with the mutant culture preceded that of the wild-type culture by approximately 0.5 h. After 24 h of growth, no lactate was present in either wild-type or mutant media anymore, while acetate levels were reduced to similar amounts, namely, 3.8 ± 0.86 mM for COLn and 3.75 ± 0.76 mM for MST23, signaling that the ccpA mutation did not affect the ability of the mutant to assimilate acetate and lactate that was excreted into the media during earlier growth stages.

Although the growth rate and yield of the ccpA mutant MST23 were only slightly affected by the deletion, the slower glucose consumption and delayed lactate secretion of MST23, paired with the slightly increased acetate production, suggested that CcpA seemed to exert a positive effect on lactate production and secretion whereas acetate production and secretion seemed to be negatively affected by this protein, taking into account that MST23 possessed lower cell densities and consumed glucose slower than the wild type at almost all time points analyzed. Preliminary Northern analyses supported the hypothesis that CcpA might stimulate lactate and suppress acetate formation in the presence of glucose. Expression of ldh1 (saCOL0222), thought to encode L-lactate dehydrogenase 1 (EC 1.1.1.27) that is believed to catalyze the conversion from pyruvate to lactate when S. aureus is grown in the presence of a rapidly catabolizable carbon source under anaerobic conditions, was highly induced in wild-type cells grown in the presence of glucose but was not detectable in the absence of glucose or in the {Delta}ccpA mutant under either growth condition. Expression of genes encoding enzymes involved in acetate formation, such as pdhABCD, encoding components of the pyruvate dehydrogenase multienzyme complex thought to catalyze the conversion of pyruvate to acetyl coenzyme A, and the adhE homologue (saCOL0135) thought to encode alcohol-acetaldehyde dehydrogenase (EC 1.2.1.10) that catalyzes the formation of acetyl-coenzyme A to acetaldehyde, on the other hand, was found to be repressed in the wild type in the presence of glucose, while no differences in expression were observed in the {Delta}ccpA mutant in either the presence or absence of glucose (K. Seidl, unpublished data).

Effect of ccpA on antibiotic resistance. CcpA was among a series of auxiliary factors reported to reduce methicillin resistance in strain COL upon Tn551 inactivation (11). We demonstrated here a fourfold reduction of the oxacillin MIC for strain COLn upon ccpA inactivation and full restoration to its original level by trans-complementation with the wild-type ccpA allele under the control of its native promoter (Fig. 3A), thus confirming experimentally that the effect was indeed solely CcpA dependent. Moreover, the population analysis profile (Fig. 3B) showed that the homogenous oxacillin resistance of strain COLn was reduced to heterogeneous resistance by ccpA inactivation.


Figure 3
View larger version (17K):
[in this window]
[in a new window]
  		FIG. 3. Susceptibility of COLn and NM143 and their isogenic {Delta}ccpA mutants. (A) Effects of ccpA inactivation and complementation on oxacillin resistance. The methicillin-resistant strain COLn, its {Delta}ccpA mutant MST23, control strains MST31 (COLn pAW17) and MST35 [COLn {Delta}ccpA::tet(L) complemented with control plasmid pAW17], and strain MST36 [COLn {Delta}ccpA::tet(L) complemented with plasmid pMST1] were swabbed along a plate containing an oxacillin gradient. The corresponding MICs of oxacillin (in micrograms per milliliter as determined by E-Test and broth microdilution) for the strains are indicated. (B) Population analysis profiles of COLn (squares) and MST23 [COLn {Delta}ccpA::tet(L); triangles] on oxacillin. (C) Population analysis profiles of the step-selected glycopeptide-intermediate-resistant S. aureus derivative NM143 (squares) and its {Delta}ccpA mutant KS30 (triangles) on teicoplanin.

 
Transduction of ccpA::tet(L) into NM143, the step-selected teicoplanin-resistant derivative of strain Newman, yielding strain KS30, had a similar negative effect on teicoplanin resistance. The ccpA inactivation reduced the MIC of teicoplanin from 24 to 12 µg ml–1, and the population analysis profile teicoplanin showed that the number of more highly resistant variants was reduced by a magnitude of over 103 (Fig. 3C).

Effect of glucose and ccpA on virulence determinant production. Previous studies showed that fermentation of glucose and/or the accompanying decrease in pH affected expression of the global regulator agr and of virulence factors such as {alpha}-hemolysin (hla) and the staphylococcal enterotoxins A, B, and C (sea, seb, and sec) (12, 24, 27, 44, 45). More recently, Weinrick and coworkers (62) showed that mildly acidic conditions (pH 5.5) influenced the expression of a variety of genes, including agr, hla, and spa, encoding protein A, and concluded that changes in staphylococcal gene expression formerly thought to represent a glucose effect might be largely the result of declining pH of the growth medium due to the fermentation of the supplemented carbon source. The effect of glucose on hla and spa expression is further complicated by the fact that both genes are affected by a complex regulatory network including agr and further regulatory elements such as ArlRS (15), MgrA (25), MsrR (46), Rot (37, 47), SaeRS (18, 20), SarA (reference 8 and references therein), SarS (7, 36, 53), SarT (48), SvrA (16), TcaR (36), and the alternative transcription factor {sigma}B (2, 20, 23), with rot, sae and tcaR expression being pH dependent as well (62).

To elucidate whether agr, hla, and spa transcription was affected by glucose independently from the fermentation-dependent pH and to find out whether CcpA may be involved in mediating such a glucose effect, we monitored the expression levels of these genes in COLn and its ccpA mutant MST23, grown in buffered LB in the presence or absence of glucose (Fig. 4). pckA, encoding a phosphoenolpyruvate carboxykinase, shown to be affected by glucose, and predicted to be regulated by CcpA in S. aureus (49), was included in this study as well. The 50 mM HEPES concentration used here to buffer the medium to pH 7.5 had no inhibiting effect on the growth kinetics and kept the pH fairly constant (Fig. 4A), while concentrations higher than 50 mM were growth inhibitory (data not shown). No changes in pH were observed when COLn and MST23 were grown in LB in the absence of glucose. In the glucose-supplemented wild-type culture, the pH started to drop slightly after 5 h to a final pH of 7, while the pH of the glucose-supplemented MST23 culture dropped from hour 7 on to pH 7.25. Growth of COLn in unbuffered glucose-supplemented LB medium would have caused a drop in pH to 5.5 already after 3 h of growth (data not shown). Addition of glucose produced a higher growth rate and growth yield in the wild type from hour 5 on (Fig. 4A). Surprisingly, glucose seemed to have a slightly inhibiting effect on the growth rate of the mutant MST23 during exponential-growth phase, as seen in the lower A600 values for MST23 grown in LB supplemented with glucose compared to those in LB alone; this effect might be due to a slightly increased lag phase.


Figure 4
View larger version (64K):
[in this window]
[in a new window]
  		FIG. 4. Northern blot analyses of COLn and its {Delta}ccpA mutant MST23 during growth. (A) Growth characteristics of COLn (squares) and MST23 (triangles) grown in HEPES-buffered LB (open symbols) and HEPES-buffered LB supplemented with 10 mM glucose (closed symbols). At 1-h intervals, an aliquot (2 ml) was removed, the absorbance at 600 nm was measured, and the pH in the culture supernatants (circles) was determined. The results presented are mean values of at least three independent experiments. Time points of sampling for the Northern blot analyses are indicated by arrows. (B) Transcription of ccpA, hla, pckA, RNAIII, and spa in COLn and MST23 during growth in HEPES-buffered LB (–Glc) and in HEPES-buffered LB supplemented with 10 mM glucose (+Glc). Relevant transcript sizes and time points of sampling are indicated. Ethidium bromide-stained 16S rRNA patterns are shown as an indication of RNA loading.

 
Expression of ccpA, pckA, RNAIII, spa, and hla was monitored after 1, 3, 5, and 8 h of growth. In COLn grown in LB, expression of ccpA was found to be fairly constant at all growth stages analyzed (Fig. 4B). Supplementation of glucose seemed to increase the expression of this gene during the early growth stages (i.e., 1 to 3 h), suggesting that expression of ccpA might be positively affected by glucose, a phenomenon that has been observed for ccpA in other gram-positive organisms (14). No ccpA transcripts were detectable in MST23, confirming that the deletion had occurred as intended. Expression of pckA increased with growth and was clearly higher when COLn cells were grown in the absence of glucose, confirming previous findings indicating that pckA expression is negatively affected by glucose (49). MST23, on the other hand, already produced pckA transcripts at a rather constant and high level early in growth. No significant differences in pckA expression were observed in the presence or absence of glucose, signaling that the effect of glucose on pckA expression was mediated via CcpA, as has been suggested by Scovill et al. (49). The idea of CcpA-dependent repression of pckA transcription is further supported by the presence of a putative CRE in the promoter region of pckA (49) that almost perfectly matched (17/18 nt) the CRE consensus of B. subtilis (38), suggesting that the CRE sequences might be similar in the two organisms.

Expression of RNAIII from the agr locus is known to increase during later growth stages (reference 63 and references within). Accordingly, in COLn cells grown in LB, RNAIII transcripts were first detected after 5 h of growth and increased with time. Unexpectedly, in glucose-grown cells, a strong increase in RNAIII expression occurred at hour 8, contrary to the findings of Regassa et al. (45), who reported unchanged RNAIII transcript levels for S. aureus cells grown in a fermentor in either the presence or absence of glucose at a constant pH. However, the discrepancy in RNAIII expression might be explained by the differing growth conditions, since Regassa and coworkers kept the glucose concentration at a constant level of 100 mM. It is noteworthy that the increase in RNAIII expression was detected at a time point when all glucose was exhausted from the medium, indicating that the depletion of glucose might have triggered a signal that induced RNAIII transcription, which was not present under the conditions used by Regassa et al. (45). The slight decrease in pH observed in the later growth stages of the wild-type culture in response to glucose (Fig. 4A) could be excluded as a reason for the strong induction of RNAIII transcription, since control experiments performed using a fermenter and an equivalent experimental setup allowed the pH to be kept constant and yielded the same induction pattern (data not shown).

No RNAIII transcripts were detectable in MST23 cells grown in LB, and addition of glucose yielded only traces of RNAIII transcripts after 8 h of growth. The clear differences in RNAIII transcript levels observed for COLn and MST23 suggest that the presence of a functional CcpA had a positive effect on RNAIII expression. However, this effect was likely to be indirect, since the screening of the agr locus did not reveal any apparent CRE in this genomic region that would fit with the CRE consensus of B. subtilis (38). Expression of spa was noticeably affected by glucose in the wild type. While spa transcripts were clearly detectable in COLn during later growth stages (hours 5 to 8) in the absence of glucose, spa transcripts were hardly detectable in the presence of glucose (Fig. 4B). However, in MST23, spa transcripts were detectable over the whole growth cycle independently of the presence or absence of glucose, although the spa transcription profiles seemed to differ between these two conditions to a certain degree. While spa expression in MST23 grown in LB peaked around hour 5, analogous to the situation found for the wild type, in the glucose-supplemented LB, spa transcription seemed to be highest at the latest time point monitored (i.e., hour 8). Interestingly, significant amounts of spa transcripts were already detectable in MST23 during the early growth stages (hours 1 to 3), irrespective of whether glucose was present in the growth media or not, signaling that CcpA might act as a negative regulator for spa expression during these growth stages. Expression of hla was found to be less affected by glucose and/or CcpA compared with RNAIII and spa expression results. In COLn grown in unsupplemented LB, hla transcripts were detectable from hour 1 on and peaked around the transition from late logarithmic-growth phase to stationary phase. Supplementation of glucose shifted the peak expression of hla to the last growth point monitored and seemed to increase the expression level at this growth stage, probably due to the action of RNAIII, which was, as shown above, found to be highly expressed under these conditions. Essentially the same hla expression patterns as those identified in the wild type were found for MST23, although the overall amounts of hla transcripts seemed to be slightly reduced in the ccpA mutant.

Since hla and spa expression are known to be under multiple levels of control, including that by RNAIII, it was difficult to judge from the results described above whether the effects observed for hla and spa were the result of direct CcpA-mediated regulation in response to glucose or whether they might represent a secondary effect of RNAIII and other regulatory elements that were not part of this study. To better define the impact of glucose and, in particular, of CcpA on the expression of these two virulence factors, we performed a second series of Northern blot experiments, this time monitoring the expression of ccpA, hla, pckA, and spa in cells that were grown to midexponential-growth phase (A600 = 1). At this time point, glucose was added to one half of the culture, and cells were harvested in 10-min intervals from 0 to 30 min (Fig. 5). This procedure was likely to abolish the effect of agr on hla and spa expression, since RNAIII expression was barely detectable at this time point and under these conditions (data not shown). Moreover, by analyzing the gene expression immediately after the addition of glucose, we assumed that we would be more likely able to identify direct CcpA-dependent effects, since secondary effects were expected to occur with a certain delay. Addition of glucose to the exponentially growing cultures did not result in a temporary growth arrest of these cultures, indicating that the glucose addition did not trigger any inhibition of the primary metabolism (data not shown).


Figure 5
View larger version (66K):
[in this window]
[in a new window]
  		FIG. 5. Effect of glucose addition on the gene expression of COLn and its {Delta}ccpA mutant MST23. (A) Northern blot analyses of ccpA, hla, pckA, and spa. Cells were grown in HEPES-buffered LB to midexponential-growth phase (A600 = 1), cultures were split in half, and 10 mM glucose was added to one half (+Glc) while the other half was left unchanged (–Glc). Relevant transcript sizes and time points of sampling are indicated. Ethidium bromide-stained 16S rRNA patterns are shown as an indication of RNA loading. (B) Putative CREs identified upstream of ccpA, hla, pckA, and spa. Nucleotides fitting with the CRE consensus of B. subtilis (38) are highlighted in bold type. Inverted repeats are indicated by arrows. Ambiguity codes are as follows: W denotes A or T, R denotes G or A, and N denotes A, C, G, or T.

 
In this second series of Northern blot analyses, the level of expression of spa in unsupplemented LB appeared to be roughly constant, only slightly increasing with time, in agreement with the previous findings. Addition of glucose, however, resulted in a clear decrease in spa transcription that was already visible after 10 to 20 min in the wild type; this effect was not seen with MST23 (Fig. 5A). Moreover, the levels of spa transcripts in MST23 seemed clearly to be higher than those found in COLn, supporting the hypothesis that CcpA of S. aureus acts as a direct negative regulator of spa expression, either in the absence of glucose or, in a stronger way, in the presence of glucose. Further support for CcpA regulating spa expression is given by the fact that the spa coding region is preceded by a putative CRE consensus sequence (Fig. 5B). Interestingly, a potential CRE element that perfectly matched with the B. subtilis CRE consensus was further identified in the genomic region upstream of the open reading frame of hla, signaling that {alpha}-hemolysin production might be subjected to a CcpA-dependent CCR as well. In line with this assumption, we identified a slight decrease in hla transcription in COLn after the addition of glucose that was not detected either in the wild type grown in unsupplemented LB or in the ccpA mutant under both sets of conditions (Fig. 5A).

In agreement with the results of the first series of Northern blot analyses, we found pckA expression to decrease in the wild type in response to the presence of glucose and to be constant in the absence of this sugar. Equivalent to the situation found with spa, expression of pckA appeared to be clearly increased in MST23, and addition of glucose again failed to exert an effect on pckA expression in the mutant, confirming the findings of Scovill et al. (49) showing that the presence of glucose represses pckA transcription and supporting the hypothesis raised by these authors that this negative regulatory effect is exerted via CcpA. Interestingly, expression of ccpA itself seems to be positively affected by glucose, since transcription of ccpA appeared to be increased in response to the addition of glucose, although this increase was only of a transient nature and was only detectable at 10 min after the sugar was added (Fig. 5A). A potential CRE was identifiable in the promoter region of ccpA (Fig. 5B), sharing 15 out of 18 nucleotides with the CRE consensus of B. subtilis but lacking the palindromic nature of CREs that was detectable in the CRE candidates of hla, pckA, and spa and leaving the question currently open of whether the observed increase in transcription was mediated via CcpA by itself or not.

Effect of ccpA and glucose on capsular polysaccharide production. The majority of clinical isolates produce capsular polysaccharides of serotype 5 (CP5) or serotype 8 (CP8), which protect S. aureus against opsonophagocytic killing by polymorphonuclear leukocytes (28, 29, 34, 54, 59) and have been shown in a number of animal models of infection to enhance its virulence (40, 43, 54, 56, 61). Expression of CPs is influenced by various environmental signals in vitro and in vivo (reviewed in references 41 and 59), and transcription of the cap operon was shown to be affected by regulatory elements such as agr, mgr, sae, sarA, and the alternative {sigma}-factor {sigma}B (2, 10, 33, 34, 35, 42, 51, 57). Both biosynthetic pathways for CP5 and CP8 production utilize precursors of the cell wall, such as UDP-N-acetylglucosamine, suggesting that CP-producing proteins might compete with cell-wall-producing enzymes for the availability of UDP-N-acetylglucosamine, thereby affecting the carbon flux of S. aureus. We therefore analyzed whether glucose, and specifically ccpA, might affect CP synthesis, as has been shown recently for the low G+C-content gram-positive pathogen Clostridium perfringens (58).

Since the CP serotype 5 strain COLn was found to produce only a little CP under the conditions tested (C. Wolz, unpublished data), the CP5 prototypic strain Newman and its {Delta}ccpA derivative MST14 were used to investigate the effect of glucose and CcpA on CP formation (Fig. 6). Newman wild-type and MST14 cells were grown for 24 h in LB medium in the presence or absence of 10 mM glucose, and the CP5 production was determined by indirect immunofluorescence using monoclonal antibodies raised against CP5 (Fig. 6A). In the absence of glucose, most of the wild-type cells produced CP5. However, in the presence of glucose, CP5 was abolished, indicating that glucose repressed CP formation. Interestingly, CP5 production in the ccpA mutant MST14 was not affected and was present irrespective of the presence or absence of glucose in the growth medium. The immunofluorescence data were confirmed by real-time PCR (Fig. 6B). The expression of the cap operon was almost indistinguishable between the wild type and mutant in the absence of glucose, whereas in the presence of glucose, strain Newman produced significantly fewer cap transcripts (P < 0.05) than MST14, which expressed cap in roughly the same amounts as in the absence of glucose. Both findings strongly suggested that the presence of glucose repressed CP formation and that this effect was, at least in part, mediated via CcpA on the transcriptional level, adding a further regulator to the complex network of regulatory elements and environmental conditions that control cap operon expression. However, since no apparent CRE was identifiable within the genomic region encoding the cap operon, it is again likely that the CcpA effect on cap transcription was of an indirect nature and might be mediated by downstream regulators.


Figure 6
View larger version (24K):
[in this window]
[in a new window]
  		FIG. 6. Capsule production and cap expression of Newman and its {Delta}ccpA mutant MST14 in response to glucose. (A) CP5 expression determined by indirect immunofluorescence of strain Newman and its isogenic {Delta}ccpA mutant grown for 24 h at 37°C in HEPES-buffered LB (–Glc), or in HEPES-buffered LB supplemented with 10 mM glucose (+Glc). Bacteria were stained with 4',6'-diamidino-2-phenylindole (DAPI), and marked with CP5-specific monoclonal antibodies and stained with Cy3-conjugated anti-mouse antibodies (CY3). (B) Quantitative transcript analysis of capA by LightCycler RT-PCR of strain Newman and its isogenic {Delta}ccpA mutant grown for 8 h at 37°C in HEPES-buffered LB (–Glc) or in HEPES-buffered LB supplemented with 10 mM glucose (+Glc). Transcripts were quantified in reference to the transcription of gyrase (in copies per copy of gyrB). Values from two separate RNA isolations and two independent RT-PCRs each were used to calculate the mean expression (±standard errors of the mean). Glc, glucose; asterisk, P < 0.05 for Newman without Glc, MST14 without Glc, and MST14 with Glc.

 
Concluding remarks. Deletion of ccpA had a clear impact on the expression of RNAIII and on virulence factors of S. aureus, some of which have previously been shown to be affected by glucose. Interestingly, the deletion of ccpA produced an effect on gene expression not only in the presence but also in the absence of glucose, indicating that the function of CcpA might not be restricted to CCR. Our findings that CcpA of S. aureus influenced the transcription of at least five genes and operons, with most of them being involved in virulence of this pathogen, suggests that CcpA might represent an important global regulator of gene expression in S. aureus that, like that of its homologue in B. subtilis, may not be limited to regulating carbon uptake and metabolization. A preliminary computational screening of the S. aureus COL genome with the CRE consensus of B. subtilis (38) indeed indicated more than 110 CREs to be present in the promoter or N-terminal coding regions of genes and operons encoded by S. aureus, if allowing one mismatch to occur. Whole genome and proteomic analyses are currently ongoing to identify the CcpA regulon in S. aureus.


   ACKNOWLEDGMENTS
 
The study was supported by the Novartis Foundation, the Swiss National Science Foundation (grants 3100A0-100234 and 31-105390), the Deutsche Forschungsgemeinschaft (Wo 578/5-1), and the European Community (grant EU-LSH-CT2003-50335) (BBW 03.0098).

We thank Jean-Michel Fournier for kindly donating the monoclonal anticapsular antibody and the AO Research Institute for providing us with the necessary materials for the scanning electron microscopy experiments.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Medical Microbiology, University of Zürich, Gloriastr. 32, CH-8006 Zürich, Switzerland. Phone: 41 44 634 26 70. Fax: 41 44 634 49 06. E-mail: Bischoff{at}immv.unizh.ch. Back

{dagger} K.S. and M.S. contributed equally to the article. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussion
 References
 

  1. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl. 1987. Current protocols in molecular biology. John Wiley and Sons, Inc., New York, N.Y.
  2. Bischoff, M., P. Dunman, J. Kormanec, D. Macapagal, E. Murphy, W. Mounts, B. Berger-Bächi, and S. Projan. 2004. Microarray-based analysis of the Staphylococcus aureus {sigma}B regulon. J. Bacteriol. 186:4085-4099.[Abstract/Free Full Text]
  3. Blencke, H. M., G. Homuth, H. Ludwig, U. Mader, M. Hecker, and J. Stulke. 2003. Transcriptional profiling of gene expression in response to glucose in Bacillus subtilis: regulation of the central metabolic pathways. Metab. Eng. 5:133-149.[CrossRef][Medline]
  4. Brückner, R. 1997. Gene replacement in Staphylococcus carnosus and Staphylococcus xylosus. FEMS Microbiol. Lett. 151:1-8.[Medline]
  5. Chan, P. F., and S. J. Foster. 1998. Role of SarA in virulence determinant production and environmental signal transduction in Staphylococcus aureus. J. Bacteriol. 180:6232-6241.[Abstract/Free Full Text]
  6. Cheung, A. L., K. J. Eberhardt, and V. A. Fischetti. 1994. A method to isolate RNA from gram-positive bacteria and mycobacteria. Anal. Biochem. 222:511-514.[CrossRef][Medline]
  7. Cheung, A. L., K. Schmidt, B. Bateman, and A. C. Manna. 2001. SarS, a SarA homolog repressible by agr, is an activator of protein A synthesis in Staphylococcus aureus. Infect. Immun. 69:2448-2455.[Abstract/Free Full Text]
  8. Cheung, A. L., and A. C. Manna. 2005. Role of the distal sarA promoters in SarA expression in Staphylococcus aureus. Infect. Immun. 73:4391-4394.[Abstract/Free Full Text]
  9. CLSI. 2003. Methods for dilution antimicrobial susceptibility tests for bacteria that grow aerobically. Approved standard—sixth edition, CLSI document M7-A6. CLSI, Wayne, Pa.
  10. Dassy, B., T. Hogan, T. J. Foster, and J. M. Fournier. 1993. Involvement of the accessory gene regulator (agr) in expression of type-5 capsular polysaccharide by Staphylococcus aureus. J. Gen. Microbiol. 139:1301-1306.
  11. De Lencastre, H., S. W. Wu, M. G. Pinho, A. M Ludovice, S. Filipe, S. Gardete, R. Sobral, S. Gill, M. Chung, and A. Tomasz. 1999. Antibiotic resistance as a stress response: complete sequencing of a large number of chromosomal loci in Staphylococcus aureus strain COL that impact on the expression of resistance to methicillin. Microb. Drug Resist. 5:163-175.[Medline]
  12. Duncan, J. L., and G. J. Cho. 1972. Production of staphylococcal alpha toxin II. Glucose repression of toxin formation. Infect. Immun. 6:689-694.[Abstract/Free Full Text]
  13. Duthie, E. S., and L. L. Lorenz. 1952. Staphylococcal coagulase: mode of action and antigenicity. J. Gen. Microbiol. 6:95-107.[Medline]
  14. Egeter, O., and R. Brückner. 1996. Catabolite repression mediated by the catabolite control protein CcpA in Staphylococcus xylosus. Mol. Microbiol. 21:739-749.[CrossRef][Medline]
  15. Fournier, B., A. Klier, and G. Rapoport. 2001. The two-component system ArlS-ArlR is a regulator of virulence gene expression in Staphylococcus aureus. Mol. Microbiol. 41:247-261.[CrossRef][Medline]
  16. Garvis, S., J. M. Mei, J. Ruiz-Albert, and D. W. Holden. 2002. Staphylococcus aureus svrA: a gene required for virulence and expression of the agr locus. Microbiology 148:3235-3243.[Abstract/Free Full Text]
  17. Giachino, P., S. Engelmann, and M. Bischoff. 2001. {sigma}B activity depends on RsbU in Staphylococcus aureus. J. Bacteriol. 183:1843-1852.[Abstract/Free Full Text]
  18. Giraudo, A. T., A. L. Cheung, and R. Nagel. 1997. The sae locus of Staphylococcus aureus controls exoprotein synthesis at the transcriptional level. Arch. Microbiol. 168:53-58.[CrossRef][Medline]
  19. Goerke, C., S. Campana, M. G. Bayer, G. Döring, K. Botzenhart, and C. Wolz. 2000. Direct quantitative transcript analysis of the agr regulon of Staphylococcus aureus during human infection in comparison to the expression profile in vitro. Infect. Immun. 68:1304-1311.[Abstract/Free Full Text]
  20. Goerke, C., U. Flückiger, A. Steinhuber, V. Bisanzio, M. Ulrich, M. Bischoff, J. M. Patti, and C. Wolz. 2005. Role of Staphylococcus aureus global regulators sae and SigmaB in virulence gene expression during device-related infection. Infect. Immun. 73:3415-3421.[Abstract/Free Full Text]
  21. Harris, L. G., S. Tosatti, M. Wieland, M. Textor, and R. G. Richards. 2004. Staphylococcus aureus adhesion to titanium oxide surfaces coated with non-functionalized and peptide-functionalized poly(L-lysine)-grafted-poly(ethylene glycol) copolymers. Biomaterials 25:4135-4148.[CrossRef][Medline]
  22. Hoeger, P. H., W. Lenz, A. Boutonnier, and J. M. Fournier. 1992. Staphylococcal skin colonization in children with atopic dermatitis: prevalence, persistence, and transmission of toxigenic and nontoxigenic strains. J. Infect. Dis. 165:1064-1068.[Medline]
  23. Horsburgh, M. J., J. L. Aish, I. J. White, L. Shaw, J. K. Lithgow, and S. J. Foster. 2002. SigmaB modulates virulence determinant expression and stress resistance: characterization of a functional rsbU strain derived from Staphylococcus aureus 8325-4. J. Bacteriol. 184:5457-5467.[Abstract/Free Full Text]
  24. Iandolo, J. J., and W. M. Shafer. 1977. Regulation of staphylococcal enterotoxin B. Infect. Immun. 16:610-616.[Abstract/Free Full Text]
  25. Ingavale, S., W. van Wamel, T. T. Luong, C. Y. Lee, and A. L. Cheung. 2005. Rat/MgrA, a regulator of autolysis, is a regulator of virulence genes in Staphylococcus aureus. Infect. Immun. 73:1423-1431.[Abstract/Free Full Text]
  26. Jankovic, I., and R. Brückner. 2002. Carbon catabolite repression by the catabolite control protein CcpA in Staphylococcus xylosus. J. Mol. Microbiol. Biotechnol. 4:309-314.[Medline]
  27. Jarvis, A. W., R. C. Lawrence, and G. G. Pritchard. 1975. Glucose repression of enterotoxins A, B and C and other extracellular proteins in staphylococci in batch and continuous culture. J. Gen. Microbiol. 86:75-87.[Medline]
  28. Kampen, A. H., T. Tollersrud, and A. Lund. 2005. Staphylococcus aureus capsular polysaccharide types 5 and 8 reduce killing by bovine neutrophils in vitro. Infect. Immun. 73:1578-1583.[Abstract/Free Full Text]
  29. Karakawa, W. W., A. Sutton, R. Schneerson, A. Karpas, and W. F. Vann. 1988. Capsular antibodies induce type-specific phagocytosis of capsulated Staphylococcus aureus by human polymorphonuclear leukocytes. Infect. Immun. 56:1090-1095.[Abstract/Free Full Text]
  30. Katayama, Y., H. Z. Zhang, and H. F. Chambers. 2004. PBP 2a mutations producing very-high-level resistance to beta-lactams. Antimicrob. Agents Chemother. 48:453-459.[Abstract/Free Full Text]
  31. Kim, J. H., Y. K. Yang, and G. H. Chambliss. 2005. Evidence that Bacillus catabolite control protein CcpA interacts with RNA polymerase to inhibit transcription. Mol. Microbiol. 56:155-162.[CrossRef][Medline]
  32. Kreiswirth, B. N., S. Löfdahl, M. J. Betley, M. O'Reilly, P. M. Schlievert, M. S. Bergdoll, and R. P. Novick. 1983. The toxic shock syndrome exotoxin structural gene is not detectably transmitted by a prophage. Nature 305:709-712.[CrossRef][Medline]
  33. Luong, T., S. Sau, M. Gomez, J. C. Lee, and C. Y. Lee. 2002. Regulation of Staphylococcus aureus capsular polysaccharide expression by agr and sarA. Infect. Immun. 70:444-450.[Abstract/Free Full Text]
  34. Luong, T. T., and C. Y. Lee. 2002. Overproduction of type 8 capsular polysaccharide augments Staphylococcus aureus virulence. Infect. Immun. 70:3389-3395.[Abstract/Free Full Text]
  35. Luong, T. T., S. W. Newell, and C. Y. Lee. 2003. mgr, a novel global regulator in Staphylococcus aureus. J. Bacteriol. 185:3703-3710.[Abstract/Free Full Text]
  36. McCallum, N., M. Bischoff, H. Maki, A. Wada, and B. Berger-Bächi. 2004. TcaR, a putative MarR-like regulator of sarS expression. J. Bacteriol. 186:2966-2972.[Abstract/Free Full Text]
  37. McNamara, P. J., K. C. Milligan-Monroe, S. Khalili, and R. A. Proctor. 2000. Identification, cloning, and initial characterization of rot, a locus encoding a regulator of virulence factor expression in Staphylococcus aureus. J. Bacteriol. 182:3197-3203.[Abstract/Free Full Text]
  38. Miwa, Y., A. Nakata, A. Ogiwara, M. Yamamoto, and Y. Fujita. 2000. Evaluation and characterization of catabolite-responsive elements (cre) of Bacillus subtilis. Nucleic Acids Res. 28:1206-1210.[Abstract/Free Full Text]
  39. Moreno, M. S., B. L. Schneider, R. R. Maile, W. Weyler, and M. H. Saier, Jr. 2001. Catabolite repression mediated by the CcpA protein in Bacillus subtilis: novel modes of regulation revealed by whole-genome analyses. Mol. Microbiol. 39:1366-1381.[CrossRef][Medline]
  40. Nilsson, I.-M., J. C. Lee, T. Bremell, C. Ryden, and A. Tarkowski. 1997. The role of staphylococcal polysaccharide microcapsule expression in septicemia and septic arthritis. Infect. Immun. 65:4216-4221.[Abstract]
  41. O'Riordan, K., and J. C. Lee. 2004. Staphylococcus aureus capsular polysaccharides. Clin. Microbiol. Rev. 17:218-234.[Abstract/Free Full Text]
  42. Pohlmann-Dietze, P., M. Ulrich, K. B. Kiser, G. Doring, J. C. Lee, J. M. Fournier, K. Botzenhart, and C. Wolz. 2000. Adherence of Staphylococcus aureus to endothelial cells: influence of the capsular polysaccharide, the global regulator agr, and the bacterial growth phase. Infect. Immun. 68:4865-4871.[Abstract/Free Full Text]
  43. Portoles, M., K. B. Kiser, N. Bhasin, K. H. N. Chan, and J. C. Lee. 2001. Staphylococcus aureus Cap5O has UDP-ManNAc dehydrogenase activity and is essential for capsule expression. Infect. Immun. 69:917-923.[Abstract/Free Full Text]
  44. Regassa, L. B., J. L. Couch, and M. J. Betley. 1991. Steady-state staphylococcal enterotoxin type C mRNA is affected by a product of the accessory gene regulator (agr) and by glucose. Infect. Immun. 59:955-962.[Abstract/Free Full Text]
  45. Regassa, L. B., R. P. Novick, and M. J. Betley. 1992. Glucose and nonmaintained pH decrease expression of the accessory gene regulator (agr) in Staphylococcus aureus. Infect. Immun. 60:3381-3388.[Abstract/Free Full Text]
  46. Rossi, J., M. Bischoff, A. Wada, and B. Berger-Bächi. 2003. MsrR, a putative cell envelope-associated element involved in Staphylococcus aureus sarA attenuation. Antimicrob. Agents Chemother. 47:2558-2564.[Abstract/Free Full Text]
  47. Said-Salim, B., P. M. Dunman, F. M. McAleese, D. Macapagal, E. Murphy, P. J. McNamara, S. Arvidson, T. J. Foster, S. J. Projan, and B. N. Kreiswirth. 2003. Global regulation of Staphylococcus aureus genes by Rot. J. Bacteriol. 185:610-619.[Abstract/Free Full Text]
  48. Schmidt, K. A., A. C. Manna, S. Gill, and A. L. Cheung. 2001. SarT, a repressor of alpha-hemolysin in Staphylococcus aureus. Infect. Immun. 69:4749-4758.[Abstract/Free Full Text]
  49. Scovill, W. H., H. J. Schreier, and K. W. Bayles. 1996. Identification and characterization of the pckA gene from Staphylococcus aureus. J. Bacteriol. 178:3362-3364.[Abstract/Free Full Text]
  50. Senn, M. M., M. Bischoff, C. von Eiff, and B. Berger-Bächi. 2005. {sigma}B activity in a Staphylococcus aureus strain Newman hemB mutant. J. Bacteriol. 187:7397-7406.[Abstract/Free Full Text]
  51. Steinhuber, A., C. Goerke, M. G. Bayer, G. Doring, and C. Wolz. 2003. Molecular architecture of the regulatory locus sae of Staphylococcus aureus and its impact on expression of virulence factors. J. Bacteriol. 185:6278-6286.[Abstract/Free Full Text]
  52. Stülke, J., and W. Hillen. 2000. Regulation of carbon catabolism in Bacillus species. Annu. Rev. Microbiol. 54:849-880.[CrossRef][Medline]
  53. Tegmark, K., A. Karlsson, and S. Arvidson. 2000. Identification and characterization of SarH1, a new global regulator of virulence gene expression in Staphylococcus aureus. Mol. Microbiol. 37:398-409.[CrossRef][Medline]
  54. Thakker, M., J.-S. Park, V. Carey, and J. C. Lee. 1998. Staphylococcus aureus serotype 5 capsular polysaccharide is antiphagocytic and enhances bacterial virulence in a murine bacteremia model. Infect. Immun. 66:5183-5189.[Abstract/Free Full Text]
  55. Titgemeyer, F., and W. Hillen. 2002. Global control of sugar metabolism: a gram-positive solution. Antonie Leeuwenhoek 82:59-71.
  56. Tzianabos, A. O., J. Y. Wang, and J. C. Lee. 2001. Structural rationale for the modulation of abscess formation by Staphylococcus aureus capsular polysaccharides. Proc. Natl. Acad. Sci. USA 98:9365-9370.[Abstract/Free Full Text]
  57. van Wamel, W., Y. Q. Xiong, A. S. Bayer, M. R. Yeaman, C. C. Nast, and A. L. Cheung. 2002. Regulation of Staphylococcus aureus type 5 capsular polysaccharides by agr and sarA in vitro and in an experimental endocarditis model. Microb. Pathog. 33:73-79.[CrossRef][Medline]
  58. Varga, J., V. L. Stirewalt, and S. B. Melville. 2004. The CcpA protein is necessary for efficient sporulation and enterotoxin gene (cpe) regulation in Clostridium perfringens. J. Bacteriol. 186:5221-5229.[Abstract/Free Full Text]
  59. Voyich, J. M., K. R. Braughton, D. E. Sturdevant, A. R. Whitney, et al. 2005. Insights into mechanisms used by Staphylococcus aureus to avoid destruction by human neutrophils. J. Immunol. 175:3907-3919.[Abstract/Free Full Text]
  60. Warner, J. B., and J. S. Lolkema. 2003. CcpA-dependent carbon catabolite repression in bacteria. Microbiol. Mol. Biol. Rev. 67:475-490.[Abstract/Free Full Text]
  61. Watts, A., D. Ke, Q. Wang, A. Pillay, A. Nicholson-Weller, and J. C. Lee. 2005. Staphylococcus aureus strains that express serotype 5 or serotype 8 capsular polysaccharides differ in virulence. Infect. Immun. 73:3502-3511.[Abstract/Free Full Text]
  62. Weinrick, B., P. M. Dunman, F. McAleese, E. Murphy, S. J. Projan, Y. Fang, and R. P. Novick. 2004. Effect of mild acid on gene expression in Staphylococcus aureus. J. Bacteriol. 186:8407-8423.[Abstract/Free Full Text]
  63. Xiong, Y. Q., W. van Wamel, C. C. Nast, M. R. Yeaman, A. L. Cheung, and A. S. Bayer. 2002. Activation and transcriptional interaction between agr RNAII and RNAIII in Staphylococcus aureus in vitro and in an experimental endocarditis model. J. Infect. Dis. 186:668-677.[CrossRef][Medline]

Antimicrobial Agents and Chemotherapy, April 2006, p. 1183-1194, Vol. 50, No. 4
0066-4804/06/$08.00+0     doi:10.1128/AAC.50.4.1183-1194.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:


This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Seidl, K.
Right arrow Articles by Bischoff, M.