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Infection and Immunity, September 2007, p. 4528-4533, Vol. 75, No. 9
0019-9567/07/$08.00+0     doi:10.1128/IAI.00603-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Mutation of traP in Staphylococcus aureus Has No Impact on Expression of agr or Biofilm Formation

Laura H. Tsang, Sonja T. Daily, Elizabeth C. Weiss, and Mark S. Smeltzer^*

Department of Microbiology and Immunology, University of Arkansas for Medical Sciences, 4301 W. Markham, Little Rock, Arkansas 72205

Received 27 April 2007/ Returned for modification 14 May 2007/ Accepted 25 May 2007

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
To investigate the regulatory role of traP (target of RNAIII-activating peptide) in Staphylococcus aureus, we generated traP mutations in the clinical isolates UAMS-1 and USA300. In neither case did mutation of traP affect expression of the accessory gene regulator (agr) or the ability to form a biofilm. We were also unable to confirm that mutation of traP in the prototype 8325-4 laboratory strain RN6390 results in reduced expression of agr, reduced hemolytic activity, or an altered capacity to form a biofilm.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
There is a growing body of literature indicating that mutation or inhibition of the Staphylococcus aureus regulatory locus designated traP (target of RNAIII-activating peptide) results in reduced expression of the accessory gene regulator (agr), a reduced capacity to form a biofilm, and reduced virulence in animal models of staphylococcal disease (3, 5, 6, 17, 23, 28). The traP gene reportedly encodes a response regulator (TraP) that is triply phosphorylated as a result of the accumulation of a protein designated RAP (RNAIII-activating peptide), resulting in activation of the agr regulatory system and induction of toxin synthesis (21). A number of studies have also concluded that clinical isolates of S. aureus are responsive to RAP and that a modified peptide derivative designated RIP (RNAIII-inhibiting peptide) can be used to limit phosphorylation of TraP and thereby limit induction of agr, the capacity to form a biofilm, and virulence (1, 3, 12, 14, 24).

To the extent that both RAP and RIP reportedly function by modulating the activity of TraP, studies indicating that clinical isolates are responsive to RAP and RIP would suggest that the traP regulatory system functions in the same fashion in most if not all S. aureus strains. However, the primary focus to date, particularly in terms of direct mutagenesis studies, has been on derivatives of the prototype 8325-4 laboratory strain such as RN6390, and recent studies from our laboratory have confirmed that regulatory circuits in RN6390 are different than those observed in at least some clinical isolates (9, 11). One such difference is the impact of agr on biofilm formation. Specifically, mutation of agr enhances biofilm formation in RN6390 but has little impact in clinical isolates (7, 29). Since TraP reportedly functions through an agr-dependent pathway, we wanted to investigate whether similar, strain-dependent differences also existed with respect to traP. To that end, we mutated traP in two clinical isolates (UAMS-1 and USA300-0114) as well as our version of the prototype laboratory strain RN6390. The impact of mutating traP in all three strains was assessed based on biofilm formation, hemolytic activity, and production of the agr-encoded regulatory molecule RNAIII.

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Bacterial strains and growth conditions. The first of the two clinical isolates targeted in this study was UAMS-1, which is a virulent musculoskeletal isolate genetically similar to other prominent clinical isolates, including MRSA252 (EMRSA-16) (8, 10, 11, 19). The second was USA300-0114 (hereafter referred to as USA300), which is the single most predominant strain responsible for the emergence of serious, community-acquired infections caused by methicillin-resistant strains (13, 15, 16). The former is available from the American Type Culture Collection (Manassas, VA), (http://www.atcc.org/) as strain ATCC 49230. The latter was obtained from the culture collection maintained by the Network on Antimicrobial Resistance in Staphylococcus aureus (Herndon, VA) (http://www.narsa.net/). The specific RN6390 strain used in this study was described in previous reports (9, 10, 11). Derivatives of UAMS-1 and RN6390 carrying deletions of the accessory gene regulator (agr) were previously generated by 11-mediated transduction of the tetM agr-null mutation from RN6911 (9). All strains were maintained on tryptic soy agar either without antibiotic selection, with 10 µg erythromycin ml^–1 for USA300 and its mutants, or with 5 µg tetracycline ml^–1 for agr strains.

Mutagenesis. In all cases, traP mutants were generated de novo using the pKOR1 mutagenesis system described by Bae and Schneewind (2). For mutagenesis, opposite ends of the chromosomal region that includes traP were PCR amplified using attB-defined, inward-facing primers corresponding to the attB1 and attB2 sites engineered into pKOR1 together with internal, outward-facing primers containing an engineered SacII site (Table 1). Primer pairs were designed such that an internal region of traP defined by the SacII-containing primers was deleted. Because the traP gene in UAMS-1 is divergent by comparison to 8325 strains and USA300 (10), it was necessary to design specific primers for mutagenesis of UAMS-1 by comparison to both USA300 and RN6390 (Table 1). The two amplified fragments from each strain were cut with SacII, ligated together, reamplified using the flanking attB-defined primers, and then cloned into the pKOR1 vector using the Gateway BP Clonase system (Invitrogen, Carlsbad, CA). The constructs were then transformed into Escherichia coli DH5 (Invitrogen). Plasmids were isolated from E. coli, introduced into RN4220 by electroporation, and then moved from RN4220 into the strain of interest by 11-mediated transduction as previously described (9). After growth at 43°C to drive integration of the plasmid into the chromosome, cultures were shifted to 30°C to promote excision of the plasmid (2). An aliquot of this culture was then plated on medium containing anhydrotetracycline and grown at 30°C to select against clones that retained the plasmid either free or integrated into the traP gene. Confirmation of plasmid loss was subsequently verified by plating on medium containing chloramphenicol, selecting sensitive clones, and verifying the absence of the cat gene by PCR using the primers described by Bae and Schneewind (2). Genotypic confirmation of the mutation in each strain was obtained by PCR using primers flanking the region targeted for deletion (Table 1) and by DNA sequencing of the amplification products. The genetic background of each parent strain and its corresponding mutants was subsequently confirmed by PCR analysis of cna, sarT, and pvl; of the strains targeted in this work, cna is present only in UAMS-1, sarT is present only in RN6390 and USA300, and pvl is present only in USA300.

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TABLE 1. Sequence of primers used in these experiments

Phenotypic characterization. Biofilm formation was assessed using a microtiter plate assay as previously described (7). For determination of hemolytic activity, colonies from overnight streak plates of bacteria grown on tryptic soy agar with antibiotic selection where appropriate were picked to nonselective sheep or rabbit blood agar plates. Blood agar plates were incubated overnight at 37°C. Hemolytic activity was assessed visually as clearing of red blood cells surrounding the area of bacterial growth. To assess relative levels of RNAIII production, total bacterial RNA was isolated using the QIAGEN RNeasy minikit with slight modification. Briefly, each strain was grown in tryptic soy broth to an optical density at 600 nm of 3.0, which represents the postexponential growth phase. An aliquot of each culture containing 5 x 10⁹ cells was mixed with an equal volume of 1:1 acetone-ethanol and stored at –20°C for at least 1 h. Cells were then harvested by centrifugation for 10 min at 4°C. After removal of the supernatant, the pellet was air dried for 10 min and then washed twice in 500 µl Tris-EDTA buffer. The pellet was then resuspended in 500 µl RLT buffer (QIAGEN) containing 10 µl ß-mercaptoethanol per ml. The resuspended cells were then lysed using the FastPrep FP120 cell disruptor as previously described (11). Cellular debris was removed by centrifugation at 13,000 x g for 10 min at 4°C. Two hundred microliters of the upper phase was transferred to a fresh 1.5-ml microcentrifuge tube, and 500 µl of RLT buffer was added, followed by addition of 500 µl ethanol. Samples were then applied to a QIAGEN RNeasy minicolumn and processed according to the manufacturer's instructions, including on-column DNase digestion using the QIAGEN RNase-free DNase kit. Quantitative, real-time reverse transcription-PCR (qRT-PCR) was then performed using the RNAIII-specific primers and probe described by Cassat et al. (11).

Complementation. To complement the traP mutation, the region of DNA extending 438 bp upstream and 352 bp downstream of the traP open reading frame (ORF) was amplified using the primers described in Table 1. The amplification product was cloned into the pCR2.1-TOPO vector (Invitrogen, Carlsbad, CA), excised by digestion with XbaI and HindIII, and then subcloned into pLI50 (9). Ligated constructs were transformed into E. coli DH5 and then into S. aureus RN4220 before 11-mediated transduction (9) from RN4220 into the strains of interest. Expression of traP was subsequently confirmed by qRT-PCR as detailed above, using the traP-specific primers and probe described in Table 1.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION REFERENCES

 
The pKOR1 mutagenesis system was used to generate traP mutations in each of three strains, including the clinical isolates UAMS-1 and USA300. This system allows for the generation of allele replacement mutants at a frequency that eliminates the reliance on antibiotic selection for the identification of mutants (2). This is an important consideration in the specific context of clinical isolates, many of which are resistant to multiple antibiotics and are therefore recalcitrant to conventional allele replacement mutagenesis. Deletion of the targeted region in each strain was verified by PCR (Fig. 1) using primers that flank the region targeted for deletion (Table 1) and by sequencing of the amplified fragments from both the wild-type and mutated strains (data not shown). Sequence analysis confirmed that the UAMS-1 traP allele corresponds to the MRSA252 ORF annotated SAR1926 in The Institute for Genomic Research (TIGR) database (http://www.tigr.org/) and the deletion of a 589-bp region that includes the entire UAMS-1 traP gene. Sequencing also confirmed that the USA300 and RN6390 traP alleles correspond to the ORF annotated NT05SA1752 (8325 genome) and SAUSA3001784 (USA300 genome) in the TIGR database and the deletion of a 415-bp region that includes the 412 bp at the 3' end of the traP gene. PCR analysis of the cna, sarT, and pvl genes also confirmed the genetic background of each mutant (data not shown).

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FIG. 1. Genotypic confirmation of traP mutants. PCR-based confirmation was done by amplifying genomic DNA from the indicated strains using primers that flank the region targeted for mutagenesis (Table 1). The smaller size of the products in the traP mutants relative to those in the corresponding wild-type (WT) strain confirms the internal traP deletion (traP). With respect to RN6390, the mutant labeled traP is the original mutant isolated in the first screen, while those labeled 1, 15, 16, and 20 are the mutants isolated in the second screen. The superscript C (e.g., traPC) indicates a mutant with the complementing plasmid; amplification of the wild-type gene in these strains is indicated by an arrow. Lanes C, E, and M refer to a negative control (no template DNA), empty lanes, and a lane containing molecular size markers, respectively.

Biofilm formation plays an important role in many forms of staphylococcal infection both in terms of contributing to the disease process and in terms of limiting the efficacy of antimicrobial therapy (25). Moreover, previous studies have suggested that mutation of traP or inhibition of TraP phosphorylation using RIP eliminates the capacity of S. aureus to form a biofilm (1, 3, 4, 12, 14, 24). For these reasons, we focused our initial efforts on examining the role of traP in biofilm formation. These experiments were performed using a well-established microtiter plate assay previously shown to yield results consistent with both flow cell assays and in vivo assays employing a murine model of catheter-associated infection (7, 11, 27). Both UAMS-1 and USA300 formed a more robust biofilm than RN6390, and mutation of traP had no impact on biofilm formation in either strain (Fig. 2). As in our previous studies (7), mutation of agr in RN6390 enhanced biofilm formation, while mutation of traP had no impact in four of the five mutants generated in two successive rounds of mutagenesis (Fig. 2). The exception was a single RN6390 traP mutant isolated in the first round of mutagenesis. This mutant exhibited an increased capacity to form a biofilm that was comparable to that of the corresponding RN6390 agr mutant (Fig. 2). The results observed with this mutant are therefore consistent with the suggestion that traP is required for expression of agr (5, 23, 24) However, they also contradict previous reports indicating that mutation or inhibition of traP results in a reduced capacity to form a biofilm (3, 5, 12, 18). More directly, the results that we observed with this mutant are consistent with the hypothesis, put forth by others, that mutation of traP results in reduced expression of agr and with our demonstration that reduced expression of agr enhances biofilm formation in RN6390, but they are in direct contrast to the suggestion that reduced traP function, whether achieved by mutation or inhibition of TraP phosphorylation, can be used as a means of limiting biofilm formation in S. aureus. Indeed, one important issue that has not been addressed in the current traP literature is the discrepancy between reports concluding that traP is required both for activation of agr and for biofilm formation (20, 21, 23, 24, 28) and independent reports from several different laboratories indicating that expression of agr has a negative impact on biofilm formation (7, 29, 30).

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FIG. 2. Biofilm formation as a function of traP. Biofilm formation was assessed using a microtiter plate assay. Results represent quantitative spectrophotometric analysis based on optical density at 595 nm after elution of crystal violet (7). Results with the corresponding RN6390 agr mutant are shown for comparison. Strain order is shown below the graph. The RN6390 derivative designated traP is the original mutant, while those with numerical designators (e.g., traP1) were isolated after the second round of mutagenesis. The identity of all mutants was confirmed by PCR and DNA sequencing as described in the legend to Fig. 1. The inset shows results obtained in an independent assay with RN6390, the original traP mutant (traP), and the complemented mutant (traPC). Complementation was done by introducing a pLI50 construct (9) containing the entire traP gene along with 438 bp of the region upstream of the traP start codon. The asterisk indicates statistical significance relative to the corresponding wild-type strain (P < 0.05). Error bars indicate standard deviations. WT, wild type.

As noted above, mutation of traP in RN6390 had no impact on biofilm formation in four of five mutants. In an attempt to address the discrepancy between the RN6390 traP mutant isolated in the first round of mutagenesis and the four isolated in the subsequent round (see below), we introduced a cloned version of traP consisting of the entire traP gene together with 438 bp upstream of the traP ORF into the original traP mutant (Fig. 1). Although subsequent studies using qRT-PCR confirmed restoration of traP expression (data not shown), complementation of the biofilm phenotype was not successful. This suggested that the increased capacity of this mutant to form a biofilm might be due to a second-site mutation rather than a direct result of the traP mutation itself. One disadvantage of the pKOR1 mutagenesis system is that the resulting mutations are unmarked by a resistance determinant, which effectively eliminates the ability to use pKOR1-generated mutants as transduction donors. This disadvantage is offset by the ability to generate mutants at high frequency, and based on this we took the alternative approach of regenerating the RN6390 traP mutant using the pKOR1 vector described above. We identified four additional RN6390 traP mutants in this second screen (designated the traP1, traP15, traP16, and traP20 mutants). The nature of the traP mutation and the RN6390 background of all four mutants were verified as described above (Fig. 1). It is not possible to definitively determine whether these mutants represent siblings or arose from independent mutagenesis events. Nevertheless, none of these mutants exhibited an altered phenotype by comparison to the RN6390 parent strain with respect to biofilm formation (Fig. 2).

One obvious explanation for the results discussed above is a spontaneous mutation in agr in the original traP mutant, particularly since agr is known to be subject to a high mutation rate in 8325-4 strains such as RN6390 (26). Based on this, and because hemolytic activity in S. aureus is tightly controlled by agr (26), we determined whether mutation of traP in any of the strains we examined also resulted in reduced hemolytic activity. In UAMS-1, hemolytic activity is not apparent on blood agar plates because UAMS-1 is lysogenized with an hlb-converting phage and, like MRSA252 (EMRSA-16), also carries a nonsense mutation in the gene (hla) that encodes alpha-toxin (11). However, in both RN6390 and USA300, mutation of traP had no effect on hemolytic activity as judged using either rabbit (Fig. 3) or sheep (data not shown) blood agar plates. With respect to RN6390, this was true of all five mutants, including the original traP mutant (Fig. 3). These results are important for two reasons. First, they suggest that mutation of traP had no impact on expression of agr in any of the strains we examined. This was subsequently confirmed by qRT-PCR indicating that all three of the wild-type strains, including UAMS-1, expressed RNAIII at levels comparable to those in their corresponding traP mutants (Fig. 4). Second, the results indicate that the increase in biofilm formation observed in the original RN6390 traP mutant was due to an undefined mutation rather a mutation in traP or agr.

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FIG. 3. Hemolytic activity as a function of traP. Hemolytic activity was assessed on rabbit blood agar after overnight incubation. The RN6390 traP mutant in the left panel is the original mutant that exhibited an increased capacity to form a biofilm (Fig. 2). The RN6390 traP mutant in the middle panel (traP1) is representative of the four mutants isolated in the second round of mutagenesis; results with this mutant are shown in comparison to both the RN6390 parent strain and its corresponding agr mutant. Results obtained with the remaining traP mutants isolated in the second round of mutagenesis (traP1, traP15, traP16, and traP20) are shown in the right panel. WT, wild type.

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FIG. 4. Production of RNAIII as a function of traP. Production of RNAIII in the postexponential growth phase was assessed by qRT-PCR as previously described (11). Results with the corresponding UAMS-1 agr mutant (agr), which was generated in a previous study (19), are shown for comparison. Results obtained with the original RN6390 traP mutant (traP) and the original mutant carrying a plasmid-borne version of the wild-type (WT) traP gene (traPC) are shown for comparison to both the corresponding RN6390 agr mutant and each of the four traP mutants (1, 15, 16, and 20) isolated in the second round of mutagenesis.

There are reports concluding that the negative impact of agr on biofilm formation is a function of the surfactant properties of the RNAIII-encoded delta-toxin rather than the regulatory functions of RNAIII itself (29), and it has been demonstrated that the regulatory functions of RNAIII can be retained even after introducing mutations that eliminate the production of delta-toxin (22). Based on this, we sequenced the hld gene from both wild-type RN6390 and the original traP mutant, but sequence analysis confirmed that the hld sequence in both strains corresponds precisely to that in the sequenced strain 8325 (data not shown). At present, we do not know what accounts for the apparent increase in biofilm formation observed in the original traP mutant. However, the more important point in the context of this work is that the inability to complement the mutation, together with the fact that the phenotype was not reproducible, clearly indicates that mutation of traP was not responsible for the increased biofilm formation in the traP mutant. Moreover, like the UAMS-1 and USA300 traP mutants, all of the RN6390 traP mutants, including the traP mutant, exhibited wild-type levels of hemolytic activity and RNAIII production. Based on this, we conclude that mutation of traP has no impact on expression of agr and that inhibition of TraP function is not a viable way to limit biofilm formation in S. aureus, at least in the strains we examined. Importantly, USA300 is a clinically predominant strain, particularly in the context of community-acquired methicillin-resistant S. aureus infection, and UAMS-1 is closely related to other prominent clinical isolates, including the epidemic nosocomial isolate EMRSA-16, but these two strains are not closely related to each other (10, 11, 15, 16). This suggests that our results would be characteristic of a significant proportion of S. aureus clinical isolates and that traP would not be a viable therapeutic target for the treatment of S. aureus biofilm-associated infection. At the same time, our results do not address the suggestion that RIP limits biofilm formation even in clinical isolates of S. aureus (1, 3, 12, 14). It will be interesting to examine the regulatory impact of RIP in these isolates, although it would be anticipated, based on the results reported here, that any alterations in gene expression that might be observed would not be dependent on traP.

 
This work was supported by grant AI074087 to M.S.S. from the National Institute of Allergy and Infectious Disease.

We thank Taeok Bae and Olaf Schneewind for the kind gift of pKOR1. The willingness of Les Shaw and George Stewart to provide helpful comments and to share unpublished data is also greatly appreciated.

 
* Corresponding author. Mailing address: Department of Microbiology and Immunology, University of Arkansas for Medical Sciences, 4301 W. Markham, Mail Slot 511, Little Rock, AR 72205. Phone: (501) 686-7958. Fax: (501) 686-5359. E-mail: smeltzermarks{at}uams.edu

Published ahead of print on 4 June 2007.

Editor: J. B. Bliska

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Infection and Immunity, September 2007, p. 4528-4533, Vol. 75, No. 9
0019-9567/07/$08.00+0     doi:10.1128/IAI.00603-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

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