| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
,
1 Department of Diagnostic and Biological Sciences, School of Dentistry, Medical School, University of Minnesota, Minneapolis, MN 55455, USA
2 Department of Genetics, Cell Biology and Development, Medical School, University of Minnesota, Minneapolis, MN 55455, USA
3 Department of Oral Biology, Joint Health Science Center, University of Florida, Gainesville, FL 32611, USA
4 Mucosal and Vaccine Research Center, Minneapolis VA Medical Center, Minneapolis, MN 55417, USA
Correspondence
Mark C. Herzberg
mcherzb{at}umn.edu
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
These authors contributed equally to this work.
Present address: Novartis Vaccines, Centro Ricerche, Via Fiorentina 1, 53100 Siena, Italy.
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Schachtele et al., 2007). Following entry into the blood, S. gordonii can be isolated from prosthetic heart valves and is a frequent cause of infective endocarditis (Douglas et al., 1993; Sommer et al., 1992). The ability of this species to adhere is essential for survival in all known environments. Among the LPXTG-motif-containing surface proteins, SspA/B, CshA/B and Hsa are expressed by S. gordonii and are essential for adhesion and colonization (Demuth et al., 1996; Holmes et al., 1998; Jakubovics et al., 2005; McNab & Jenkinson, 1998). S. gordonii also utilizes non-LPXTG adhesins, including ScaA, AbpA, AbpB and FbpA (Schachtele et al., 2007). Expressed in S. gordonii and virtually all Gram-positive bacteria (Mazmanian et al., 2002; Scott et al., 2002), sortase A (SrtA) is a membrane-localized transpeptidase that specifically cleaves the LPXTG sequence, and transfers the processed exported N-terminal polypeptide to peptidoglycan, where it covalently cross-links to the cell wall (Paterson & Mitchell, 2004).
Sortase A is responsible for the anchoring of LPXTG-containing surface proteins to the cell wall (Paterson & Mitchell, 2004). In Staphylococcus aureus, a mutation in the srtA gene results in defective anchoring of several proteins, including a number of surface-associated adherence factors (Mazmanian et al., 1999). In Streptococcus mutans biofilms in vitro, genes encoding three putative sortase A-dependent proteins are upregulated when compared to planktonic cells, and mutations in srtA reduce biofilm biomass (Levesque et al., 2005). Consistent with sortase A anchoring of specific adhesive proteins, a recombinant M6 protein of Streptococcus pyogenes containing the LPXTG motif was expressed in S. gordonii GP1223 (Bolken et al., 2001). An isogenic srtA knockout expressed markedly less surface M6 protein and showed reduced binding to immobilized human fibronectin and oral colonization of mice.
The LPXTG surface proteins of S. gordonii can be modelled by SspA and SspB (El-Sabaeny et al., 2000; Holmes et al., 1998; Williams et al., 1994). SspA and SspB are members of the antigen I/II family of proteins (70 % sequence identity), which are expressed by virtually all species of oral streptococci (Demuth et al., 1997). SspA and SspB contain a specific 35-residue C-terminal sorting signal, consisting of an LPXTG sequence motif and hydrophobic and positively charged domains (Fischetti et al., 1990; Schneewind et al., 1992, 1993).
To show how sortase A affects the distribution and function of an endogenous adhesin, we tested the hypothesis that disruption of the srtA gene in S. gordonii changes the localization of SspA and SspB adhesins, biofilm formation, and binding to a specific salivary agglutinin receptor in vitro. Since S. gordonii must adhere to saliva-coated teeth to survive and avoid extinction, we noted that while the srtA– mutants showed loss of function, the organism appeared to adapt to the abnormalities in expression of the LPXTG adhesins. Since we have reported (Zhang et al., 2005) that mutations in S. gordonii SspA and SspB caused upregulation of other adhesin genes to complement loss of adhesion function, we also explored if sortase A might regulate compensating expression of adhesin genes.
|
METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
) were routinely grown in chemically defined synthetic media (FMC) (Terleckyj et al., 1975) for 16–20 h at 37 °C in 5 % CO2. Escherichia coli DH5
cells were grown aerobically at 37 °C in Luria–Bertani (LB) medium. When required, antibiotics were added to the medium at the following concentrations: erythromycin (Em), 5 µg ml–1 (DL1srtA–/V288srtA–); kanamycin (Km), 50 µg ml–1 (E. coli) or 250 µg ml–1 (DL1srtA+).
|
). Plasmids (listed in Table 1
) were purified from E. coli cells using the QIAquick Spin Miniprep Purification Kit (Qiagen). Oligonucleotides were synthesized by Integrated DNA Technologies (listed in Table 2).
|
The V288srtA– mutant was constructed in wtV288 by insertion duplication using the pCR2.1 : B210 plasmid (a gift from Sigma) (Bolken et al., 2001). The mutant was selected for resistance to 5 µg erythromycin ml–1 and confirmed by PCR as described previously (Bolken et al., 2001). Unfortunately, attempts to complement the srtA– mutation in strain V288 were unsuccessful. Thus, the closely related strain, DL1 (Challis), was used for this purpose.
The srtA gene of S. gordonii DL1 was inactivated by allelic exchange with the erythromycin resistance determinant, ermAM. PCR amplification with primers (Table 2) srtA.F1/srtA.R1 and srtA.F2/srtA.R2 of S. gordonii DL1 chromosomal DNA template generated two fragments comprising the flanking sequences of the srtA gene (324 bp, 381 bp), which were ligated via a unique ClaI site and cloned into pGEM-T Easy. A DNA fragment (1023 bp) containing the ermAM gene was PCR-amplified from plasmid pVA891 using primers ermAM.F/ermAM.R that incorporated ClaI restriction sites. The PCR product was digested with ClaI and ligated into the unique site within the combined srtA fragments, generating plasmid pGEM-srtA.ermAM. The insert DNA from this plasmid was PCR-amplified using primers srtA.F1/srtA.R2, purified and transformed into S. gordonii DL1, thus generating the srtA deletion mutant, DL1srtA–. Confirmation of predicted insertions was obtained by PCR amplification and sequencing.
To complement the DL1srtA– mutant, a DNA fragment (1484 bp) incorporating the entire srtA gene was PCR-amplified from S. gordonii DL1 chromosomal DNA using primers srtA.F1/srtA.R2. The PCR product was cloned into E. coli–streptococcal shuttle vector pDL276, generating plasmid pDL276-srtA. This construct was purified and used to transform the DL1srtA– mutant. Confirmation of predicted insertions was obtained by PCR amplification and sequencing. Complementation was confirmed by detection of srtA RNA transcript using primers RealsrtA.F/RealsrtA.R (data not shown). RNA extraction and cDNA synthesis were performed as described below.
No significant difference was found in the growth of any of the mutants compared to their respective wild-type strains (data not shown).
Biofilm formation assay.
As reported previously (Loo et al., 2000), bacteria were grown in 96-well plastic plates at 37 °C for 16–18 h. The biomass was then stained with 25 µl of 1 % (w/v) crystal violet for 15 min and washed to remove planktonic cells and unincorporated stain. To estimate sessile biomass, 250 µl of 70 % ethanol was added to each well. From each well, 125 µl of the extracted crystal violet was transferred to a new well in a polystyrene microtitre plate and the absorbance at 568 nm was determined with an ELISA reader.
Preparation of salivary agglutinin.
Salivary agglutinin was prepared by a modification of a technique reported previously (Rundegren & Arnold, 1987). Unstimulated saliva was collected on ice from healthy volunteers and clarified by centrifugation at 8000 g for 20 min at 4 °C. Agglutinin was prepared by adsorption of the clarified saliva with S. mutans as described previously (Brady et al., 1992). Equal volumes of clarified saliva, phosphate-buffered saline (PBS; 2.7 mM KCl, 1.5 mM KH2PO4, 137 mM NaCl, 6.5 mM Na2HPO4, pH 7.2) and PBS-washed S. mutans NG8 cells (Klett reading of 600) were mixed and incubated on a rotor at 37 °C for 30 min. Cells with adherent agglutinin were removed from the saliva by centrifugation at 2000 g for 15 min and washed once with PBS. Adsorbed agglutinin was eluted from the cells with one volume of PBS containing 1 mM EDTA. Cells were removed by centrifugation (4000 g, 20 min) and the agglutinin preparation was filter-sterilized with a 0.2 µm acrodisc filter (Pall), dialysed against PBS to remove EDTA, and stored in aliquots at –20 °C. Total protein concentration was determined by the bicinchoninic acid (BCA) protein assay (Pierce) and adjusted to 100 ng ml–1.
Binding to salivary agglutinin.
The interaction of S. gordonii with immobilized agglutinin was analysed using the BIAcore 3000 machine with a Pioneer F1 sensor chip, as described by Oli et al. (2006). Salivary agglutinin, isolated as described above, was covalently bound to the activated carboxyl groups of the dextran molecules in the flow cell (FC) 2. The dextran matrix was activated with 35 µl of an equal mixture of N-hydrosuccinimide (115 mg ml–1) and N-ethyl-N'-(dimethylaminopropyl)carbodiimide (750 mg ml–1), as suggested by the manufacturer. Agglutinin (100 ng ml–1) was diluted 1 : 5 (v/v) in 10 mM sodium acetate buffer (pH 5) and 2–3x20 µl was injected manually until the change in resonance units (RU) was 
1500, indicating the immobilization of 1.5 ng protein. The remaining activated dextran was inactivated by injection of 2x35 µl of 1 M ethanolamine. All experiments were conducted at 25 °C in modified adhesion buffer (0.78 mM KH2PO4, 1.22 mM K2HPO4, 50 mM KCl, 1 mM CaCl2 . 6H2O, pH 7.2) (Clark et al., 1978) and the flow rate was maintained at 10 µl min–1 throughout. To serve as a reference surface, FC 1 was treated in the same way, but agglutinin was omitted.
To determine the binding of cells of S. gordonii to purified salivary agglutinin, wtV288 and V288srtA– mutant were grown in 10 ml cultures for 16 h at 37 °C in Todd–Hewitt broth (BBL) supplemented with 0.3 % (w/v) yeast extract, and with 5 µg erythromycin ml–1 for V288srtA–. Cells were harvested by centrifugation and washed once in modified adhesion buffer. After resuspending cells in modified adhesion buffer to 1x1010 c.f.u. ml–1 (5 ml), the bacterial suspension was sonicated for 30 s to disrupt chains and form a single-cell suspension (Sonic Dismembrator, model 100, Fisher Scientific, setting 5) and washed again. Cells were then resuspended in 5 ml modified adhesion buffer and diluted 1 : 10 before use.
S. gordonii cells (1x109 c.f.u. ml–1) in modified adhesion buffer were injected at a flow rate of 10 µl min–1 for 60 s, totalling approx. 1x107 c.f.u. per injection. Adherent bacterial cells were removed by injection of 10 µl regeneration solution [PBS-0.3 % (v/v) Tween-20 with 10 mM Na2EDTA, 100 mM NaCl and 100 mM NaOH], which allowed repeated reuse of the sensor chip surface. S. mutans NG8 cells were analysed for binding as a positive control. Binding of cells to the sensor chip surface was monitored by the change in refractive index over time with resonance signal (RU) plotted on the y-axis and time on the x-axis.
Fractionation of S. gordonii cells.
Cells were fractionated by a modifiction of our previously reported method (Herzberg et al., 1990). Overnight bacterial cultures (100 ml) in FMC were harvested (3000 g, 20 min) and the supernatant and cell pellet collected separately. Supernatants were dialysed against 0.2 % (w/v) Na2EDTA, followed by distilled water, concentrated by lyophilization, and designated the extracellular fraction. Cell pellets were resuspended in 10 ml spheroplasting buffer (20 mM Tris/HCl pH 6.8, 10 mM MgCl2, 26 % (w/v) raffinose), and mutanolysin (Sigma) was added at a final concentration of 80 U ml–1. After incubation at 37 °C for 1 h, suspensions were chilled on ice, and the protoplasts and supernatants then separated by centrifugation (12 000 g, 20 min, 4 °C). Supernatants were collected, dialysed, concentrated by lyophilization and designated the cell-wall fraction. Protoplasts were resuspended in 10 ml of 50 mM triethanolamine/250 mM sucrose, lysed by sonication (50 W, 8 min), and insoluble materials were removed by centrifugation (12 000 g, 20 min, 4 °C). Supernatants were collected, dialysed, concentrated by lyophilization and designated the cytoplasmic fraction. Protein concentrations for each fraction were determined using BCA Protein Assay (Pierce). To compare protein profiles, each fraction (20 µg protein) was resolved by 8 % (w/v) SDS-PAGE and stained with Coomassie blue.
Western immunoblotting.
Proteins were extracted from S. gordonii strains and resolved on 8 % (w/v) SDS-PAGE gels, as described above. Proteins were transferred onto Trans-Blot nitrocellulose membrane (Bio-Rad) at 15 V for 30 min using a Trans-Blot SD Semi-Dry Electrophoretic Transfer Cell (Bio-Rad). Membranes were then probed with rabbit anti-S. mutans P1 serum (1 : 1000 dilution) (gift of Neil Hunter, Millenium Institute of Dental Research, Westmead and Sydney, Australia), and the results visualized with goat anti-rabbit alkaline phosphatase-conjugated IgG (Bio-Rad) and 5-bromo-4-chloro-3-indolylphosphate (BCIP)-Nitro Blue Tetrazolium (Sigma) (Brady et al., 1998; Lee & Boran, 2003). As reported previously, the rabbit anti-S. mutans P1 antibodies (Forester et al., 1983) specifically recognize SspA and SspB of S. gordonii (Heddle et al., 2003; Holmes et al., 1998; Jakubovics et al., 2005).
Immunogold labelling.
Bacteria were grown overnight in FMC medium at 37 °C and diluted to OD620 0.3. Bacterial cultures (1 ml) were pelleted by centrifugation, washed with PBS, resuspended in PBS containing 5 % (v/v) goat serum, and incubated with a 20 µg ml–1 solution of rabbit anti-P1 antibody for 2 h. At 2 h, bacteria were washed three times with PBS-5 % (v/v) goat serum, incubated for 1 h with goat anti-rabbit immunoglobulin conjugated to 10 nm diameter colloidal gold particles (Sigma) diluted 1 : 50, and then washed three times with PBS. The bacteria were examined by backscatter electron imaging using a Hitachi S-900 field emission scanning microscope (5 keV), as described previously (Olmsted et al., 1993). For quantification, gold particles were counted on 30 randomly selected wtV288 and V288srtA– bacterial cells, the identities of which were blinded from the observer.
Whole-cell ELISA.
Overnight cultures (3 ml) of S. gordonii strains were harvested, washed and immobilized in the wells (4x108 cells per well) of 96-well Maxisorp microtitre plates (Nunc), as described previously (Holmes et al., 1995). The wells were then incubated with 0.1 ml blocking buffer [10 mM Tris/HCl pH 8.0, 0.15 M NaCl, 0.1 % (v/v) Tween-20, 2.5 % (w/v) BSA] for 16 h at 4 °C. Next, 50 µl of a 1 : 25 dilution of rabbit anti-P1 antibody in blocking buffer was added to each well and plates were incubated for 2.5 h at room temperature. After three washes with TBST buffer [10 mM Tris/HCl pH 8.0, 0.15 M NaCl, 0.1 % (v/v) Tween-20], 50 µl of a 1 : 125 dilution of goat anti-rabbit IgG antibody conjugated with horseradish peroxidase (Santa Cruz Biotechnology) in blocking buffer was added to each well. After 2.5 h incubation at room temperature, bound antibody was detected using the Stable Peroxide Substrate Buffer (Pierce), following the manufacturer's instructions.
RNA extraction and cDNA synthesis.
RNA extraction from bacterial cultures, confirmation of RNA integrity and purity on agarose gels, and cDNA synthesis were performed as described previously (Zhang et al., 2004, 2005). For each RNA sample, a control cDNA reaction in the absence of reverse transcriptase was performed to check for DNA contamination.
Real-time PCR.
Real-time quantitative PCR was modified from the method of Zhang et al. (2005) using the Mx3000 Real-Time PCR System (Stratagene). PCR reactions (25 µl) comprised 12.5 µl 2x Brilliant SYBR Green QPCR Master Mix (Stratagene), 50 pmol each forward and reverse primer and 3 µl template. The 16S rRNA gene was used as the endogenous reference control, and real-time PCR results were quantified based on standard curves, as described previously (Zhang et al., 2005). The primer set used to amplify the fbpA gene is given in Table 2, whilst the primer sets used for sspA, sspB, scaA, abpA, abpB, hsa, cshA, cshB and 16S rRNA were as described previously (Zhang et al., 2005).
Statistics.
S. gordonii strains were compared for differences in gene expression levels and cell-surface P1 antigen using the Student's t-test.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
).
|
). Salivary agglutinin is the specific receptor on saliva-coated hydroxyapatite for the antigen I/II family of adhesins (Demuth & Irvine, 2002) and for Hsa (Jakubovics et al., 2005; Nobbs et al., 2007). The wtV288 cells bound rapidly to the sensor chip coated with salivary agglutinin. The wtV288 binding curve (sensogram) reflected a change in refractive index (
RU) of 206 during the first 60 s, significantly faster than the V288srtA– mutant (RU=3). Overall binding of the V288srtA– mutant was reduced by 97 %. Similarly, wtDL1 showed a 60 s RU=184. These data were then compared against those of S. mutans, for which the antigen I/II adhesin, SpaP, has been shown to mediate binding to salivary agglutinin (Oli et al., 2006). S. mutans NG8 (wild-type) was used as a positive control and the salivary agglutinin-binding negative S. mutans PC3370 spaP– (isogenic) mutant (Oli et al., 2006) was used as a negative control and these were analysed under identical conditions. Like the V288srtA– mutant, the mutation in the adhesin spaP gene resulted in loss of S. mutans binding to salivary agglutinin. When the sensor was prepared without agglutinin, overall binding of bacterial cells was 90 % less (data not shown). Hence, srtA is required for functional expression of an S. gordonii adhesin(s) for salivary agglutinin with a binding profile similar to SpaP of S. mutans.
Structural effects
Electron microscopy.
To learn whether the srtA– mutation influences cell-wall morphology, the wtV288 and V288srtA– mutant strains were studied with bright-field microscopy, field emission scanning electron microscopy (FESEM) and transmission electron microscopy (TEM). The wild-type and the mutant strains showed similar ultrastructure (data not shown), but the V288srtA– mutant showed fewer immunogold-labelled particles conjugated to anti-P1 reacting with SspA and SspB proteins than wtV288 (Fig. 2). The wtV288 strain bound 8.9±2.5 (mean±SD) gold particles per cell; the V288srtA– mutant bound 2.8±1.7 particles per cell. This difference was statistically significant (P<0.05).
|
). In all strains, lower molecular mass proteins were over-represented in the cytoplasmic and cell-wall fractions when compared to the supernatant fractions. Higher molecular mass proteins were over-represented in the supernatant fractions of V288srtA– and DL1srtA– when compared to the wild-type strains. Complementation of srtA (DL1srtA+) restored the wild-type phenotype protein profile (Fig. 3b).
|
). Anti-P1 antibody reacted with a cell-wall antigen in the wild-type strains, the srtA– mutants and the DL1srtA+ strain. The antigen was not detected in the cytoplasm of any of the strains. P1 antigen was prominent in the supernatant of the deletion mutants (V288srtA–, DL1srtA–) when compared to the wild-types or DL1srtA+ strains, suggesting abundant release of SspA and SspB into the supernatant during growth. The wtDL1 and complemented DL1srtA+ showed a similar distribution of P1 antigen.
To clarify the seeming inconsistency in expression of cell-surface P1 antigen by the srtA– mutant and wild-type, as observed by electron microscopy and Western immunoblotting, a whole-cell ELISA experiment was carried out. When analysed by whole-cell ELISA, the srtA– mutants expressed less cell-surface P1 antigen than the wild-types. Complemented mutant DL1srtA+ expressed the same level of surface P1 antigen as the wild-type DL1 (Fig. 4).
|
We therefore studied the possibility that sortase A might be a general regulator of adhesin gene expression. Using real-time PCR, expression of specific mRNA for the LPXTG-bearing adhesin proteins SspA/B, CshA/B and Hsa, and the non-LPXTG adhesins AbpA/B, FbpA and ScaA, were compared in wtV288 and V288srtA– (Table 3a) and wtDL1, DL1srtA– and DL1srtA+ (Table 3b). Expression of sspA/B, cshA/B, abpA/B and fbpA were significantly upregulated in V288srtA– relative to wtV288 (Table 3a). Expression of scaA and hsa was similar in wtV288 and V288srtA–. The expression profile of adhesin genes in wtDL1 and DL1srtA– (Table 3b) was similar but not identical to wtV288 and V288srtA–. When compared to wtDL1, DL1srtA– genes sspA, cshA/B, abpA/B, hsa and fbpA were all upregulated, although the typical fold-change was less than in the V288 background. Interestingly, scaA was also upregulated 3.2-fold in DL1srtA–. Confirming that srtA regulated expression of the adhesin genes, DL1srtA+ and wtDL1 showed similar levels of expression of adhesin genes (Table 3b).
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Holmes et al., 1998; Williams et al., 1994). The tandem sspA and sspB genes (Demuth et al., 1996) possess individual promoters that are differentially regulated in response to environmental conditions (El-Sabaeny et al., 2000). We hypothesized that functional presentation of SspA/B, the protein products of sspA and sspB genes, requires accessory genes such as srtA. Disruption of the srtA gene has now been shown to change the destination and functions of the SspA/B adhesins, consistent with our hypothesis. Furthermore, sortase A functions more globally in the regulation of adhesin gene expression, apparently optimizing adhesion function in challenging environments and conditions.
Consistent with loss of function of surface adhesins, the sortase A-deficient mutant showed decreased adhesion to immobilized salivary agglutinin, and reduced biofilm-forming ability in vitro. Among these adhesins, SspA and SspB from the srtA– mutants appeared to be over-represented in the extracellular milieu, while still expressed on the cell surface in reduced amounts, as shown by immunoelectron microscopy (Fig. 2) and whole-cell ELISA (Fig. 4). The reduction in cell-surface P1 antigen is consistent with the loss of sortase function. On the other hand, P1 antigen appeared to be overexpressed in the srtA– mutants (Table 3), perhaps accumulating in the interior layers of the cell wall. This speculation is consistent with the observation that total wall-associated SspA and SspB were similar in the wild-type and srtA– mutants when cell-wall fractions were analysed by Western immunoblotting (Fig. 3c, d). These data are consistent with the observations reported previously (Mazmanian et al., 2000), indicating that srtA– mutants of Staphylococcus aureus accumulate non-functional surface protein precursors in their cell wall. Loss of sortase function in the srtA– mutants, therefore, causes improper routeing of the adhesins through the wall.
In the srtA– mutants, LPXTG-containing proteins are synthesized and exported across the plasma membrane into the medium since they cannot bind peptidoglycan in the cell wall. In the absence of sortase A, cell-surface SspA/B is detectable, but less available to bind specific anti-P1 antibodies than on the wild-type strain. Furthermore, the SspA/B expressed on the srtA– mutant cell surface appears non-functional, since V288srtA–, like the prototype S. mutans PC3370 spaP– mutant (Oli et al., 2006), was unable to bind its preferred (but not exclusive) salivary receptor, high molecular mass agglutinin. Hence, sortase A is required for functional presentation of SspA/B and likely other LPXTG adhesins on the cell surface of S. gordonii. In most cases, loss of function could be attributed specifically to srtA since DL1srtA+, which complemented the srtA– mutation, showed complete rescue of adhesin gene expression at the transcriptional level, and was indistinguishable from the wild-type strain. Furthermore, a restoration of adhesion function has been specifically demonstrated for DL1srtA+ (Nobbs et al., 2007).
While the srtA– mutation affected adhesin-related functions and functional expression of SspA/B, cell morphology was not remarkably altered. The V288srtA– and wtV288 cells showed similar bright-field and ultrastructural morphology (data not shown). Furthermore, free-growing V288srtA– and wtV288 cells (and wtDL1, DL1srtA– and DL1srtA+) grew similarly in suspension culture (data not shown). Loss of sortase A activity was, however, associated with a reduction in biofilm formation. During growth of srtA– mutants, our data indicated that SspA and SspB and other LPXTG adhesins may be inappropriately presented on the cell wall and also overexpressed and exported. Soluble adhesins are likely to competitively inhibit cell binding by inappropriately expressed adhesins that are present on the cell wall. Consequently, cells prepared for electron microscopy and salivary agglutinin adhesion assays were washed in advance to remove extracellular proteins, including released SspA and SspB in the culture media. Yet, the presence of poorly functional LPXTG adhesins and soluble exported LPXTG adhesins released by growing cells remained throughout the in vitro biofilm formation assays. Given that adhesion and biofilm formation are adhesin-dependent and that LPXTG adhesins predominate, a sortase A deficiency might, therefore, have been expected to result in greater loss of function than the 36 % reduction observed for V288srtA– relative to wtV288. These observations implied the existence of a compensatory mechanism(s) to maintain at least some level of adhesion.
The data presented here clearly demonstrate that the srtA mutation was accompanied by compensatory changes in adhesin gene expression. Deletion of srtA caused a strain-specific upregulation in adhesin gene mRNAs (abpA/B; LPXTG adhesins sspA/B, cshA/B, hsa). Furthermore, these changes were restored upon complementation of srtA mutation. In V288srtA–, the magnitude of increase in expression of LPXTG-adhesin genes was greater than in DL1srtA–, but in DL1srtA–, scaA, a non-LPXTG-adhesive protein, was substantially upregulated. Hence, S. gordonii cells recognized the loss of srtA, or inappropriate processing, presentation or routeing of LPXTG adhesins, and responded. Several S. gordonii adhesins are known to contribute to transcriptional regulation of alternative adhesins. For example, deletion of sspA/B increases scaA expression at the gene and protein levels (Zhang et al., 2005), HppA and FbpA appear to regulate expression of CshA (McNab & Jenkinson, 1998), while SspB is regulated by SspA (El-Sabaeny et al., 2001). Thus, one might speculate that those adhesin proteins affected by the sortase A mutation subsequently functioned in transcriptional regulation of alternative compensatory adhesins.
While this finding is consistent with a role for sortase A in a regulated network of proteins and genes involved in adhesin-related functions, it remains to be determined how adhesin expression is transcriptionally regulated, although such studies are currently under way. Extensive changes in gene expression are required for S. gordonii to achieve a biofilm state (Gilmore et al., 2003; Zhang et al., 2005), including genes encoding adhesins and transcriptional regulators. Its role as a gene regulator may, therefore, provide a further explanation as to the importance of sortase A in enabling biofilm formation.
Collectively, these data strongly suggest that the adhesins of S. gordonii contribute to the transcriptional control of an array of adhesin genes that form an adhesin maintenance system (AMS). Furthermore, these adhesin protein domains function independently, with no one adhesin solely responsible for adhesion of a streptococcal cell to a specific biological substrate (Gong & Herzberg, 1997). By modulating the expression of these genes, sortase A serves as an additional component of this AMS. Thus, upon improper presentation of adhesins, as occurs upon loss of srtA, the AMS appears to enable the cell wall to remodel its specificity. The cell-wall array of adhesins can be viewed as ‘plastic’ and accommodating to abnormal adhesion. We posit, therefore, that the AMS provides compensatory fail-safe mechanisms that this commensal species needs to persist within the host in the face of constantly changing environmental conditions, and that sortase A forms an essential part of this mechanism.
|
ACKNOWLEDGEMENTS |
|---|
Edited by: P. E. Kolenbrander
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
Brady, L. J., Piacentini, D. A., Crowley, P. J., Oyston, P. C. & Bleiweis, A. S. (1992). Differentiation of salivary agglutinin-mediated adherence and aggregation of mutans streptococci by use of monoclonal antibodies against the major surface adhesin P1. Infect Immun 60, 1008–1017.
Brady, L. J., Cvitkovitch, D. G., Geric, C. M., Addison, M. N., Joyce, J. C., Crowley, P. J. & Bleiweis, A. S. (1998). Deletion of the central proline-rich repeat domain results in altered antigenicity and lack of surface expression of the Streptococcus mutans P1 adhesin molecule. Infect Immun 66, 4274–4282.
Clark, W. B., Bammann, L. L. & Gibbons, R. J. (1978). Comparative estimates of bacterial affinities and adsorption sites on hydroxyapatite surfaces. Infect Immun 19, 846–853.
Demuth, D. R. & Irvine, D. C. (2002). Structural and functional variation within the alanine-rich repetitive domain of streptococcal antigen I/II. Infect Immun 70, 6389–6398.
Demuth, D. R., Duan, Y., Brooks, W., Holmes, A. R., McNab, R. & Jenkinson, H. F. (1996). Tandem genes encode cell-surface polypeptides SspA and SspB which mediate adhesion of the oral bacterium Streptococcus gordonii to human and bacterial receptors. Mol Microbiol 20, 403–413.[Medline]
Demuth, D. R., Duan, Y., Jenkinson, H. F., McNab, R., Gil, S. & Lamont, R. J. (1997). Interruption of the Streptococcus gordonii M5 sspA/sspB intergenic region by an insertion sequence related to IS1167 of Streptococcus pneumoniae. Microbiology 143, 2047–2055.
Douglas, C. W., Heath, J., Hampton, K. K. & Preston, F. E. (1993). Identity of viridans streptococci isolated from cases of infective endocarditis. J Med Microbiol 39, 179–182.
Dunny, G. M., Lee, L. N. & LeBlanc, D. J. (1991). Improved electroporation and cloning vector system for gram-positive bacteria. Appl Environ Microbiol 57, 1194–1201.
El-Sabaeny, A., Demuth, D. R., Park, Y. & Lamont, R. J. (2000). Environmental conditions modulate the expression of the sspA and sspB genes in Streptococcus gordonii. Microb Pathog 29, 101–113.[CrossRef][Medline]
El-Sabaeny, A., Demuth, D. R. & Lamont, R. J. (2001). Regulation of Streptococcus gordonii sspB by the sspA gene product. Infect Immun 69, 6520–6522.
Fischetti, V. A., Pancholi, V. & Schneewind, O. (1990). Conservation of a hexapeptide sequence in the anchor region of surface proteins from gram-positive cocci. Mol Microbiol 4, 1603–1605.[Medline]
Forester, H., Hunter, N. & Knox, K. W. (1983). Characteristics of a high molecular weight extracellular protein of Streptococcus mutans. J Gen Microbiol 129, 2779–2788.
Gilmore, K. S., Srinivas, P., Akins, D. R., Hatter, K. L. & Gilmore, M. S. (2003). Growth, development and gene expression in a persistent Streptococcus gordonii biofilm. Infect Immun 71, 4759–4766.
Gong, K. & Herzberg, M. C. (1997). Streptococcus sanguis expresses a 150-kilodalton two-domain adhesin: characterization of several independent adhesin epitopes. Infect Immun 65, 3815–3821.[Abstract]
Heddle, C., Nobbs, A. H., Jakubovics, N. S., Gal, M., Mansell, J. P., Dymock, D. & Jenkinson, H. F. (2003). Host collagen signal induces antigen I/II adhesin and invasin gene expression in oral Streptococcus gordonii. Mol Microbiol 50, 597–607.[CrossRef][Medline]
Herzberg, M. C., Erickson, P. R., Kane, P. K., Clawson, D. J., Clawson, C. C. & Hoff, F. A. (1990). Platelet-interactive products of Streptococcus sanguis protoplasts. Infect Immun 58, 4117–4125.
Holmes, A. R., Gopal, P. K. & Jenkinson, H. F. (1995). Adherence of Candida albicans to a cell surface polysaccharide receptor on Streptococcus gordonii. Infect Immun 63, 1827–1834.[Abstract]
Holmes, A. R., Gilbert, C., Wells, J. M. & Jenkinson, H. F. (1998). Binding properties of Streptococcus gordonii SspA and SspB (antigen I/II family) polypeptides expressed on the cell surface of Lactococcus lactis MG1363. Infect Immun 66, 4633–4639.
Jakubovics, N. S., Kerrigan, S. W., Nobbs, A. H., Stromberg, N., van Dolleweerd, C. J., Cox, D. M., Kelly, C. G. & Jenkinson, H. F. (2005). Functions of cell surface-anchored antigen I/II family and Hsa polypeptides in interactions of Streptococcus gordonii with host receptors. Infect Immun 73, 6629–6638.
Kuboniwa, M., Tribble, G. D., James, C. E., Kilic, A. O., Tao, L., Herzberg, M. C., Shizukuishi, S. & Lamont, R. J. (2006). Streptococcus gordonii utilizes several distinct gene functions to recruit Porphyromonas gingivalis into a mixed community. Mol Microbiol 60, 121–139.[CrossRef][Medline]
Lee, S. F. & Boran, T. L. (2003). Roles of sortase in surface expression of the major protein adhesin P1, saliva-induced aggregation and adherence, and cariogenicity of Streptococcus mutans. Infect Immun 71, 676–681.
Levesque, C. M., Voronejskaia, E., Huang, Y. C., Mair, R. W., Ellen, R. P. & Cvitkovitch, D. G. (2005). Involvement of sortase anchoring of cell wall proteins in biofilm formation by Streptococcus mutans. Infect Immun 73, 3773–3777.
Loo, C. Y., Corliss, D. A. & Ganeshkumar, N. (2000). Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J Bacteriol 182, 1374–1382.
Macrina, F. L., Evans, R. P., Tobian, J. A., Hartley, D. L., Clewell, D. B. & Jones, K. R. (1983). Novel shuttle plasmid vehicles for Escherichia-Streptococcus transgeneric cloning. Gene 25, 145–150.[CrossRef][Medline]
Mazmanian, S. K., Liu, G., Ton-That, H. & Schneewind, O. (1999). Staphylococcus aureus sortase, an enzyme that anchors surface proteins to the cell wall. Science 285, 760–763.
Mazmanian, S. K., Liu, G., Jensen, E. R., Lenoy, E. & Schneewind, O. (2000). Staphylococcus aureus sortase mutants defective in the display of surface proteins and in the pathogenesis of animal infections. Proc Natl Acad Sci U S A 97, 5510–5515.
Mazmanian, S. K., Ton-That, H., Su, K. & Schneewind, O. (2002). An iron-regulated sortase anchors a class of surface protein during Staphylococcus aureus pathogenesis. Proc Natl Acad Sci U S A 99, 2293–2298.
McNab, R. & Jenkinson, H. F. (1998). Altered adherence properties of a Streptococcus gordonii hppA (oligopeptide permease) mutant result from transcriptional effects on cshA adhesin gene expression. Microbiology 144, 127–136.
Nobbs, A. H., Zhang, Y., Khammanivong, A. & Herzberg, M. C. (2007). Streptococcus gordonii Hsa environmentally constrains competitive binding by Streptococcus sanguinis to saliva-coated hydroxyapatite. J Bacteriol 189, 3106–3114.
Oli, M. W., McArthur, W. P. & Brady, L. J. (2006). A whole cell BIAcore assay to evaluate P1-mediated adherence of Streptococcus mutans to human salivary agglutinin and inhibition by specific antibodies. J Microbiol Methods 65, 503–511.[CrossRef][Medline]
Olmsted, S. B., Erlandsen, S. L., Dunny, G. M. & Wells, C. L. (1993). High-resolution visualization by field emission scanning electron microscopy of Enterococcus faecalis surface proteins encoded by the pheromone-inducible conjugative plasmid pCF10. J Bacteriol 175, 6229–6237.
Pakula, R. & Walczak, W. (1963). On the nature of competence of transformable streptococci. J Gen Microbiol 31, 125–133.
Paterson, G. K. & Mitchell, T. J. (2004). The biology of Gram-positive sortase enzymes. Trends Microbiol 12, 89–95.[CrossRef][Medline]
Rundegren, J. & Arnold, R. R. (1987). Differentiation and interaction of secretory immunoglobulin A and a calcium-dependent parotid agglutinin for several bacterial strains. Infect Immun 55, 288–292.
Sambrook, J., Fritsch, E. F. & Maniatis, T. (1989). Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.
Schachtele, C. F., Nobbs, A. H., Zhang, Y., Costalonga, M. & Herzberg, M. C. (2007). Oral streptococci: commensals and opportunistic pathogens. In Molecular Biology of Streptococci, pp. 411–462. Edited by R. H. S. Chhatwal. Norfolk, UK: Horizon Bioscience.
Schneewind, O., Model, P. & Fischetti, V. A. (1992). Sorting of protein A to the staphylococcal cell wall. Cell 70, 267–281.[CrossRef][Medline]
Schneewind, O., Mihaylova-Petkov, D. & Model, P. (1993). Cell wall sorting signals in surface proteins of gram-positive bacteria. EMBO J 12, 4803–4811.[Medline]
Scott, C. J., McDowell, A., Martin, S. L., Lynas, J. F., Vandenbroeck, K. & Walker, B. (2002). Irreversible inhibition of the bacterial cysteine protease-transpeptidase sortase (SrtA) by substrate-derived affinity labels. Biochem J 366, 953–958.[Medline]
Sommer, P., Gleyzal, C., Guerret, S., Etienne, J. & Grimaud, J. A. (1992). Induction of a putative laminin-binding protein of Streptococcus gordonii in human infective endocarditis. Infect Immun 60, 360–365.
Terleckyj, B., Willett, N. P. & Shockman, G. D. (1975). Growth of several cariogenic strains of oral streptococci in a chemically defined medium. Infect Immun 11, 649–655.
Williams, M. D., Ouyang, T. X. & Flickinger, M. C. (1994). Starvation-induced expression of SspA and SspB: the effects of a null mutation in sspA on Escherichia coli protein synthesis and survival during growth and prolonged starvation. Mol Microbiol 11, 1029–1043.[Medline]
Zhang, Y., Lei, Y., Khammanivong, A. & Herzberg, M. C. (2004). Identification of a novel two-component system in Streptococcus gordonii V288 involved in biofilm formation. Infect Immun 72, 3489–3494.
Zhang, Y., Lei, Y., Nobbs, A., Khammanivong, A. & Herzberg, M. C. (2005). Inactivation of Streptococcus gordonii SspAB alters expression of multiple adhesin genes. Infect Immun 73, 3351–3357.
Received 30 April 2007;
revised 27 July 2007;
accepted 28 August 2007.
This article has been cited by other articles:
|
|
 |
|
  L. S. Turner, T. Kanamoto, T. Unoki, C. L. Munro, H. Wu, and T. Kitten Comprehensive Evaluation of Streptococcus sanguinis Cell Wall-Anchored Proteins in Early Infective Endocarditis Infect. Immun., November 1, 2009; 77(11): 4966 - 4975. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. H. Nobbs, R. J. Lamont, and H. F. Jenkinson Streptococcus Adherence and Colonization Microbiol. Mol. Biol. Rev., September 1, 2009; 73(3): 407 - 450. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. R. Davies, G. Svensater, and M. C. Herzberg Identification of novel LPXTG-linked surface proteins from Streptococcus gordonii Microbiology, June 1, 2009; 155(6): 1977 - 1988. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Zhang, M. Whiteley, J. Kreth, Y. Lei, A. Khammanivong, J. N. Evavold, J. Fan, and M. C. Herzberg The two-component system BfrAB regulates expression of ABC transporters in Streptococcus gordonii and Streptococcus sanguinis Microbiology, January 1, 2009; 155(1): 165 - 173. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. J. Munoz-Elias, J. Marcano, and A. Camilli Isolation of Streptococcus pneumoniae Biofilm Mutants and Their Characterization during Nasopharyngeal Colonization Infect. Immun., November 1, 2008; 76(11): 5049 - 5061. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S.-J. Ahn, S.-J. Ahn, Z. T. Wen, L. J. Brady, and R. A. Burne Characteristics of Biofilm Formation by Streptococcus mutans in the Presence of Saliva Infect. Immun., September 1, 2008; 76(9): 4259 - 4268. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. Steinberg, D. Moreinos, J. Featherstone, M. Shemesh, and O. Feuerstein Genetic and Physiological Effects of Noncoherent Visible Light Combined with Hydrogen Peroxide on Streptococcus mutans in Biofilm Antimicrob. Agents Chemother., July 1, 2008; 52(7): 2626 - 2631. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
| J MED MICROBIOL | ALL SGM JOURNALS | |
