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Infection and Immunity, April 2009, p. 1623-1635, Vol. 77, No. 4
0019-9567/09/$08.00+0     doi:10.1128/IAI.01036-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Interconnections between Sigma B, agr, and Proteolytic Activity in Staphylococcus aureus Biofilm Maturation {triangledown}

Katherine J. Lauderdale,1 Blaise R. Boles,1 Ambrose L. Cheung,2 and Alexander R. Horswill1*

Department of Microbiology, Roy J. and Lucille A. Carver College of Medicine, University of Iowa, Iowa City, Iowa 52242,1 Department of Microbiology and Immunology, Dartmouth Medical School, Hanover, New Hampshire 037552

Received 19 August 2008/ Returned for modification 29 September 2008/ Accepted 22 January 2009


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ABSTRACT
 
Staphylococcus aureus is a proficient biofilm former on host tissues and medical implants. We mutagenized S. aureus strain SH1000 to identify loci essential for ica-independent mechanisms of biofilm maturation and identified multiple insertions in the rsbUVW-sigB operon. Following construction and characterization of a sigB deletion, we determined that the biofilm phenotype was due to a lack of sigma factor B (SigB) activity. The phenotype was conserved in a sigB mutant of USA300 strain LAC, a well-studied community-associated methicillin-resistant S. aureus isolate. We determined that agr RNAIII levels were elevated in the sigB mutants, and high levels of RNAIII expression are known to have antibiofilm effects. By introducing an agr mutation into the SH1000 or LAC sigB deletion strain, S. aureus regained biofilm capacity, indicating that the biofilm phenotype was agr dependent. Protease activity is linked to agr activity and ica-independent biofilm formation, and we observed that the protease inhibitors phenylmethylsulfonyl fluoride and {alpha}-macroglobulin could reverse the sigB biofilm defect. Similarly, inactivating genes encoding both the aureolysin and Spl extracellular proteases in the sigB mutant restored biofilm capacity. Due to the growing link between murein hydrolase activity and biofilm maturation, autolysin zymography was performed, which revealed an altered profile in the sigB mutant; again, the phenotype could be repaired through protease inactivation. These findings indicate that the lack of SigB activity results in increased RNAIII expression, thus elevating extracellular protease levels and altering the murein hydrolase activity profile. Altogether, our observations demonstrate that SigB is an essential regulator of S. aureus biofilm maturation.


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INTRODUCTION
 
Staphylococcus aureus is the causative agent of a diverse array of acute and chronic infections (48). The ability of S. aureus to attach to surfaces and form a biofilm is a critical determinant of the course of chronic disease (16). The colonized surfaces can be medical implants, such as catheters and orthopedic implants (15), or host tissue, as exemplified by osteomyelitis and infective endocarditis (12, 58). The challenge presented by biofilms is their remarkable resistance to both host defenses and antimicrobial therapies, limiting available treatment options (21).

In recent studies, it has become clear that there are at least two mechanisms of biofilm development in S. aureus (54). One mechanism requires the production of an extracellular polysaccharide, termed polysaccharide intercellular adhesin (PIA), or polymeric N-acetyl-glucosamine. The ica gene cluster is required for the production of PIA, and this locus is essential for biofilm formation in PIA-producing strains (17, 68). Later studies determined that mutations in the ica locus in multiple S. aureus strains do not impair biofilm capacity (6, 11, 19), revealing a second, ica-independent mechanism of biofilm formation. Examination of methicillin-resistant S. aureus (MRSA) strains indicates that these isolates predominantly form the ica-independent biofilms (24, 55), suggesting that there is a need to decipher mechanisms of biofilm maturation and dispersal in the absence of PIA.

Some of the best-studied factors involved in the ica-independent biofilm processes are the agr and sarA global regulatory systems. Both regulators are known to modulate attachment to surfaces but in opposing manners, with SarA being essential for attachment and the agr system controlling the dispersal mechanism (5, 11, 70, 75). Other global regulators, such as the two-component autolysis-related locus ArlRS, have also been implicated in ica-independent biofilms (67). Additionally, surface adhesins, such as the biofilm adhesin protein (Bap) and the SasG adhesin (Aap in Staphylococcus epidermidis), are required for the ica-independent mechanism (14, 18, 19, 62). Proteolytic processing of the SasG adhesin is necessary for attachment (14, 62), and recently we demonstrated that extracellular proteases have an additional role in the dispersal from ica-independent biofilms (11), further outlining the complex nature of biofilm maturation and dispersal in the absence of PIA.

Based on the emerging importance of ica-independent biofilms, we have initiated a screen for chromosomal loci essential for biofilm formation. In preliminary tests, we identified multiple transposon insertions in the rsbUVW-sigB operon. The result of these insertions was a defect in SigB activity, a known global regulator induced in stationary phase and by a variety of environmental stresses, including heat, alkaline, and high-salt conditions (57). Although SigB-defective strains are reported to form biofilms (70), we observed a pronounced biofilm phenotype in sigB mutants using both a methicillin-susceptible strain and a community-associated MRSA (CA-MRSA) USA300 isolate. The characterization of the SigB role in biofilm maturation is the focus of this report.


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MATERIALS AND METHODS
 
Strains and growth conditions. The bacterial strains used in this study are described in Table 1. Strains of Escherichia coli were grown in Luria-Bertani broth or Luria agar plates, and growth medium was supplemented with ampicillin (100 µg/ml) as needed for maintenance of plasmids. Strains of S. aureus were grown in tryptic soy broth (TSB) or tryptic soy agar (TSA). For selection of chromosomal markers or maintenance of plasmids, S. aureus antibiotic concentrations were (in µg/ml) the following: chloramphenicol (Cam), 10; erythromycin (Erm), 10; kanamycin (Kan), 20; and tetracycline (Tet), 5. All reagents were purchased from Fisher Scientific (Pittsburg, PA) and Sigma (St. Louis, MO) unless otherwise indicated.


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  		TABLE 1. Strains and plasmids

Recombinant DNA and genetic techniques. Restriction and modification enzymes were purchased from New England Biolabs (Beverly, MA) and were used according to the manufacturer's instructions. All DNA manipulations were performed in E. coli strain DH5{alpha}-E (Invitrogen, Carlsbad, CA) or strain BW25141 (20). All oligonucleotides were synthesized at Integrated DNA Technologies (Coralville, IA). Plasmids were transformed into S. aureus RN4220 by electroporation as described previously (63). Plasmids were moved from strain RN4220 to other strains using transduction by bacteriophage 80{alpha} as described previously (53). In some cases, chromosomal markers were also moved by bacteriophage 80{alpha} transduction between S. aureus strains. Nonradioactive sequencing was performed at the DNA sequencing facility at the University of Iowa.

Generation of transposon insertions. A plasmid for generating Tn5 insertions in S. aureus was generated using the plasmid pKOR1 (3) as a template. The hypermutable Tn5 transposase gene from plasmid pGRTYB35 (7) was amplified using the oligonucleotides ARH87 (5'-GTTGTTGGTACCTTTCCCGGGAATAATTTTGTTTAACTTTAAGAAGGAG-3') and ARH88 (5'-GTTGTTGGGCCCTTTAGATCTTTTCGTACGTCATATCTTGATCCCCTGCGCC-3'). The PCR product was digested with KpnI and ApaI and ligated into plasmid pKOR1 cut with the same enzymes. This cloning step removed the Gateway region and placed the transposase gene immediately downstream of the Cam resistance gene on pKOR1. Using the BsiWI and BglII restriction sites encoded within oligonucleotide ARH88, a fragment containing the Tn5 mosaic ends and a Kan resistance determinant was cloned onto the plasmid. The mosaic ends were constructed using the oligonucleotides described by Goryshin et al. (28), which flanked a Kan resistance gene from pDG783 (30). The final plasmid was named pTN11 and was used for the initial Tn5 hopping experiments. For Tn5 mutagenesis, plasmid pTN11 was transformed into S. aureus SH1000 and maintained at 30°C on TSA supplemented with Cam. Individual colonies were picked and grown to an optical density at 600 nM (OD600) of 0.4, and dilutions were plated on TSA supplemented with Kan and anhydrotetracycline (400 ng/ml). Plates were incubated at 42°C, and colonies were tested for Cam sensitivity.

For mariner mutagenesis, SH1000 was transformed with plasmids pFA545 and pBursa, and mutagenesis was performed as described previously (2). Mutants were banked in deep-well microtiter titer plates in TSB with 10% glycerol and stored at –80°C. Biofilm and protease phenotypes of the banked mutants were tested in parallel. The biofilm capacity of transposon mutants in microtiter plates was measured as described previously (11). Qualitative protease activity was assessed by testing 48 mutants per plate on milk agar and estimating zones of clearing (37). Transposon insertions described in this report were reconstructed into parent strain SH1000 using bacteriophage 80{alpha}, and protease and biofilm assays were repeated.

Mapping transposon insertions. Transposon insertion sites were mapped using a modification of an arbitrary PCR technique (40). Briefly, amplification was performed using Phusion DNA Polymerase (New England Biolabs) in an MJ Mini thermocycler (Bio-Rad) on chromosomal DNA prepared with a Puregene DNA purification kit (Gentra Systems). Two rounds of PCR were performed. The first round consisted of the following program: 5 cycles of 98°C for 10 s, 30°C for 20 s, and 72°C for 45 s, followed by 30 cycles of 98°C for 10 s, 40°C for 20 s, and 72°C for 45 s, with a final elongation step of 72°C for 10 min. Oligonucleotides for mapping the Tn5 insertion were TN11-For2 (5'-GAGCTATTTTTTGACTTACTGGGG-3') for the 5' end and TN11-Rev2 (5'-GGTCCAATTCTCGTTTTCATAC-3') for the 3' end of Tn11 in conjunction with an arbitrary primer (Arb1, 5'-GGCCACGCGTCGACTAGTACNNNNNNNNNNGATAT-3'). The second round of amplification was performed with the following parameters: 30 cycles of 98°C for 10 s, 50°C for 20 s, and 72°C for 45 s, followed by elongation at 72° for 10 min. Oligonucleotides for the second round were TN11-For1 (5'-GGATGAATTGTTTTAGTACCTAGATTTAG-3') for the 5' end and TN11-Rev1 (5'-CCTATCACCTCAACAATTGAAGCTT-3') for the 3' end in conjunction with an arbitrary primer (Arb2, 5'-GGCCACGCGTCGACTAGTAC-3'). For mapping mariner transposon insertions, the oligonucleotides were changed. For the first round of amplification, the Tn5 oligonucleotides were swapped with the mariner 5' end (TnMarRev1, 5'-GTAAATCAAGTACCAAAATCCG-3') or 3' end (TnMarFor1, 5'-TCCGTATGTTGCATCACCTTCAC-3'), and for the second round, the mariner oligonucleotides were TnMarRev2 (5'-AGTTCCTATATAGTTATACGCGTCTAG-3') for the 5' end or TnMarFor2 (5'-GTGCCCATTAACATCACCATCTA-3') for the 3' end.

Construction of a sigB deletion. Chromosomal deletions of sigB were constructed in SH1000 using the pKOR1 plasmid for allelic replacement as described by Bae and Schneewind (3). Briefly, flanking regions of sigB were amplified using primers SigBupf (5'-GGGGACAAGTTTGTACAAAAAAGCAGGCTCTTAATATAGAAACAACCACTCAAG-3'), SigBupc (5'-ACGCGTGGTACCGCTAGCCATTATTTCGCACCTGCTCTTTTTTTAT-3'), SigBdownc (5'-AAATGAGCTAGCGGTACCACGCGTTAGAATTTGTTTATTAATGATACG-3'), and SigBdownf (5'-GGGGACCACTTTGTACAAGAAAGCTGGGTGTCTTTTTCTTTTGTTTAAAAGGCC-3'). The two PCR products were joined with overlap extension (69) and moved into pKOR1 using the Gateway cloning system (Invitrogen). The resulting plasmid was verified and transformed into SH1000, and the sigB deletion was constructed as described previously (3). The sigB deletion in strain LAC was constructed using the same plasmid and protocol.

Construction of reporter plasmids. (i) pAH1 and pAH6 construction. The asp23 promoter was amplified from S. aureus SH1000 genomic template using the oligonucleotides Asp-Bam (5'-GTTGTTGGATCCTAGCGTTTCTATTAATCGCGATATTATTC-3') and Asp2 (5'-GTTGTTGGTACCAATAGATTCTCCTTTTACTTGTTAATTTTTATA-3'). A second PCR was performed using plasmid pDB59 as a template with the oligonucleotides AspYFP (5'-GGAGAATCTATTGGTACCAACAACAAGAAGGAGATATACATATGAGTAAAGG-3'; YFP is yellow fluorescent protein) and YFP EcoRI (5'-GTTGTTGAATTCTTATTTGTATAGTTCATCCATGCCA-3'). The two PCR products were fused with overlap extension PCR (69), digested with BamHI and EcoRI, and ligated into pDB59 digested by the same enzymes, resulting in plasmid pAH5. To change fluorescent reporters, the mCherry gene was removed from pAH9 (10) using KpnI and EcoRI and ligated into pAH5 cut with the same enzymes. The resulting plasmid, called pAH6, has the asp23 promoter driving the mCherry reporter. To build pAH1, the agr P3 promoter region was PCR amplified from SH1000 genomic DNA with oligonucleotides incorporating HindIII and KpnI sites (for, 5'-GTTGTTAAGCTTCTGTCATTATACGATTTAGTACAATC-3'; rev, 5'-GTTGTTGGTACCTTAAACAACTCATCAACTATTTTCC-3'). The PCR product was cloned into the vector pCR2.1 using a TOPO-TA cloning kit (Invitrogen) according to manufacturer's instructions, generating an intermediate plasmid called pCR2.1-RNAIII. The promoter fragment was removed from this plasmid with BamHI and KpnI and ligated into pAH6 digested with the same enzymes. The resulting plasmid, called pAH1, has the agr P3 promoter driving the mCherry reporter.

(ii) pAH8 and pAH12 construction. A DNA fragment containing the agr P3 promoter was removed from the pCR2.1-RNAIII clone (described above) with HindIII and KpnI, and this fragment was cloned into pAH9 (10) cut with the same enzymes. The resulting vector, called pAH8, has the agr P3 promoter driving the mCherry reporter in an Erm-resistant plasmid. For pAH12, the asp23 promoter was PCR amplified with oligonucleotides Asp1 (5'-GGGAAAAAGCTTTAGCGTTTCTATTAATCGCGATATTATTC-3') and Asp2 (described above) and cloned into the vector pCR2.1 using a TOPO-TA cloning kit (Invitrogen). The promoter fragment was removed with HindIII and KpnI and cloned into pAH9 cut with the same enzymes. The resulting plasmid, called pAH12, has the asp23 promoter driving mCherry reporter in an Erm-resistant plasmid.

Hemolysis assays. Qualitative hemolysis was monitored using sheep blood agar for β-toxin or rabbit blood agar plates for {alpha}-toxin. Quantitative measurements of hemolysis were performed using rabbit blood cell fractions (modified from Jiang et al.) (35). Briefly, blood was pelleted at 8,000 rpm for 10 min in a microcentrifuge tube. Cells were resuspended in 25x volume of RPMI 1640 medium (Invitrogen). For S. aureus samples, strains were grown for 20 h in TSB, and cells were removed by centrifugation and passage through a 0.2-µm-pore-size cutoff filter (Amicon). Hemolytic titers were performed by gently mixing 500 µl of supernatant with 500 µl of blood cell dilutions and incubating samples for 15 min at 37°C. As a reference, total hemolysis was accomplished using 0.2% sodium dodecyl sulfate (SDS). Samples were then centrifuged at 8,000 rpm for 10 min to remove unlysed cells. Release of hemoglobin was measured by monitoring the absorbance at 540 nm. Hemolytic titers were defined as the dilution factor of supernatant required to lyse half of the red blood cells.

Protease assays. Quantitative protease activity measurements were determined using Azocoll (Calbiochem) reagent as described previously (26).

Carotenoid production. The carotenoid staphyloxanthin was extracted with methanol from S. aureus strains (47). Briefly, strains were grown on TSA plates at 37°C for 24 and 48 h. Cells were harvested in sterile water and adjusted to an OD600 of 10. One milliliter was pelleted and resuspended in 1 ml of 100% methanol, and the mixture was vortexed vigorously for 5 min. Cell debris was removed by centrifugation, and a spectral scan was performed using a Beckman DU 7500 spectrophotometer. The absorbance of peaks at 450- and 470-nm wavelengths was evaluated.

Real-time PCR. S. aureus strains were grown in TSB and harvested during stationary phase growth. Levels of RNAIII transcript were determined using quantitative real-time reverse transcription-PCR (qRT-PCR) as described previously (43). RNAIII transcript was amplified using primers RNAIIIfor (5'-AGTCACCGATTGTTGAAATGATATCT-3') and RNAIIIrev (5'AGGAAGGAGTGATTTCAATGGC-3'). As a control, 16S mRNA was amplified as described previously (43).

PIA production. Levels of PIA production were measured using quantitative dot blot analysis based on a modified version of a protocol reported by Cramton et al. (17). Cells were collected from overnight cultures of S. aureus strains grown in 66% TSB with 0.2% glucose. A standard number of cells (5 x 109) was collected by centrifugation and resuspended in 200 µl of 0.5 M EDTA, pH 8. Samples were boiled for 5 min, and then cell debris was cleared by centrifugation at 13,000 rpm for 15 min. After preparation, 5 µl of the sample was applied to a 0.45-µm-pore-size nitrocellulose membrane (Whatman) that was prewashed in sterile water. The membrane was blocked for 1 h in 5% milk before primary antibody was applied (rabbit-anti-PIA at 1:20,000; kindly provided by Paul Fey) and incubated for 1 h. The secondary antibody of horseradish peroxidase-conjugated goat-anti-rabbit antibody (Upstate) was applied for 1 h. Detection was performed using SuperSignal West Pico chemiluminescent substrate (Pierce) according to the manufacturer's specifications and an LAS-1000 luminescent imager from Fuji (Standford, CT) with ImageReader software (Fuji). Quantification was performed using ImageGauge software (Fuji).

Biofilm assays. Microtiter plate biofilms and flow cell biofilms were grown as described previously (11). For culture media, microtiter biofilms were grown in 66% TSB supplemented with 0.2% glucose, and flow cell biofilms were grown in 2% TSB supplemented with 0.2% glucose. For protease inhibition in microtiter biofilms, cells were added to the plate with phenylmethylsulfonyl fluoride ([PMSF] dissolved in 2% dimethyl sulfoxide [DMSO] at a final concentration of 400 µM) or {alpha}2-macroglobulin (final concentration, 0.25 units/ml; Roche). Confocal fluorescence laser microscopy (CLSM) was performed using a Nikon Eclipse E600 microscope with a Radiance 2100 system (Bio-Rad). CSLM and image analysis were performed as described previously (11). For in situ monitoring of biofilm growth using CSLM, S. aureus strains were transformed with the pALC2084 plasmid, which encodes a green fluorescent protein (GFP) reporter under the control of a Tet-inducible promoter (4). For plasmid maintenance and GFP expression, flow cell growth medium was supplemented with 1 µg/ml Cam and 20 ng/ml anhydrotetracycline. For biofilm poststaining, biofilms were treated with 330 nM Syto9 (LIVE/DEAD BacLight Bacterial Viability Kit; Molecular Probes) 15 min prior to visualization.

Protease and autolysin zymography. S. aureus overnight cultures containing the appropriate plasmid were inoculated into 20 ml of TSB medium supplemented with Cam. For sigB mutant complementation, 50 ng/µl anhydrotetracycline was added to induce gene expression. Cultures were grown at 37°C with shaking to an OD600 of 1.0. Cells were pelleted by centrifugation and removed by filtration using a 0.20-µm-pore-size syringe filter. Extracellular protease zymography was performed as described previously (46). Briefly, supernatants were concentrated 100-fold (15 ml to 0.15 ml) with a Centricon 3 concentrator (Millipore, Billerica, MA) at 4°C according to the manufacturer's instructions. Concentrated supernatants were mixed with an equal volume of Laemmli sample buffer, and 20 µl was separated by 8% SDS-polyacrylamide gel electrophoresis (PAGE). Gels were supplemented with 0.2% gelatin (wt/vol) as a protease substrate. To remove SDS following electrophoresis, gels were washed successively with 50-ml portions of 2.5% (vol/vol) Triton X-100 in distilled water (two times for 10 min each time), 2.5% Triton X-100 in Tris buffer (50 mM Tris-HCl, pH 7.4) (two times for 10 min each time), and Tris buffer (two times for 10 min each time). After gels were washed, they were placed in 50 ml of Tris buffer and incubated at 37°C for 15 h. For visualization of protease clearing zones, gels were stained with Coomassie brilliant blue. Autolysin (murein hydrolase) zymography was performed as described previously (42). Cell cultures were grown, and supernatants were prepared and concentrated as described above. Samples were separated on 8% SDS-PAGE gels supplemented with 0.2% Micrococcus luteus cells (wt/vol) as a substrate. SDS was removed as described above, and autolysin activity was visualized with staining using 1% methylene blue in 0.01% KOH, followed by destaining in deionized water.


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RESULTS
 
Isolation of transposon insertions in the rsbU gene. To gain insight on loci involved in biofilm formation, we utilized a new Tn5 transposon system to mutagenize S. aureus strain SH1000. In preliminary tests, several hundred insertions were isolated to verify the activity of the Tn5 system and begin assessment of biofilm capacity. Through pilot tests for biofilm growth in microtiter plates, one of the Tn5 mutants, AH528, was completely defective in biofilm formation (Fig. 1A). Using arbitrary PCR, the insertion was mapped to the rsbU gene (Fig. 1B), encoding a positive regulator of the alternative sigma factor B (SigB). Based on this finding, we began to investigate the role of the SigB system in biofilm formation, which is the focus of the studies described in this report. Unfortunately, through our continued testing of the mutagenesis system, we observed that many of the other isolated Tn5 insertions were siblings. Despite repeated attempts, the problem was not corrected, and we speculate that this is due in part to the unregulated, background expression of the transposase gene, resulting in premature hopping. As an alternative, we began using the bursa aurealis (mariner) mutagenesis system developed by Bae et al. (2), and the sibling issue was not observed using this system.


Figure 1
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  		FIG. 1. Biofilm phenotypes of transposon insertions in the rsbUVW-sigB locus. (A) Microtiter biofilm assays of the wild-type (SH1000), three transposon mutants in rsbUV genes, and constructed {Delta}sigB and {Delta}sigB {Delta}agr mutants. The sarA::Kan mutant was included as a negative control. The crystal violet in each well was solubilized and the OD595 was determined. (B) Graphic map of approximate locations of transposon insertions in the rsbUVW-sigB locus.

In our preliminary screening of ~3,500 mariner transposon mutants, we identified two additional insertions that visually matched the agar plate phenotypes of the rsbU::Tn5 mutant (data not shown). The phenotypes are striking and easily identifiable, highlighted by a lack of carotenoid, hyperprotease production, and hyperhemolysis (see below), and they are consistent with previous reports of mutations in the rsbUVW-sigB locus (56). One of the mariner insertions mapped to rsbU, approximately 600 bp downstream of the Tn5 insertion, and the second mariner insertion mapped to the rsbV gene (Fig. 1B), the anti-anti-SigB factor. Importantly, the rsbU and rsbV mariner insertions were also defective in biofilm formation (Fig. 1A). We are in the process of scaling up the mariner screen for a more complete analysis of the S. aureus loci required for ica-independent biofilm formation.

SigB activity is necessary for biofilm formation. A functional RsbU protein is known to be required for full activity of SigB (27, 56). To determine if the phenotypes observed in the rsbU::Tn5 strain were due to a loss of SigB activity, a sigB deletion was generated using the pKOR1 plasmid (3). Agar plate phenotypes of the {Delta}sigB mutant, such as hemolysis, proteolysis, and carotenoid production, were indistinguishable from the rsbU::Tn5 strain (data not shown). To assess SigB activity, we utilized the asp23 promoter, which is known to be exclusively controlled by SigB (27). An asp23 promoter fusion plasmid (pAH12) was transformed into strains SH1000 (sigB+) and AH1012 ({Delta}sigB). The {Delta}sigB mutant showed the expected loss of asp23-dependent transcription (Fig. 2), indicating that SigB was not functional. As anticipated, the asp23 promoter was also not induced in the rsbU::Tn5 mutant (data not shown).


Figure 2
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  		FIG. 2. Confirmation of the constructed {Delta}sigB mutant. Plasmid pAH12 (Pasp23-mCherry) was used to measure SigB activity from the asp23 promoter in strains SH1000 (sigB+) and AH1012 ({Delta}sigB). Fluorescence readings for the mCherry reporter were taken after 20 h of growth of cells. For complementation, strain AH1012 (with pAH12) was transformed with plasmid pALC2109 (Ptet-sigB), and sigB expression from the tet promoter was induced with 50 ng/ml anhydrotetracycline.

To examine biofilm phenotypes, strains SH1000 (sigB+), AH528 (rsbU::Tn5), and AH1012 ({Delta}sigB) were compared in microtiter assays. Again, the rsbU::Tn5 and {Delta}sigB mutant strains showed similar defects in biofilm formation (Fig. 1A). For a more rigorous examination of biofilm maturation, strains SH1000 and AH1012 were transformed with the Tet-inducible GFP plasmid pALC2084 (4), and the strains were grown in a once-through flow cell system as described previously (11). The biofilms were examined by CLSM, and similar to previously reported results (11), SH1000 formed a robust biofilm ~20 µm thick after 4 days of growth (Fig. 3A). Consistent with the microtiter assay, the {Delta}sigB mutant was defective in flow cell biofilm formation (Fig. 3B).


Figure 3
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  		FIG. 3. Biofilm phenotypes of the {Delta}sigB mutant. For flow cells, strains SH1000 (sigB+) (A) and AH1012 ({Delta}sigB) (B) were transformed with plasmid pALC2084 (induced with 20 ng/ml anhydrotetracycline) to image biofilms with GFP. Biofilms were grown for 4 days in a once-through flow cell setup, and a z-series of images was taken by CLSM and reconstructed with Volocity software. Each side of a grid square is 20 µm in the image reconstruction. For complementation of the {Delta}sigB mutant, strain AH1012 was transformed with plasmid pALC2109. Complementation was assessed with microtiter (C) and flow cell (D) biofilm assays. In the microtiter biofilm, levels of uninduced and induced (50 ng/ml anhydrotetracycline) sigB expression were compared. For the flow cell, a biofilm of strain AH1012 with the sigB expression plasmid pALC2109 (induced with 50 ng/ml anhydrotetracycline) was grown and poststained with Syto9.

To verify the phenotypes of the constructed {Delta}sigB mutant, a complementing plasmid with sigB under the control of the Tet-inducible promoter, plasmid pALC2109 (4), was transformed into AH1012. By performing dose-response tests, we determined that 50 ng/ml anhydrotetracycline inducer was optimal (Fig. 3C), and this induction level was used to complement other {Delta}sigB mutant phenotypes. Levels of carotenoid production, extracellular protease, and {alpha}-hemolysin activity all returned to near-wild-type levels (Table 2). Expression of sigB from pALC2109 also complemented the asp23 promoter defect in the {Delta}sigB mutant (Fig. 2). Importantly, the biofilm phenotype of the {Delta}sigB mutant was complemented in both microtiter and flow cell models (Fig. 3C and D). Altogether, these findings indicate that SigB is essential for S. aureus biofilm formation and that phenotypes of the rsbU::Tn5 mutant were due to lack of SigB activity.


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  		TABLE 2. Phenotypes

The SigB biofilm phenotype is independent of exopolysaccharide. The PIA is important for biofilm formation in S. epidermidis and some strains of S. aureus (17, 29). SigB is known to regulate PIA production, and biofilm phenotypes in SigB-defective strains have been attributed to altered PIA levels (38, 39, 60, 71). Considering these factors, we measured the production of PIA in the {Delta}sigB deletion strain to evaluate whether PIA levels were linked to the biofilm phenotype. As positive producers of PIA, we used S. epidermidis strain ATCC R97-03 (65) and the S. aureus PIA-overproducing strain MN8m (50). For testing PIA levels, S. aureus strains MN8 (sigB+), SH1000 (sigB+), and AH1012 ({Delta}sigB) were used, and for comparison, {Delta}ica::tet deletions in both strain backgrounds were examined as negative controls. PIA dot blot s assays were performed and quantified on a phosphorimager (Fig. 4). Overall, the level of PIA production in S. epidermidis was notably higher than observed in S. aureus, and this difference in levels was also observed using immunofluorescence (25). As anticipated, the MN8 and SH1000 {Delta}ica::tet mutants did not produce detectable levels of PIA. Surprisingly, the AH1012 ({Delta}sigB) strain produced slightly higher levels of PIA than the wild-type SH1000 strain although this difference was not statistically significant (Fig. 4). These observations indicate that changes in PIA levels are not likely the source of the biofilm phenotype in an S. aureus sigB mutant, and they support our recent report that PIA is not essential to form a biofilm in strain SH1000 (11).


Figure 4
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  		FIG. 4. PIA production in {Delta}sigB mutants. Quantitative dot blot s analysis was performed with anti-PIA antiserum. S. epidermidis R97-03 was used as a positive control producer. For S. aureus, MN8m was used as a positive control, and a {Delta}ica mutation served as a negative control for the MN8, SH1000, and LAC genetic backgrounds. For testing, dot blot S assays were performed on strains SH1000 (wild-type [WT]), AH1012 (SH1000 {Delta}sigB), LAC, and AH1096 (LAC {Delta}sigB), along with the indicated controls. The dot blot s results were quantitated with ImageGauge software (Fuji) and plotted relative to S. epidermidis R97-03 (S. epi). P values of <0.05 as calculated by a Student's t test were considered statistically significant.

The interconnection of SigB with the agr quorum-sensing system. Defects in SigB are known to have an impact on the agr quorum-sensing system in S. aureus. It has been reported that a sigB mutation results in increased expression from the agr P3 promoter (33), meaning that RNAIII levels should be higher when SigB is defective. We recently demonstrated that high RNAIII levels have antibiofilm effects (11), suggesting that the induction of the agr system in a sigB mutant could explain the observed biofilm phenotype. To investigate this question, we transformed a P3agr-mCherry reporter plasmid (pAH8) into SH1000 (sigB+) and AH1012 ({Delta}sigB) and compared P3 expression levels. At 8 h of growth in TSB, the relative levels of the P3 promoter were sixfold higher in the SigB-defective strains (Fig. 5A). As further verification, levels of the RNAIII mRNA transcript were evaluated using qRT-PCR. A comparison of SH1000 to AH1012 showed that RNAIII levels increased 11.8-fold in the {Delta}sigB mutant (Fig. 5B), supporting the agr P3 promoter fusion results. In a similar qRT-PCR experiment, RNAIII levels increased nearly 10-fold in the rsbU::Tn5 mutant (AH528) versus SH1000 (data not shown). These observations verify previous reports and implicate high levels of RNAIII expression as contributors to the biofilm phenotype in SigB-defective strains.


Figure 5
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  		FIG. 5. Interconnection between the SigB and agr regulatory systems. (A) Effect of the {Delta}sigB mutation on agr expression. Plasmid pAH8 (P3agr-mCherry) was used to monitor agr activity in strains SH1000 (sigB+) and AH1012 ({Delta}sigB). Fluorescence readings for the mCherry reporter were taken after 8 h of growth in TSB. For complementation, strain AH1012 (with pAH8) was transformed with plasmid pALC2109 (Ptet-sigB), and sigB expression was induced with 50 ng/ml anhydrotetracycline. (B) Analysis of RNAIII levels with quantitative RT-PCR using strains SH1000, AH1012, and AH1012 with complementation plasmid pALC2109. RNAIII levels are relative to 16S RNA control transcript. (C) Flow cell biofilm of strain AH1106 ({Delta}sigB {Delta}agr). Flow cell biofilms were grown for 4 days in a once-through setup, and cells were visualized with CLSM using GFP labeling with plasmid pALC2084 (induced with 20 ng/ml anhydrotetracycline). Each side of a grid square is 20 µm in the CLSM image reconstruction.

To test the RNAIII overproduction hypothesis, a {Delta}sigB {Delta}agr double mutant was constructed and examined for biofilm maturation in both microtiter and flow cell formats. In microtiter plates, introduction of the agr mutation restored biofilm formation compared to the {Delta}sigB single mutant (Fig. 1A). In flow cells, a similar enhancement in biofilm formation was observed (Fig. 5C). For quantitative flow cell assessment, COMSTAT analysis indicated that the {Delta}sigB {Delta}agr double mutant biofilm had 80% of the biomass of the wild-type strain. In summary, our findings indicate that the {Delta}sigB mutant biofilm phenotype is primarily due to the enhanced activity of the agr system.

Increased protease activity contributes to the sigB mutant biofilm phenotype. When agr RNAIII levels are high, S. aureus has increased extracellular protease activity. We recently demonstrated that these increased protease levels have antibiofilm effects (11). Considering that the {Delta}sigB mutation leads to a dramatic increase in RNAIII levels, we reasoned that higher protease levels could contribute to the biofilm phenotype. To address this question, the serine protease inhibitor PMSF was incubated with strains SH1000 (sigB+), SH1002 (sarA::Kan), AH528 (rsbU::Tn5), and AH1012 ({Delta}sigB). The addition of 400 µM PMSF reversed the biofilm phenotype of the rsbU::Tn5 and {Delta}sigB mutants (Fig. 6) but did not alter the phenotype of the sarA::Kan mutant. The 400 µM level of PMSF did not affect growth of the strains, but higher PMSF levels were toxic (data not shown). The solvent used for the PMSF experiment, 2% DMSO, had a minimal effect on biofilm formation. In repeated experiments, we have also found that general protease inhibitor {alpha}2-macroglobulin could repair the biofilm phenotype of SigB-defective strains (data not shown), supporting the PMSF results. Collectively, these observations implicate protease activity as a contributor to the sigB mutant biofilm phenotype.


Figure 6
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  		FIG. 6. Effect of PMSF on biofilm formation in a sarA mutant and SigB-defective strains. Microtiter biofilm assays were performed with SH1000 (sarA+ sigB+), SH1002 (sarA::Kan), AH528 (rsbU::Tn5), and AH1012 ({Delta}sigB). Biofilms were grown with TSB alone, 2% DMSO, or 400 µM PMSF dissolved in DMSO. Biofilm quantitation is plotted relative to the OD595 of SH1000 biofilms.

Inactivation of Aur and Spl proteases restores biofilm formation in a sigB mutant. S. aureus secretes at least 10 extracellular proteases (23). These include the metalloprotease aureolysin (Aur), V8 (SspA), the six Spl serine proteases, and the cysteine proteases staphopain A (ScpA) and staphopain B (SspB). These proteases are encoded in four separate transcripts (aur, sspABC, scpAB, and splABCDEF), and each of these transcripts is upregulated in a SigB-defective strain (8). Based on our PMSF findings, we began inactivating protease genes in the {Delta}sigB mutant strain AH1012. Introduction of a individual {Delta}aur, {Delta}splABCDEF::Erm, or sspA::Erm mutation did not restore biofilm formation in AH1012 (data not shown). However, simultaneous deletion of both the aur and spl loci in a {Delta}sigB mutant restored biofilm capacity in the microtiter assay to wild-type levels (Fig. 7A). The agr system is still functional in the aur spl mutants (11; also data not shown), indicating that the effect is not due to secondary mutations in agr. For a more careful examination of the biofilm phenotype, the GFP plasmid pALC2084 was introduced into strains SH1000 (sigB+), AH1012 ({Delta}sigB), AH750 ({Delta}aur {Delta}spl::Erm), and AH1136 ({Delta}sigB {Delta}aur {Delta}spl::Erm). The strains were grown in flow cells for 4 days and visualized each day by CLSM. As anticipated based on our previous study (11), strains SH1000 and AH750 were robust biofilm formers. In accordance with the microtiter assay results, introduction of the aur and spl protease knockouts restored biofilm capacity to the sigB mutant (Fig. 7B). Altogether, these observations provide evidence that upregulation of the extracellular proteases in SigB-defective strains contributes to the biofilm phenotype.


Figure 7
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  		FIG. 7. Effect of protease mutations on biofilm formation in a {Delta}sigB mutant. (A) Microtiter biofilm assays of SH1000, AH1012 ({Delta}sigB), AH750 ({Delta}aur {Delta}spl), and AH1136 ({Delta}sigB {Delta}aur {Delta}spl) strains. (B) Flow cell biofilm of strain AH1136. Flow cell biofilms were grown for 4 days in a once-through setup, and cells were visualized with CLSM using GFP labeling with plasmid pALC2084 (induced with 20 ng/ml anhydrotetracycline). Each side of a grid square is 20 µm in the CLSM image reconstruction.

Comparisons to a CA-MRSA USA300 strain. In recent years, the CA-MRSA USA300 strain lineage has been identified as the dominant cause of S. aureus infections in many community and hospital settings (22, 44, 59, 64). To determine whether our observed phenotypes in the SH1000 genetic background were consistent in a clinically relevant strain, a {Delta}sigB mutant was constructed in isolate LAC (72). The agar plate phenotypes of the LAC strain versus its {Delta}sigB mutation (strain AH1096) were similar to those in the SH1000 genetic background. Quantitative tests supported this observation as strain AH1096 showed a dramatic reduction in carotenoid production but increased hemolysis and proteolytic activity (Table 2). Strain AH1096 was transformed with the sigB expression plasmid pALC2109, and the plasmid complemented the phenotypes, indicating that the constructed strain did not have secondary defects. To confirm that SigB was not active in strain AH1096, a Pasp23-mCherry promoter fusion plasmid (pAH6) was transformed and tested in strains LAC and AH1096. Again, similar to result in the SH1000 background, the asp23 promoter did not activate in AH1096 under stationary phase conditions (Fig. 8A). RNAIII production was also monitored using a P3agr-mCherry promoter fusion plasmid (pAH1). The levels rose twofold in the {Delta}sigB mutant (Fig. 8B), but the relative increase was not as high as that observed in the SH1000 background (Fig. 5A). The natural level of RNAIII expression in LAC is known to be higher than in methicillin-susceptible S. aureus strains and other MRSA isolates (52, 73).


Figure 8
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  		FIG. 8. LAC wild-type and {Delta}sigB mutant phenotypes. (A) Plasmid pAH6 (Pasp23-mCherry) was used to measure SigB activity from the asp23 promoter in strains LAC (sigB+) and AH1096 ({Delta}sigB). Fluorescence readings for the mCherry reporter were taken after 20 h of growth of cells. (B) Plasmid pAH1 (P3agr-mCherry) was used to monitor agr activity in LAC and AH1096 after 8 h of growth in TSB. (C to G) CLSM reconstructions of flow cell biofilms grown for 4 days. Strains LAC, AH1096, AH1203, and AH1204 were visualized by CLSM using GFP labeling with plasmid pALC2084 (induced with 20 ng/ml anhydrotetracycline). For complementation assessment (E), a biofilm of strain AH1096 with the sigB expression plasmid pALC2109 (induced with 50 ng/ml anhydrotetracycline) was grown in the flow cell and poststained with Syto9. Each side of a grid square is 20 µm in the CLSM image reconstruction.

The LAC strain has not been characterized in models of biofilm formation. We were unable to grow biofilms in microtiter plates, but LAC was a robust biofilm former in the flow cell apparatus. To monitor flow cell biofilms, the LAC (sigB+) and AH1096 ({Delta}sigB) strains were transformed with the GFP-expressing plasmid pALC2084 and grown for 4 days. Through visualization of the biofilms with CLSM, the {Delta}sigB mutation introduced a striking biofilm phenotype compared to the wild-type LAC (Fig. 8C and D). Notably, introduction of the sigB-expressing plasmid complemented the biofilm phenotype (Fig. 8E), demonstrating that the phenotype was due to loss of SigB activity.

To investigate the interconnection between the SigB and agr system in the LAC background, an {Delta}agr mutation was introduced into strains LAC and AH1096, resulting in new strains AH1203 ({Delta}agr) and AH1204 ({Delta}sigB {Delta}agr), respectively. As anticipated, the LAC {Delta}agr mutant (AH1203) was a proficient biofilm former (Fig. 8F). Similar to the result with SH1000, introduction of an {Delta}agr mutation into AH1096 restored biofilm capacity (Fig. 8G). While the {Delta}sigB {Delta}agr double mutant biofilm was as thick as the LAC wild-type biofilm and appeared normal in CLSM image reconstructions (Fig. 8G), closer examination indicated that the biofilm of the double mutant was porous. COMSTAT analysis showed that the {Delta}sigB {Delta}agr double mutant had 50% of the biofilm biomass of the LAC wild-type strain, which is lower than the 80% recovery observed in the SH1000 genetic background (Fig. 5C).

To further compare LAC with SH1000, the {Delta}ica::tet deletion was moved to LAC, generating strain AH1308. Through quantitative dot blot s analysis, the PIA production profile in LAC was determined to be similar to that of SH1000, with the {Delta}ica::tet mutant yielding undetectable levels of PIA and the {Delta}sigB mutation resulting in minor PIA enhancement (Fig. 4). Notably, the LAC {Delta}ica::tet mutant displayed no defect in the flow cell biofilm model compared to wild-type (data not shown). Overall, the {Delta}sigB mutant phenotypes of the LAC strain mirrored those of the SH1000 strain, suggesting that both SigB regulatory systems function in a similar manner.

Murein hydrolases as a protease target. Thus far, we linked the {Delta}sigB mutant biofilm phenotype to the upregulation of the agr system and increased extracellular protease activity. Going forward, what are the proteases cleaving that disables the ability to develop a biofilm? While there are numerous target possibilities, several S. aureus studies have linked defects in murein hydrolase activity to a biofilm phenotype (9, 32, 61, 68). We reasoned that the increase in protease activity in the {Delta}sigB mutant could be impacting the function of murein hydrolases, paralleling observations made in Bacillus subtilis (36, 41). To investigate this hypothesis, we prepared cell-free supernatants of wild-type and {Delta}sigB mutants of strains SH1000 and LAC and performed protease zymography (Fig. 9A) and autolysin zymography (Fig. 9B) on these samples. As anticipated, the {Delta}sigB mutants of both the SH1000 and LAC strains displayed increased protease activity versus the parental wild-type strains (Fig. 9A, lane 2 versus lane 3 for SH1000 and lane 5 versus lane 6 for LAC). The protease enhancement was more significant for SH1000 than LAC, supporting our observations from protease assays with Azocoll reagent (Table 2). When the sigB-complementing plasmid pALC2109 was provided, protease activity decreased (Fig. 9A, lanes 4 and 7). Induction of sigB expression in the LAC background appeared to slightly overcompensate and repress protease production beyond wild-type levels (Fig. 9A, compare lanes 5 and 7), similar to what we observed using Azocoll reagent (Table 2). To further investigate the protease connection, we utilized a double knockout of the {Delta}aur {Delta}spl::Erm loci, which we previously reported to have low extracellular protease activity (11). When these protease knockouts were moved to a {Delta}sigB mutant (strain AH1136), extracellular protease activity was not detectable by zymography in the triple mutant (Fig. 9A, lane 1).


Figure 9
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  		FIG. 9. Protease and autolysin zymography in {Delta}sigB mutants. Gel images are black-white inverted to aid visualization. (A) Extracellular protease zymography using 8% SDS-PAGE supplemented with 0.2% gelatin as a substrate. (B) Autolysin zymography using 8% SDS-PAGE supplemented with 0.2% M. luteus cells as a substrate. Lanes are identical in both protease and autolysin gels. Labels at the top of panel A indicate the genetic background. The presence or absence of SigB is indicated as follows: +, chromosomal sigB; +C, plasmid-encoded sigB. Lane 1, AH1136 ({Delta}sigB {Delta}aur {Delta}spl::Erm); lane 2, SH1000 with pALC2073; lane 3, AH1012 with pALC2073; lane 4, AH1012 with pALC2109; lane 5, LAC with pALC2073; lane 6, AH1096 with pALC2073; lane 7, AH1096 with pALC2109. Expression of sigB from plasmid pALC2109 was induced with 50 ng/ml anhydrotetracycline.

To assess the function of murein hydrolases, the same samples were tested in autolysin zymography using M. luteus cells as a substrate (Fig. 9B). In the {Delta}sigB mutants, we observed a significant increase in the activity of two lower-molecular-mass murein hydrolases in the 30- to 37-kDa size range (Fig. 9B, lanes 3 and 6). In the SH1000 {Delta}sigB mutant (AH1012), a corresponding decrease in the activity of a 50-kDa protein was apparent, but this decrease was not significant in the LAC {Delta}sigB mutant (AH1096). Thus, the lack of SigB activity altered the autolysin profile, resulting in an enhancement in low-molecular-mass murein hydrolases. When sigB was provided on a plasmid, the activity trend changed back to the wild-type profile, demonstrating complementation of the phenotype. Importantly, when the Aur and Spl proteases were deleted in the AH1012 {Delta}sigB mutant, the murein hydrolase profile also returned to the wild-type trend, suggesting that the changes were a direct reflection of the extracellular protease activity level. Altogether, the {Delta}sigB mutants display altered murein hydrolase profiles, which may be a contributing factor to the biofilm phenotype in these mutants.


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DISCUSSION
 
Through random mutagenesis, we identified the SigB system as an important biofilm regulator under ica-independent conditions. We linked the SigB biofilm phenotype to the overexpression of agr RNAIII and demonstrated that increased protease activity was a contributor to the biofilm phenotype. Finally, we observed that the SigB biofilm phenotypes were consistent in a clinical CA-MRSA USA300 isolate.

Our findings clarify the mixed reports on the role of SigB in biofilm maturation (12, 49, 54, 60, 70). Under ica-independent conditions, our results demonstrate that SigB is essential for biofilm formation in multiple strain backgrounds. Valle et al. reported that a sigB mutant of S. aureus strain 15981 formed a biofilm (70), but it was later observed that this strain has an agr mutation (67). As we demonstrated in this report, when the agr system is defective, the SigB biofilm phenotype is masked, perhaps explaining the observed phenotype in strain 15981. Mutations in the ica locus of strain 15981 are also known to alter biofilm development (71), suggesting that this strain forms an ica-dependent biofilm, which may be another explanation for the observation that 15981 sigB mutants retain biofilm capacity.

Our findings implicate the elevated protease levels in a sigB mutant as important contributors to the biofilm phenotype. Considering the results with sigB agr double mutants, the altered protease regulation is likely a result of high RNAIII expression and not direct regulation of the protease genes by SigB. These findings parallel our recent report that demonstrates that extracellular proteases are required for biofilm dispersal (11). Putting these observations together, we propose that regulatory mutations resulting in high protease levels will block S. aureus biofilm attachment and maturation under ica-independent conditions.

Our experiments with the sarA mutant indicate that the biofilm defect results from a different mechanism than that observed for SigB. The protease inhibitor PMSF recovered the sigB mutant defect but did not alter an sarA mutant biofilm, suggesting that the sarA phenotype is more complex. It is important to note that lower sarA expression could have some contribution to the sigB biofilm phenotype. In a sigB agr double mutant background, the biofilm capacity recovered to approximately 80% of the wild-type level in strain SH1000, and the recovery was even lower in strain LAC. SigB is known to be required for full sarA expression (8, 57), and the lower levels of sarA could explain the reduced biomass in the double mutant biofilm.

To compare our results with a clinical isolate, we examined biofilm maturation and the role of SigB in the CA-MRSA strain LAC. USA300 strains are emerging as causative agents of biofilm-related infections (31), and strain LAC is a representative isolate that has recently been the subject of several studies (13, 72, 73). In our hands, LAC attached poorly to microtiter plates, and thus we performed all experiments under flow cell conditions. We speculate that extracellular enzymes and the agr autoinducing peptide are washed away in the flow cells, enabling the LAC strain to attach and develop a biofilm. We have experienced similar microtiter problems with other S. aureus strains that are also alleviated using flow cell conditions (data not shown). Importantly, the extracellular enzyme and biofilm phenotypes of the LAC sigB mutant were similar to those of SH1000, indicating that they are not a strain-specific issue and are consistent in clinical isolates. Although preliminary, these results suggest that the SigB regulatory system functions in an identical manner in strains SH1000 and LAC.

While we have linked the SigB biofilm phenotype to extracellular proteases, the target of the proteases is still unknown. A theme emerging from numerous S. aureus biofilm studies is that murein hydrolases are important for biofilm maturation (9, 61, 68). This link is not limited to S. aureus as similar observations have been made in many gram-positive organisms, including S. epidermidis (32), Streptococcus mutans (1), Enterococcus faecalis (66), and Lactococcus lactis (51). We hypothesized that murein hydrolase function could be modulated by the extracellular protease enzymes, contributing to the observed sigB mutant biofilm phenotypes. In support of this argument, S. aureus and B. subtilis murein hydrolases show susceptibility to proteolytic degradation (41, 74). In S. aureus sigB mutants, the murein hydrolase profile changed, resulting in increased levels of low-molecular-weight activities. When extracellular proteases were inactivated in combination with the sigB mutation, the murein hydrolase profile returned to the wild-type trend, supporting a link between extracellular protease activity and murein hydrolase function. It is possible that the increased protease levels cleaved a high-molecular-weight murein hydrolase to a smaller size or, alternatively, triggered the activation of a dormant enzyme. Considering the numerous secreted protease and murein hydrolase activities, how all the cross-talk occurs at a mechanistic level and further interfaces with biofilm maturation remains an open question. While the autolytic profiles have changed in sigB mutants, the corresponding effect on cell lysis and DNA release are not known and could also be a contributor to the biofilm phenotype.

Altogether, our findings demonstrate that the SigB system is essential for ica-independent biofilm formation in S. aureus. The results of the mutant epistasis studies suggest that SigB is operating upstream of the agr quorum sensing, presumably to respond to environmental cues. Based on our observations and published reports, stress conditions that induce the SigB cascade will favor biofilm formation due to lower agr RNAIII levels, resulting in reduced extracellular protease activity. Conversely, conditions that inhibit SigB activity will increase RNAIII expression and have antibiofilm effects. Thus, conditions that favor low SigB activity will block biofilm maturation although the environmental or host factors that lead to SigB inhibition remain to be defined. It is possible that SigB inhibition will also disperse an established biofilm. In support of this proposal, repression of SigB in established S. epidermidis biofilms led to a pronounced dispersal event (34), suggesting that a similar mechanism may occur in S. aureus. Further insight into the SigB regulatory mechanism would improve our understanding of biofilm maturation and dispersal pathways.


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ACKNOWLEDGMENTS
 
We thank D. Missiakas for providing the mariner transposon plasmids. We thank A. Roth for isolating mariner transposon insertions and M. Thoendel for assistance in sequencing insertion sites. We thank P. Fey for providing PIA antibody.

K. J. Lauderdale was supported by an NIH Predoctoral Training Program in Biotechnology Grant (number T32 GM08365-18). B. R. Boles was supported by an NIH Training Grant (number T32 AI07511). The project was supported by an American Heart Association Beginning Grant-In-Aid and by Award R01AI078921 from the National Institute of Allergy and Infections Diseases.

The content of this paper is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infections Diseases or the National Institutes of Health.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Microbiology, 540F EMRB, University of Iowa, Iowa City, IA 52242. Phone: (319) 335-7783. Fax: (319) 335-8228. E-mail: alex-horswill{at}uiowa.edu Back

{triangledown} Published ahead of print on 2 February 2009. Back

Editor: A. Camilli


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Infection and Immunity, April 2009, p. 1623-1635, Vol. 77, No. 4
0019-9567/09/$08.00+0     doi:10.1128/IAI.01036-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.



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