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Journal of Bacteriology, August 2005, p. 5301-5309, Vol. 187, No. 15
0021-9193/05/$08.00+0     doi:10.1128/JB.187.15.5301-5309.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Rot Repression of Enterotoxin B Expression in Staphylococcus aureus

Ching Wen Tseng1 and George C. Stewart2*

Department of Diagnostic Medicine/Pathobiology, Kansas State University, Manhattan, Kansas,1 Department of Veterinary Pathobiology, Life Sciences Center, University of Missouri, Columbia, Missouri2

Received 20 January 2005/ Accepted 11 May 2005


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The accessory gene regulator (Agr) system is a quorum-sensing system of Staphylococcus aureus responsible for upregulation of certain exoprotein genes and downregulation of certain cell-wall associated proteins during the post-exponential phase of growth. The enterotoxin B (seb) determinant is upregulated by the Agr system. Agr-regulated cis elements within the seb promoter region were examined by deletion analyses of the seb promoter by a hybrid promoter approach utilizing the staphylococcal lac operon promoter. To identify the regulatory pathway for enterotoxin B expression, the seb promoter fused to the chloramphenicol acetyltransferase reporter gene was introduced into mutants of S. aureus lacking agr or different members of the Sar family of transcriptional regulators. Agr control of seb promoter activity was found to be dependent upon the presence of a functional Rot protein, and Rot was shown to be able to bind to the seb promoter. Therefore, the Agr-mediated post-exponential-phase increase in seb transcription results from the Agr system's inactivation of Rot repressor activity.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Staphylococcus aureus causes a wide range of diseases, including food poisoning, cutaneous infections, endocarditis, pneumonia, septic arthritis, and osteomyelitis (15, 18). A variety of virulence factors are produced by this organism, including multiple exoproteins and cell wall-associated components (5, 22, 23). Coordinated regulation of expression of the many virulence genes is a critical feature of the pathogenicity of S. aureus, and the regulatory networks might provide sites of possible therapeutic intervention for the treatment of staphylococcal infections. To date, several global regulators have been identified that regulate the production of virulence-associated exoproteins and cell wall components. These global regulators include the accessory gene regulator (Agr), staphylococcal accessory regulator (SarA), repressor of toxins (Rot), and the SigB alternative sigma factor (2, 6, 11, 26, 28). Among these regulatory systems, Agr, a quorum-sensing system, has been the best characterized. The agr operon encodes a two-component regulatory system consisting of two distinct transcripts, RNAII and RNAIII, the former of which encodes the structural genes for the quorum-sensing system, AgrBDCA (14). The Agr system responds to an agrD-encoded 7- to 9-mer polypeptide signal, the autoinducing peptide (AIP), which is processed and possibly exported from the cell by AgrB (36, 37). AgrA is the response regulator and, when activated, upregulates the activity of the agr P2 and P3 promoters to increase the production of RNAII and RNAIII (14, 35). AgrC is the transmembrane histidine kinase responsible for sensing the level of AIP in the environment (16). Accumulation of a threshold concentration of AIP in the environment leads to an activation and autophosphorylation of AgrC, which subsequently transfers the phosphate to AgrA (16). RNAIII encodes {delta}-hemolysin, but the RNAIII itself serves as the regulatory signal of the Agr system (13, 24). As the cell density increases in a growing culture, the intracellular level of RNAIII increases due to the activity of the Agr system. Increased RNAIII levels lead to an increased transcription of many exotoxin genes and a reduced transcription of certain cell wall protein determinants (14, 21). Inactivation of the Agr system leads to a nonhemolytic and nonproteolytic phenotype, which is due to the failure of exoprotein induction (14, 21).

The sarA locus encodes a transcriptional factor which up-regulates expression of RNAIII, thus influencing the expression of Agr-regulated genes (2). SarA has also been shown to regulate gene expression in an Agr-independent fashion (3).

The repressor of toxins (rot) locus was first identified through a transposon mutagenesis study (20). Inactivation of rot was able to partially restore {alpha}-hemolysin and protease-positive phenotypes in an agr-null mutant (20). Rot is a member of the Sar family of transcriptional factors of S. aureus. An examination of Rot regulatory effects using a microarray analysis has shown that Rot acts as a global regulator that affects several virulence factors, including geh, ssp protease, alpha-toxin, protein A, and clumping factor (28). Rot acts as both a negative and positive regulator of gene expression (28). Rot activity is regulated by the Agr system. Results from Northern blot analysis and reporter gene assays have shown that transcription of Rot is not affected by the Agr system, during the post-exponential phase of growth. However, the activity of Rot is negatively regulated by the Agr system. With the induction of the Agr system during the post-exponential growth phase, the activity of Rot as a repressor of exotoxin expression is inactivated. Thus, Rot affects transcription of its target genes during the exponential phase of growth (20). The exact mechanism of Rot regulation by the Agr system is not known.

Staphylococcal enterotoxins are the virulence factors responsible for staphylococcal gastroenteritis (5, 39). The symptoms of staphylococcal gastroenteritis include emesis and abdominal cramping. A number of staphylococcal enterotoxins have been identified, and they are distinguished by serological or amino acid sequence differences. These enterotoxins are designated as staphylococcal enterotoxins A (SEA) through R (SER), excluding SEF, which was previously the designation for toxic shock syndrome toxin (5, 39). A number of the enterotoxins have been shown to be superantigens, capable of polyclonal T-cell stimulation that results in release of inflammatory cytokines and symptoms of shock (10, 25). The staphylococcal enterotoxin B (seb) determinant is carried on a pathogenicity island designated SaPI3 in S. aureus (23). SEB appears in the culture medium in the greatest quantity during the transition from exponential growth to stationary-phase growth, a characteristic of its regulation by the Agr system (14, 19). Loss of the Agr signal transduction system results in substantial reductions in seb transcription (8). In addition, strain-specific differences in expression patterns for SEB have been reported. For, example, strain S6 secretes 30-fold more SEB than does strain COL (4). However, analysis of the DNA sequences of the seb promoter elements from these strains revealed identical sequences (4). The different levels of SEB production have been attributed to different levels of RNAIII production by each strain (4).

The seb promoter element was previously characterized by Khan and coworkers. They reported that the sequences upstream of –93 are not important for the regulated expression of seb (17). Deletion of the upstream promoter element to –59 resulted in a dramatic reduction in the seb mRNA level. They concluded that sequences required for the expression and regulation of seb reside between –93 and –59 (17). Cheung and coworkers used green fluorescent protein as a reporter gene for analyzing the promoter activity of the seb gene and reported a reduced level of seb expression in a SigB+ strain (30). In addition, the level of seb expression was significantly lower in a SigB+ Agr– strain than those of an Agr– strain. Therefore, they concluded that the SigB regulation of seb expression is independent of the Agr regulation.

In this study, we created a series of deletions of the seb promoter and defined the effects of these truncated promoter activities. In addition, a hybrid promoter approach, using the core promoter sequences from a staphylococcal lactose operon (lac) fused to the sequences upstream of the seb core promoter element, was utilized to further define the regulatory role of the upstream sequences.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Media and culture conditions. The bacterial strains used are listed in Table 1. All S. aureus strains are 8325-4 derivatives. The wild-type 8325-4 strains do not produce the SigB transcriptional factor owing to a small deletion in the regulatory locus affecting this alternative sigma factor (9). The SigB-positive cultures have the genetically repaired rbsU allele from strain GP269 introduced by transduction. Escherichia coli cultures were grown in L-broth (1% NaCl, 1% tryptone, and 0.5% yeast extract) with shaking at 200 rpm or on L-agar (L-broth with 1.5% agar) at 37°C. S. aureus cultures were grow in tryptic soy broth (TSB; Difco) with shaking at 200 rpm, or tryptic soy agar (Difco) at 37°C overnight. The following antibiotic concentrations were utilized: ampicillin, 100 µg/ml; erythromycin, 400 µg/ml for E. coli and 10 µg/ml for S. aureus; kanamycin, 25 µg/ml; and tetracycline, 20 µg/ml for E. coli and 10 µg/ml for S. aureus. Skim milk plates were prepared according to the Difco formulation.


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  		TABLE 1. Bacterial strains used in this study

 
DNA manipulations and plasmids. Promoter elements and the Rot open reading frame (ORF) were cloned by PCR. Site-directed mutagenesis was carried out using oligonucleotide primers containing the desired mutations as described previously (34). PCR primer pairs are listed in Table 2. The plasmids used are listed in Table 3. PCR-amplified promoter elements were inserted into the reporter shuttle plasmid pMH109 (12) following SacI and XbaI digestion. Subsequently, the inserted DNA fragments were subjected to DNA sequencing to confirm the identity of the inserted sequence. The Rot expression plasmid pDT34 (34) was used to express Rot in an agr- and rot-independent fashion. Plasmids were electroporated into RN4220 using the method of Schenk and Laddaga (29). To verify the inserted elements, plasmid DNA was isolated from RN4220 and transformed into E. coli strain DH5{alpha}. Plasmid DNA was isolated using Wizard Plus kits (Promega) and subjected to DNA sequence analysis. The sequence-verified plasmids were then transduced from RN4220 into strains with different genetic backgrounds using the method of Rubin and Rosenblum (27).


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  		TABLE 2. Primers used for promoter cloning

 

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  		TABLE 3. Plasmids used in this study

 
CAT assay. Chloramphenicol acetyltransferase (CAT) assays were performed by the spectrophotometric method of Shaw, adapted to a microtiter plate format (31, 34). Bacterial cultures were grown in TSB with appropriate antibiotic selection overnight at 37°C. Ten milliliters of prewarmed TSB (37°C) was inoculated with an overnight culture to an A540 of 0.2, unless otherwise indicated. The cultures were then incubated at 37°C with shaking, until the A540 was 2.5, unless otherwise indicated. The growth kinetics of the different mutant strains did not differ significantly from the wild-type S. aureus strain. Five milliliters of each bacterial culture was collected, the cells were harvested by centrifugation (5,000 x g, 10 min), and the cell pellets were washed with TE buffer (50 mM Tris-HCl, 10 mM EDTA, pH 8.0). The cell pellets were then resuspended in 1 ml TE buffer, and the cells were then lysed with 0.1-mm glass beads, twice for 1 min at 4°C using an eight-sample bead beater (Biospec Products). Between each cycle, the suspensions were cooled on ice for 1 min. The lysed bacterial samples were centrifuged (2,500 x g, 10 min) at 4°C, and the supernatant was saved and stored at –70°C. Two to 25 µl of cell lysate was added to 37.5 µl of 0.4% of 5, 5' dithiobis-2-nitrobenzoic acid (DTNB; Sigma) in 100 mM Tris-HCl (pH 8.0), and 7.5 µl of 5 mM acetyl coenzyme A (Amersham Pharmacia). Distilled water was added to give a final volume of 250 µl. The reaction mixtures were then incubated at 37°C for 10 min prior to the addition of 5 pmol of chloramphenicol (in 10 ml of 70% ethanol). The changes of absorbance at 412 nm were measured using a microplate reader (Molecular Devices). Results were normalized with respect to the total protein concentration of the cell extracts as determined using a Bio-Rad protein assay kit. CAT values were calculated as the change in absorbance per minute, divided by 13.6 (the molar extinction value for DTNB), and by the amount of protein added. CAT values were expressed as nanomoles of chloramphenicol acetylated per minute per mg protein.

Expression of recombinant Rot protein. The overexpression of recombinant Rot was accomplished using Qiaexpress system (QIAGEN). A PCR fragment containing the rot ORF (primers 5p rot exp and 3p rot exp [Table 2]) was inserted into pQE30 at the BamHI and PstI sites, producing pDT112. Expression of the His-tagged Rot protein in E. coli was induced by the addition of IPTG (isopropyl-ß-D-thiogalactopyranoside), and the protein was purified using His-spin kits (Zymo Corp.). The protein was purified to apparent homogeneity as evidenced by Coomassie blue staining of sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels.

EMSAs. Electrophoretic mobility shift assays (EMSAs) were performed as follows. DNA fragments were labeled with [{gamma}-32P]ATP using T4 polynucleotide kinase. Cell lysates were prepared from post-exponential-phase cultures (A540 = 2.5). The cells were washed and resuspended in TE buffer (50 mM Tris-HCl, pH 8, 10 mM EDTA). Cell suspensions were then standardized to contain 0.5 mg/ml cells (dry weight) in lysis buffer (50 mM Tris-HCl, pH 8, 10 mM EDTA, and 10% glycerol). The cells were lysed by beating twice for 1 min using 0.1-mm-diameter glass beads in a bead beater. Binding of lysate protein to radiolabeled DNA was accomplished using a previously described protocol in a buffer system containing 10 mM Tris-HCl [pH 7.5], 1 mM EDTA, 50 mM NaCl, and 5% glycerol (17). Recombinant His6-Rot was purified from E. coli strain AbleK harboring pREP4 (QIAGEN). Protein concentrations were determined using the Bradford method (Bio-Rad Corp.) using bovine serum albumin as a standard. Different amounts of purified Rot protein or cell lysate protein were incubated with the labeled DNA fragments for 30 min at room temperature and then incubated on ice for 10 min. Samples were then electrophoresed in 6% polyacrylamide gels under native conditions. The gels were dried and subjected to radioautography.

Statistical analyses. Each result is represented as the mean ± standard deviation from three independently collected samples. At least two sets of independently collected samples were tested. Results were subjected to a pair-sampled two-tailed Student t test. Results with P values of less than 0.05 were considered to be significant.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Regulation of seb transcription by Rot. Rot is a negative regulator of sed expression (34). To determine if Rot also affects seb expression, the seb promoter-containing fragment (–122 to +7; Fig. 1) was fused to a promoterless CAT gene in pMH109 (pGS762) and introduced into Agr+, Agr–, Agr+ Rot–, and Agr– Rot– strains. Post-exponential-growth-phase expression of the CAT reporter was then determined (Fig. 2). The seb promoter activity was low during the post-exponential phase in the Agr– host, indicative of the known Agr regulation of this promoter. However, in Rot– strains, post-exponential-phase activity of the seb promoter was elevated in both Agr-negative and Agr-positive hosts. Furthermore, loss of Rot essentially eliminated the Agr effect on seb promoter expression. Thus, Rot may act as a repressor of seb transcription during the exponential phase of growth and is inactivated by the Agr system, albeit incompletely, during the post-exponential phase of growth. To confirm that Rot acts as the repressor on the seb transcription, we transduced pGS762 into the wild-type, Agr–, Rot–, and Agr– Rot– strains containing the rot expression plasmid (pDT34) or its negative control plasmid lacking the rot encoding sequence (pDT33) (34). The strains were tested for protease and urease activities to confirm their Rot– and complemented phenotypes. The seb promoter activities in the different strains are shown in Fig. 2. The presence of the rot expression plasmid reduced the seb promoter activity approximately threefold in the wild-type and the Agr– Rot– strains. To determine if growth-phase-dependent expression of the seb promoter is Rot mediated, cultures from the exponential and post-exponential phases were collected and the seb promoter activities obtained are shown in Fig. 3. Loss of Rot activity resulted in elevated exponential-phase expression in both Agr-positive and Agr-negative host strains, and the post-exponential-phase induction of seb promoter activity was found to be dependent upon the presence of a functional Rot protein. Loss of Rot activity resulted in an approximately sixfold enhancement of the seb promoter activity in the exponential phase (P < 0.05, exponential growth phase versus the activity in the Rot+ background) and an approximately 30% increase in activity in the post-exponential growth phase. Therefore, Rot appears to be a transcriptional regulator of seb transcription and the repression of the seb promoter is alleviated, albeit incompletely, by the activated Agr system.



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  		FIG. 1. seb promoter element from –122 to +7. The –35 and –10 element sequences are in boldface. Four sets of the repeat elements are shown with the arrows: A 10-bp inverted repeat element ({zjb01505494300g1}), a 5-bp repeat element as a direct repeat ({zjb01505494300g2}) and in the inverted orientation ({zjb01505494300g3}), a 6-bp repeat element ({zjb01505494300g4} and {zjb01505494300g5}), and a 7-bp directly repeated sequence ({zjb01505494300g6}).

 


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  		FIG. 2. Rot regulation of seb promoter activity. Shown is the seb promoter activity in Rot– and/or Agr– genetic backgrounds with Rot expression plasmid (pDT34) and its negative control (pDT33). +, wild-type; –, knockout. The promoter activity is expressed as nanomoles of chloramphenicol acetylated per minute per mg protein (n ≥ 3, similar from at least two independent experiments; average ± standard deviation). P < 0.05 for Agr– or Agr– Rot– compared to Agr+, Agr– Rot–(pDT33) compared to Agr–, and Agr– Rot–(pDT34) compared to Agr– Rot–.

 


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  		FIG. 3. seb promoter activity from different growth phases. Shown is the seb promoter activity from wild-type and Agr– Rot– hosts in the exponential and post-exponential phases. Prewarmed medium was inoculated to an A540 of 0.1, and samples were independently collected at A540s of 0.4 and 2.5 and tested for promoter activity. Promoter activity is expressed as nanomoles of chloramphenicol acetylated per minute per mg of protein (n ≥ 3 from at least two independent experiments; average ± standard deviation).

 
Regulation of seb promoter activity by Agr and the Sar family of transcription regulators. Western blot analysis of SEB from culture supernatants indicated that SarA is an upregulator of seb expression (1). Mutants lacking SigB displayed an increased level of seb transcription (30). Both of these transcriptional regulators may affect seb expression indirectly through their effects on agr expression. To evaluate the contribution of the Sar family proteins to the seb promoter activity, pGS762 was transduced into the strains with different genetic backgrounds: the post-exponential-phase promoter activities obtained are shown in Fig. 4. For SigB+ strains, the sigB locus from strain GP269 was introduced by transduction. In Agr+ strains, restoration of SigB production reduces the post-exponential-phase expression of the seb promoter (compare columns 1 and 2 in Fig. 4A). In the Agr– hosts, the activity of the seb promoter is unaffected by the presence of an active {sigma}B (compare the columns 3 and 4 of Fig. 4A). Therefore, the effect of SigB on seb expression appears to be an indirect consequence of the SigB reduction of Agr expression.



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  		FIG. 4. seb promoter activity from different genetic background strains. +, wild type; –, knockout. Promoter activity is expressed as nanomoles of chloramphenicol acetylated per minute per mg of protein (n ≥ 3 from at least two independent experiments; average ± standard deviation). Samples were from post-exponential-phase cultures (A540 = 2.5). All strains are SigB negative, unless indicated otherwise. The results are presented in panels A to D to facilitate result comparisons in the text.

 
SarA affects expression of the seb promoter, and the effect is dependent on the Agr status of the strain. In Agr+ hosts, loss of SarA results in a reduction in seb promoter activity (compare columns 1 and 2 of Fig. 4B), results consistent with the reduction in Agr expression in SarA-minus hosts. However, in the Agr– hosts, a SarA effect on promoter activity is still observed, indicating that this regulator acts in an Agr-independent fashion as well. The activity of the seb promoter is reduced in the presence of an active SarA protein (compare columns 4 and 5 of Fig. 4B). SarA, or a factor whose expression is dependent on SarA, negatively affects the activity of the seb promoter. The effects of SarA on the seb expression appear to be both an indirect consequence of the SarA enhancement of the Agr expression and an Agr-independent inhibition of the seb transcription. With hosts that are SigB positive and SarA negative, an intermediate level of seb promoter activity was observed and this level of expression was independent of the Agr status of the strain (compare columns 7 and 8 of Fig. 4B).

Loss of the Rot protein was shown to elevate seb promoter activity and to make the expression largely independent of the Agr system. Interestingly, the elevation in seb promoter activity observed in the Rot– hosts is dependent upon the expression of SarA. In SarA– and Rot– mutant hosts, the seb promoter activity is approximately 5- to 10-fold lower than the values obtained with the isogenic SarA+ hosts (compare Fig. 4C, columns 2 and 5 for the Agr+ and Rot– hosts and columns 4 and 6 for the Agr– and Rot– mutant hosts). These results indicate that loss of Rot repression does not result in the elevated seb promoter activity unless SarA, or a SarA-induced transcriptional factor, is present. In addition, seb promoter activity in the SarA– and Rot– double mutant was still subject to a slight (twofold) Agr control (compare columns 5 and 6 of Fig. 4C). This result suggests that the Agr control of seb transcription does involve an additional transcription factor, but that this activity is masked in the Rot+ strains.

SarS does not have an appreciable effect on seb promoter activity (compare Fig. 4D, columns 1 and 2 (Agr+ hosts) and columns 3 and 4 (Agr– hosts). However, in some of the mutant genotype combinations examined, modest SarS effects were observed. These included a twofold elevation in seb promoter activity in the SarA–/SarS–/Agr+ host relative to the isogenic SarS+ host (compare Fig. 4B columns 2 and 3), but no comparable effect was observed in the Agr– host (compare columns 5 and 6 in Fig. 4B). Loss of SarS also results in a twofold reduction in promoter activity associated with a loss of Rot repression (compare column 2 [Fig. 4C] with column 5 [Fig. 4D] and column 4 [Fig. 4C] with column 6 [Fig. 4D]). The modest role played by SarS is masked in the Rot+ strains.

cis-Regulatory elements in the seb promoter region. Putative repeat elements are found in the seb promoter element between –108 and –1 (Fig. 1). These may simply be the consequence of the A+T-rich sequences of S. aureus, especially in promoter regions, or they may represent regulatory cis elements within the seb promoter. To investigate the importance of these repeat elements for the expression and regulation of seb, we constructed truncated promoter elements fused to the CAT reporter gene. Truncated promoter constructs were evaluated for their post-exponential-growth-phase promoter activities in Agr+, Agr–, Agr+ Rot–, and Agr– Rot– hosts. The promoter containing the upstream sequence to –122 (pGS762) displayed a 13-fold Agr upregulation of promoter activity (Fig. 5B). In the Rot– hosts, the Agr stimulatory effect on this promoter was abolished. Instead, the promoter activities from both Agr+ Rot– and Agr– Rot– hosts were increased ~1.7-fold compared to the Agr+ Rot+ host (P < 0.05). This construct appears to possess all of its regulatory sequences. When the upstream sequences were truncated to –91 (pDT85), the overall promoter activity increased and Agr regulation and Rot repression were intact. Truncation to –79 (pDT80) resulted in decreased promoter activity to a level comparable to that of the –122 construct in the Agr+ background, but a higher level of expression in the Agr-negative host was observed. The increased level of expression in the Agr-negative host resulted in only a twofold Agr stimulation effect. Rot repression was still evident with this promoter construct. Truncation of the promoter to –64 resulted in a transcriptionally active promoter, defining the upstream boundary for an active promoter to between –59 (17) and –64. Truncation of the promoter to the core promoter sequences (pDT83), containing only the –35 box and Pribnow box elements, resulted in an inactive promoter in S. aureus (0 ± 0.3; Fig. 5B). This promoter-containing plasmid was also evaluated in E. coli, and the promoter activity was detectable in this host (47.3 ± 10.2). In addition, the –49 truncated promoter was not detectably active, but activity was present in a –63 construct (data not shown). Khan and coworkers had previously reported that seb promoter activity was abolished when the upstream sequence was truncated to –59 (17). Thus, there appear to be at least two distinct elements within the seb promoter. The first is a positive element necessary for binding an activator of the promoter. This element has an upstream boundary between –64 and –59. The second element is required for Agr regulation, presumably as the binding site for Rot binding.



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  		FIG. 5. (A) Promoters are depicted as rectangles representing the –35 and –10 promoter elements, and the lines indicate the extent of upstream sequences present in the constructs, with the end points indicated at the top of the figure. Open boxes correspond to the seb promoter, whereas hatched boxes correspond to the lac operon promoter in the hybrid constructs. The series numbers correspond to the numbers along the x axis in panel B. (B) seb (groups 1 to 5) and seb-lac hybrid (groups 6 to 11) promoter elements and promoter activity from Agr+ (white bars), Agr– (black bars), Agr+ Rot– (hatched bars), and Agr– Rot– (stippled bars) hosts. The promoter element upstream boundaries were –122 (groups 1 and 6), –91 (groups 2 and 7), –79 (groups 3 and 8), –64 (groups 4 and 9), –36 (group 5), –49 (group 10), and the lac core promoter (group 11). The promoter activity is expressed as nanomoles of chloramphenicol acetylated per minute per mg of protein (n ≥ 3 from at least two independent experiments; average ± standard deviation).

 
The inactivity of the shorter promoter fragments makes it difficult to precisely define the boundaries of the Agr regulatory site. Therefore, a hybrid promoter approach was undertaken to further define the cis elements in the seb promoter. The non-Agr- and non-Rot-regulated staphylococcal lactose operon core promoter was fused to cat to permit the construction of hybrid promoters. Sequences upstream of the –35 element from the seb promoter element were positioned upstream of the –35 element of the lac core promoter (Fig. 5A). The hybrid promoter elements included the upstream sequence to –122 containing all the putative repeats (pDT2), and truncations of the upstream repeat containing sequences to –91 (pDT85), –82 (pDT18), –64 (pDT19), and –49 (pDT20). The post-exponential-phase promoter activities of these truncated hybrid promoter elements are presented in Fig. 5B.

The lac core promoter was indeed active and not subject to regulation by Agr or Rot (Fig. 5B). When the hybrid promoter element included seb upstream sequences to –122, a promoter that based on the seb promoter results should retain full Agr regulation, only a 1.6-fold Agr stimulation of transcription was observed. In addition, only a 1.8-fold Rot repressive effect was observed. This may be a result of the regulatory sequences being fused to an active promoter, whereas Agr control may only work well when the promoter requires activation (see the Discussion section). The larger truncations resulted in hybrid promoters that did not demonstrate Agr regulation nor evidence of Rot repression, suggesting that the 10-bp inverted repeat (Fig. 1) may be important in Rot binding and thus Agr regulation.

Interestingly, the addition of upstream sequences to –49 (12 bp of seb sequence fused to the –35 sequence of the lac promoter) resulted in a dramatic reduction (approximately 10-fold) of promoter activity. To investigate if the reduction in promoter activity was associated with the binding of a transcriptional factor or is a consequence of a sequence inherent property, pZS2785 (lac core), pDT20 (–49 hybrid), pDT19 (–64 hybrid), and pDT18 (–79 hybrid) were transformed into E. coli DH5{alpha}, and the promoter activities from these strains were tested. The presumption was that this heterologous host would lack the putative transcription factors that would bind to the 11-bp target sequence. The promoter activities were 10,748.1 ± 621.7, 177.3 ± 13.7, 4,081.3 ± 383.6, and 1,083.7 ± 39.6, respectively. The addition of the 11-bp sequence reduced the promoter strength in E. coli. Therefore, the A+T-rich 11-bp sequence upstream of the –35 box appears to be a kill switch that inactivates the promoter rather than a binding site for a transcriptional factor.

Electrophoretic mobility shift assay of the seb promoter containing DNA fragments. Expression of the Rot protein is not well tolerated by E. coli, frequently resulting in clones that contain deletions in the plasmid insert. To minimize the toxic effect of Rot expression on the host cells, pDT112 was transformed into E. coli strain AbleK to reduce the copy number of pDT112, the recipient cells contained pREP4 (QIAGEN) to minimize the background expression of Rot and the culture was grown in the presence of 2 mM glucose to maximize the repression of the lac promoter. Over 50% of colonies of the E. coli AbleK strain bearing both pDT112 and pREP4 were translucent and smaller in size when grown overnight on selective L-agar plates. Under microscopic examination, cell lysis was evident in the translucent colonies. The opaque colonies consisted of cells bearing plasmid deletions or mutations. Subculture of the translucent colonies produced mixed populations of translucent and opaque colonies, whereas the opaque colonies when subcultured produced only opaque colonies. It was hypothesized that the leaky expression of the His6-Rot from pDT112 was not well tolerated by E. coli and resulted in cell lysis. Mutants that failed to produce an intact Rot protein had a growth advantage and outgrew the Rot-expressing cells. Sequence analysis of the rot ORF from different translucent colony-producing clones bearing pDT112 revealed a point mutation of TTT to TCT resulting in an amino acid change of Phe to Ser at position 130. Further sequence analysis revealed that the independently constructed pDT34 also bears the same mutation. This mutational change may be required for survival of the plasmid in E. coli, but this allele is functional, based on complementation studies using pDT34 (34). However, the failure to completely complement the Rot-negative phenotype may be due to this mutation.

The recombinant His6-Rot was purified and used in an EMSA with the 122-bp seb promoter containing DNA fragment and with a 210-bp S. aureus lac promoter fragment as a control. One hundred femtomoles of the promoter-containing DNA fragment was mixed with a 2x to 32x molar ratio of His6-Rot (Fig. 6A). The mobility of the seb promoter-containing DNA was detectably retarded with 2x recombinant protein. An 8:1 molar ratio of Rot to target DNA was required to detect mobility shift with the lac promoter fragment (Fig. 6B). These results suggested that the recombinant Rot is a general DNA binding protein, but the difference in mobility retardation suggested that Rot has a greater affinity for the seb target than lac promoter control DNA fragment. This finding is comparable with the analysis of SarA binding by Sterba et al. (32). They measured the SarA binding affinity to multiple gene promoters that are under SarA regulation, using the trp promoter from E coli as a negative control. Their results demonstrated that affinities of SarA binding to the S. aureus promoter fragments were 1.7- to 5.8-fold higher than the affinity for the trp promoter fragment.



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  		FIG. 6. Electrophoretic mobility shift assays of a 120-bp seb and a 210-bp lac promoter containing DNA fragments incubated with recombinant His6-Rot. Molar ratios of protein to DNA were as follows: 0 (control), lane 1; 2x, lane 2; 4x, lane 3; 8x, lane 4; 16x, lane 5; 32x, lane 6. (A) seb promoter-containing DNA fragment; (B) lac promoter-containing DNA fragment.

 
The promoter deletion studies suggested that a positive transcriptional factor is required for seb expression. To confirm the hypothesis that an additional factor or factors, other than Rot, is involved in the regulation of the seb expression, whole-cell lysates from the wild-type, Agr–, Agr– SarA–, and Agr– Rot– hosts from the post-exponential phase of growth were collected. Various amounts of the cell lysates were incubated with the 122-bp seb promoter DNA fragment (Fig. 7). Retarded electrophoretic mobility of the 122-bp seb promoter fragment was more pronounced when the DNA fragment was incubated with cell lysates collected from the Agr– Rot– hosts than from Agr– or Agr– SarA– hosts. These results suggested that another factor is involved with post-exponential-phase upregulation of the seb promoter activity, and this factor either binds to and retards the mobility of the seb DNA fragment better in the absence of Rot or is present in the lysate in greater amounts in the absence of Rot.



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  		FIG. 7. Electrophoretic mobility shift assay of the wild-type 122-bp seb promoter containing DNA fragment incubated with post-exponential-growth-phase cell lysates collected from Agr–, Agr– Rot–, and Agr– SarA– strains. (–), no lysate added. Two, 4, and 6 µl of lysate were added for the Agr– and Agr– Rot– strains, and 4 µl and 6 µl were added for the Agr– SarA– strain.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The results of this study indicate that Agr-mediated post-exponential-phase enhancement of seb promoter activity is mediated predominantly by the Rot protein. During the exponential phase, Rot represses expression of the enterotoxin B determinant. When the Agr system is activated, the Rot protein is inactivated and expression of seb ensues. However, the evidence also supports the presence of an additional factor necessary for transcription of seb. This factor is a positive transcriptional factor and is largely independent of Agr control. The lack of promoter activity by the core seb promoter and the requirement for upstream sequences for promoter activity to occur support the hypothesis of this positive regulator. The presence of only a single transcriptional start site (8) has been confirmed by us using primer extension (data not shown). This excludes the possibility of the upstream sequences containing additional promoters capable of expressing seb. The effects of other transcriptional factors, such as SigB and SarA, can be explained, for the most part, by their effects on expression of agr or rot. There is evidence in support of SarA, or a SarA-regulated protein, exerting a minor regulatory effect on seb expression.

A model for seb expression can now be made. During the exponential phase of growth, Rot binds to the seb promoter and inhibits transcription by preventing the binding of a yet-unidentified positive transcriptional factor. During the transition to the post-exponential phase of growth, production of RNAIII by the activated Agr system results in the functional inactivation of the Rot protein (20). Loss of Rot binding to the seb promoter frees the DNA to allow the binding of the positive regulator. Rot functions as a repressor of transcription by inhibiting the binding of the positive transcriptional factor. Binding of the positive regulator leads to post-exponential-phase induction of seb expression. The positive transcriptional factor is presumably required to overcome the inhibitory effect of the "kill switch" sequence immediately upstream of the –35 promoter element. The binding of this transcriptional factor, not its production, is subject to Agr control. Evidence for this is that in the absence of Rot, high-level expression of the seb promoter occurs during the exponential growth phase.

The requirement for both positive and negative transcriptional factors may be a common feature of Agr-regulated genes in S. aureus. For example, the protein A determinant (spa) is activated by the SarS protein and binding of this positive regulator is inhibited by the SarA protein, which is a repressor of spa transcription (7). In the case of spa expression, one regulator (SarS) is regulated by the Agr system whereas the other (SarA) is not. The agr upregulated system (seb) utilizes an Agr-regulated repressor (Rot) and an Agr-independent inducer, whereas the Agr downregulated system (spa) utilizes an Agr-regulated inducer (SarS) and an Agr-independent repressor (SarA).

The explanations why the seb promoter is expressed at an intermediate level when Rot-minus mutants also carry an inactive sarA or sarS allele are not known. The effects of SarA or SarS on seb expression are less pronounced in Rot-positive strains.


   ACKNOWLEDGMENTS
 
We thank David George for expert operation of the DNA sequencer and Peter McNamara, Mark Smeltzer, and Markus Bischoff for providing strains.

This work was supported by Public Health Service grant AI45778 from the National Institutes of Health.


   FOOTNOTES
 
* Corresponding author. Mailing address: Life Sciences Center 471E, 1201 Rollins Road, University of Missouri, Columbia, MO 65211-7310. Phone: (573) 884-2866. Fax: (573) 884-9395. E-mail: stewartgc{at}missouri.edu. Back


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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Journal of Bacteriology, August 2005, p. 5301-5309, Vol. 187, No. 15
0021-9193/05/$08.00+0     doi:10.1128/JB.187.15.5301-5309.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.



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