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. Author manuscript; available in PMC: 2013 Feb 1.
Published in final edited form as: Mol Microbiol. 2012 Jan 18;83(4):866–878. doi: 10.1111/j.1365-2958.2012.07970.x

Gap1 functions as a molecular chaperone to stabilize its interaction partner Gap3 during biogenesis of serine-rich repeat bacterial adhesin

Meixian Zhou 1,2, Fan Zhu 1, Yirong Li 1,3, Hua Zhang 1, Hui Wu 1,*
PMCID: PMC3285465  NIHMSID: NIHMS347328  PMID: 22251284

Summary

Serine-rich repeat glycoproteins (SRRPs) are important bacterial adhesins that are conserved in streptococci and staphylococci. Fimbriae-associated protein (Fap1) from Streptococcus parasanguinis, was the first SRRP identified; it plays an important role in bacterial biofilm formation. A gene cluster encoding glycosyltransferases and accessory secretion components is required for Fap1 biogenesis. Two glycosylation-associated proteins, Gap1 and Gap3 within the cluster, interact with each other and function in concert in Fap1 biogenesis. Here we report the new molecular events underlying contribution of the interaction to Fap1 biogenesis. The Gap1 deficient mutant rendered Gap3 unstable and degraded in vitro and in vivo. Inactivation of a gene encoding protease ClpP reversed the phenotype of the gap1 mutant, suggesting that ClpP is responsible for degradation of Gap3. Molecular chaperone GroEL was co-purified with Gap3 only when Gap1 was absent and also reacted with Gap1 monoclonal antibody, suggesting that Gap1 functions as a specific chaperone for Gap3. The N-terminal interacting domains of Gap1 mediated the Gap3 stability and Fap1 biogenesis. Gap1 homologues from Streptococcus agalactiae and Staphylococcus aureus also interacted with and stabilized corresponding Gap3 homologues, suggesting that the chaperone activity of the Gap1 homologues is common in biogenesis of SRRPs.

Introduction

Oral biofilm formation is initiated by adherence of early colonizing streptococci to the oral cavity (Jenkinson & Lamont, 1997; Carlsson et al., 1970; Cisar et al., 1985). Streptococcus parasanguinis FW213 is a primary colonizer of the tooth surface (Froeliger & Fives-Taylor, 2001); it also plays an important role in subacute infective endocarditis (Burnette-Curley et al., 1995). The adhesion of S. parasanguinis FW213 is mediated by its long, peritrichous fimbriae (Burnette-Curley et al., 1995; Fives-Taylor, 1982; Fives-Taylor & Thompson, 1985). Fap1, a serine-rich repeat glycoprotein (SRRP), is the major subunit of the long fimbriae and is required for bacterial adhesion and biofilm formation (Wu et al., 1998; Wu & Fives-Taylor, 1999; Froeliger & Fives-Taylor, 2001). Fap1-like SRRPs from many streptococcal, lactobacillus and staphylococcal species, including both commensal and pathogenic bacteria, are required for bacterial adhesion, biofilm formation and pathogenesis (Zhou & Wu, 2009).

Mature Fap1 is a glycoprotein (Wu et al., 1998; Stephenson et al., 2002). Genes that modulate the glycosylation and biogenesis of Fap1 are clustered in two loci (Zhou & Wu, 2009), gly-gtf3-galT1-galT2 and secY2-gap1-gap2-gap3-secA2-gtf1-gtf2. The gtf1 and gtf2 genes as well as genes from the gly-gtf3-galT1-galT2 locus mediate Fap1 glycosylation (Bu et al., 2008; Zhou et al., 2010; Wu et al., 2007b; Zhu et al., 2011; Chen et al., 2004; Wu et al., 2007a). The secY2-gap1-gap2-gap3-secA2 locus is responsible for secretion of Fap1 (Chen et al., 2004; Wu et al., 2007a), which is conserved in every genome that contains SRRP (Bensing & Sullam, 2002; Siboo et al., 2008; Zhou & Wu, 2009; Zhou et al., 2011).

The conserved secY2-secA2 locus encodes five putative accessory secretory proteins, SecY2, Gap1, Gap2, Gap3 and SecA2. The homologues of these proteins are also involved in the secretion of SRRPs (Chen et al., 2004; Mistou et al., 2009; Bensing & Sullam, 2002; Siboo et al., 2008; Seepersaud et al., 2010; Zhou et al., 2011). The interactions between these accessory secretory components in S. parasanguinis and S. gordonii have been reported. For instance, the interaction between Gap1 and Gap3 is required for Fap1 maturation of S. parasanguinis (Li et al., 2008). Asp1, a Gap1 homologue, also interacts with Asp3, a Gap3 homologue. The interaction is essential for export of GspB to the cell surface of S. gordonii (Seepersaud et al., 2010) implying that the interaction between Gap1 and Gap3 homologues is a common mechanism for the biogenesis of SRRPs. However, it is not understood why the interaction is important for biogenesis of SRRPs.

In this study, we determined that Gap1 functions as a molecular chaperone to interact with Gap3 and stabilize Gap3 in S. parasanguinis; and protease ClpP is responsible for the degradation of Gap3 in the Gap1 mutant. The chaperone activity of the Gap1 homologues is also conserved in S. agalactiae and S. aureus, suggesting the importance of this molecular mechanism in the biogenesis of SRRPs.

Results

Gap1 deficiency impaired production of Gap3

Interaction between Gap1 and Gap3 is required for biogenesis of Fap1 (Li et al., 2008). However, the function of Gap1 in the interaction is unknown. Examination of Gap3 from the gap1 mutant led to an unexpected finding. No endogenous Gap3 protein was detected in the gap1 insertional mutant (Fig. 1A, upper panel). Complementation of the gap1 mutant restored the production of Gap3 (Fig. 1A, upper panel). No difference in production of FimA, a control protein, was observed among different S. parasanguinis variants (Fig. 1A, lower panel). On the other hand, the Gap3 deficiency had no effect on production of Gap1 (Fig. 1B, upper panel). These data demonstrate that Gap1 modulates production of Gap3.

Fig. 1. Production of Gap3 was inhibited in the gap1 mutant.

Fig. 1

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(A) Cell lysates of wild type (WT), gap3 mutant (gap3−), gap1 mutant (gap1−) and the gap1 mutant transformed with pVT1666 (gap1−/pVT1666) or with pVPT-Gap1-GFP (gap1−/pVPT-Gap1-GFP) were subjected to Western blot analysis using Gap3 (upper panel) and FimA (lower panel) antibodies, respectively. (B) Cell lysates of wild type (WT), gap1 mutant (gap1−) and gap3 mutant (gap3−) of S. parasanguinis were subjected to Western blot analysis using anti-Gap1 (upper panel) and anti-FimA (lower panel) antibodies, respectively. (C) The Gap1 deficiency did not affect expression of gap3 by RT-PCR. Total RNA prepared from wild type and the gap1 mutant was reverse-transcribed with M-MLV reverse transcriptase, and used for PCR with gap3-specific primers for gap3 (550bp) and fimA-specific primers for fimA (930bp). Genomic DNA samples were used as PCR controls.

The effect of Gap1 on Gap3 may occur at the transcriptional or translational level. RT-PCR was used to determine expression of gap3. The gap1 mutant still maintained expression of gap3 (Fig. 1C), suggesting that Gap1 deficiency does not regulate Gap3 at the transcriptional level.

Gap1 stabilized Gap3 by preventing Gap3 from degradation

The stability of Gap3 was subsequently evaluated to determine whether Gap1 modulates Gap3 at the translational level. In both recombinant E. coli and S. parasanguinis cells, Gap3 was degraded 60 min after chloramphenicol treatment when it was expressed alone (Fig. 2A & 2C). By contrast, Gap3 remained stable when it was co-expressed with Gap1 (Fig. 2B & 2D). No difference was observed in production of a control protein, HSV-like protein in E. coli and FimA in S. parasanguinis. These results suggest that Gap1 stabilizes Gap3 by preventing it from degradation.

Fig. 2. Gap1 stabilized Gap3.

Fig. 2

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Top10 E. coli cells were transformed with pVPT-Gap3-CHSV (A), or pVPT-Gap1-3-CHSV (B). The gap1 mutant cells were transformed with pVPT-Gap3-CHSV (C) or with pVPT-Gap1-3-CHSV (D). The recombinant bacteria were grown to exponential phase (OD600 = 0.6) and treated with chloramphenicol at 200 µg/ml for 0, 10, 20, 40 and 60 min. The cell lysates were prepared and then subjected to Western blot analysis using anti-Gap3, and anti-HSV antibody, respectively.

Gap1 rendered Gap3 more resistant to in vitro proteolytic digestion

To determine whether Gap1 prevents Gap3 from degradation in vitro, we selected protease endoproteinase Glu-C and trypsin to digest Gap1, Gap3 and the Gap1 and Gap3 complex (Fig. 3). Gap3 alone is sensitive to both proteases (Fig. 3A & B, upper panel). Gap1 is sensitive to trypsin (Fig. 3A, bottom panel) but resistant to endoproteinase Glu-C (Fig. 3B, bottom panel). Gap3 from the Gap1/3 complex was readily degraded when incubating with trypsin (Fig. 3A, middle panel), however the degradation was retarded when treated with endoproteinase Glu-C (Fig. 3B, middle panel) in which Gap1 is resistant to the protease. These data further support the notion that Gap1 stabilizes Gap3 by preventing it from degradation.

Fig. 3. Gap3 alone was more susceptible to proteolytic digestion in vitro.

Fig. 3

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Proteolytic assays for degradation of Gap1, Gap3 and Gap1/3 complex. Purified Gap1, Gap3 and Gap1/3 complex were treated with trypsin (A) and (B) endoproteinase Glu-C, and subjected to SDS-PAGE analysis and stained with Coomassie brilliant blue.

ClpP protease regulated the degradation of Gap3

Proteases are often involved in degradation of misfolded proteins. We hypothesize that a well conserved protease ClpP may degrade Gap3. To test the hypothesis, we constructed a clpP mutant and a gap1/clpP double mutant(VT324/clpP−) to examine the involvement of ClpP in the degradation of Gap3 (Fig. 4). No difference in the Gap3 production was observed between wild type and the clpP mutant. In comparison with a gap1 mutant, VT324, the ClpP deficiency restored the production of Gap3 in the gap1/clpP double mutant, suggesting ClpP is responsible for the degradation of Gap3 in VT324.

Fig. 4. The Clp deficiency restored production of Gap3 in the gap1 mutant.

Fig. 4

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Cell lysates prepared from wild type (WT), clpP mutant (clpP−), VT324 (gap1Δ513–525), double mutant (VT324/clpP−), gap1 mutant (gap1−) and gap3 mutant (gap3−) were subjected to Western blot analysis using Gap3 (upper panel) and FimA (bottom panel) antibody, respectively.

The N-terminal interacting domain of Gap1 modulated the Gap3 stability

Gap1 may stabilize Gap3 through its ability to interact with Gap3. Gap1 deletion variants at Gap1 C-terminal region did not abolish the interactions, and a deletion of amino acid residues 182–190 abrogates its interaction with Gap3 (Li et al., 2008). To determine whether other regions in N-terminus of Gap1 are also involved in the interaction, we constructed four deletions (Δ1–10, Δ11–20, Δ21–28 and Δ29–45) and determined their ability to interact with Gap3 by in vitro GST pull-down assays. The deletion of 1–10, 11–20 and 29–45 reduced the interaction, while deletion of 21–28 had no impact on the interaction (Fig. 5A). The deletion of 1–10, 11–20 and 29–45 significantly reduced the production of Gap3 (Fig. 5B, upper panel) but didn’t impair the Gap1 expression (Fig. 5B, middle panel). These data suggest that the interaction between Gap1 and Gap3 mediates the Gap3 stability.

Fig. 5. The N-terminus of Gap1 was required for the Gap3 stability, for the interaction with Gap3 and for Fap1 maturation.

Fig. 5

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(A) Interaction between Gap1 mutant variants and Gap3. Equal amounts of purified GST, GST-Gap1 and GST-tagged Gap1 mutant variants (Gap1Δ1–10, 11–20, 21–28 and 29–45) bound to glutathione Sepharose 4B beads were incubated with in vitro translated c-Myc-Gap3. The pull-down protein complexes were analyzed by Western blotting using c-Myc monoclonal antibody. Input represents the in vitro translated protein product. (B) Effect of the Gap1 deletions on production of Gap3. Cell lysates of gap1 mutant transformed with the full-length Gap1 and with various Gap1 deletion constructs (Δ1–10, Δ11–20, Δ21–28, Δ29–45) were subjected to Western blot analysis using anti-Gap3 (upper panel), anti-Gap1 (middle panel) and anti-FimA (bottom panel) antibody respectively. (C) Effect of the Gap1 deletions on production of Fap1. Complementation of the gap1 mutant was carried out using the gap1 deletion mutants. Cell lysates of the gap1 mutant transformed with the full-length Gap1 and with various Gap1 deletion mutants were subjected to Western blot analysis using mature Fap1-specific antibody F51.

mAbF51 antibody reacts with mature Fap1. The production of Fap1 was further examined by mAbF51 to determine functional contribution of the Gap3 stability to the biogenesis of Fap1. Deletion of the amino acid motifs 1–10 and 29–45 completely abolished Fap1 maturation, and deletion of 11–20 significantly reduced the production of mature Fap1 (Fig. 5C), suggesting that the motifs that modulated the Gap3 stability were also required for production of mature Fap1. Deletion of the amino acid motifs 21–28 did not affect production of mature Fap1, suggesting the motif that did not regulate the Gap3 stability was not required for the production of mature Fap1 (Fig. 5C). These data indicate that Gap3 stability modulated by the protein-protein interaction is important for biogenesis of Fap1.

Gap1 functioned as GroEL-like molecular chaperone

In the process of purifying recombinant Gap3, we found that a 60 kDa protein consistently co-purified with the recombinant Gap3 protein when Gap1 was absent (Fig. 6A, lane 1). This protein was identified as GroEL by LC/MS/MS analysis. GroEL is a molecular chaperone that mediates folding of many proteins, and is found in numerous bacteria (Zeilstra-Ryalls et al., 1991). Association of GroEL with Gap3 suggested that Gap3 requires a GroEL-like molecular chaperone to maintain its proper folding. No GroEL was co-purified when Gap3 is co-expressed with Gap1 (Lane3). As Gap1 interacts with Gap3 and modulates the Gap3 stability, we hypothesized that Gap1 functions as a molecular chaperone. In fact, a Gap1 monoclonal antibody that reacted with Gap1 (Fig. 6B, lane 2), and Gap1 from the Gap1/3 complex (lane 3), also recognized the GroEL protein co-purified with Gap3 (Fig. 6B, lane 1). These data suggest that Gap1 partially mimics GroEL.

Fig. 6. Gap1 monoclonal antibody recognized GroEL co-purified with Gap3.

Fig. 6

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His-SUMO-tagged Gap3 (Lane 1), Gap1Δ21–28 (Lane 2) and Gap1/Gap3 (Lane 3) proteins were purified with Ni-NTA resin, eluted by cleavage of the SUMO tag and subjected to SDS-PAGE (A) and Western blot analysis using anti-Gap1 (B). A protein band co-purified with Gap3 (Lane 1, top band) was excised, digested with trypsin and subjected to mass spectrometry (LC/MS/MS) to determine the protein identity.

Homologues of Gap1 and Gap3 from S. agalactiae J48 and S. aureus COL interacted with each other

Gap1 and Gap3 are highly conserved in SRRP-containing Gram-positive bacteria. We hypothesize that Asp1 and Asp3, homologues of Gap1 and Gap3, may interact with each other. Indeed, in vitro translated Asp1 (J48) and Asp3 (J48) proteins bound to GST fused -Asp3 (J48) and -Asp1 (J48), respectively, but not to GST (Fig. 7A and B). In addition, HSV-tagged Asp3 (COL) co-purified with GST-Asp1 (COL) but not with GST (Fig. 7C). These data demonstrate that interaction between Asp1 and Asp3 is common in S aureus and S. agalactiae.

Fig. 7. Asp1 interacted with Asp3 from S. agalactiae J48 and S. aureus COL.

Fig. 7

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(A) GST-Asp3 interacted with c-Myc-Asp1. Equal amounts of purified GST and GST-Asp3 (J48) bound to glutathione Sepharose 4B beads were each incubated with in vitro translated c-Myc-Asp1 (J48). (B) GST-Asp1 interacted with c-Myc-Asp3. Equal amounts of purified GST and GST-Asp1 (J48) bound to glutathione Sepharose 4B beads were each incubated with in vitro translated c-Myc-Asp3 (J48). The pull-down protein complexes were analyzed by Western blotting using c-Myc monoclonal antibody. Inputs represent the in vitro translated protein products. (C) Asp1 of S. aureus interacted with Asp3. pGEX-Asp1 (COL) and pVPT-Asp3-CHSV (COL) were co-transformed into E. coli Top10, and recombinant GST-Asp1 was purified using glutathione Sepharose 4B beads and subjected to Western blot analysis with anti-HSV monoclonal antibody. The recombinant E. coli strain harboring pGEX-5X-1 (GST) and pVPT-Asp3-CHSV (COL) was used as a control.

Homologues of Gap1 from S. agalactiae and S. aureus stabilized homologues of Gap3

In order to determine if the Gap1 homologues also stabilize the Gap3 homologues, the stability of Asp3 from S. agalactiae J48 and S. aureus COL was evaluated. Asp3 was readily degraded 20 min after chloramphenicol treatment when only Asp3 was expressed (Fig. 8A & C). By contrast, the Asp3 protein remained stable 60 min after chloramphenicol treatment when it was co-expressed with Asp1 (Fig. 8B & D), suggesting that Asp1 stabilizes Asp3, and that the chaperone-like activity is common in Gram-positive bacteria that produce SRRPs.

Fig. 8. Gap1 homologues stabilized Gap3 homologues.

Fig. 8

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Top10 E. coli cells transformed with pVPT-Asp3-CHSV (J48) (A), pVPT-Asp3-CHSV/pGEX-Asp1 (J48) (B), pVPT-Asp3-CHSV(COL) (C) or pVPT-Asp3-CHSV/pGEX-Asp1 (COL) (D) respectively were grown to exponential phase (OD600 = 0.6), treated with 200 µg/ml of chloramphenicol for 0, 10, 20, 40, and 60 min, harvested and subjected to Western blot analysis using anti-HSV antibody.

Discussion

Biogenesis of SRRPs is mediated by a glycosylation and accessory secretory locus (Zhou & Wu, 2009). The accessory Sec components are highly conserved in many streptococci and staphylococci. Among the accessory components, we and others have determined that Gap1 and Gap3 and their homologues interact with each other and are important for export of SRRPs (Li et al., 2008; Seepersaud et al., 2010). Here we reported a new finding underlying the interaction between Gap1 and Gap3: Gap1 positively regulates production of Gap3; the presence of Gap1 renders Gap3 more stable. A general molecular chaperone GroEL, which is required for the proper folding of many proteins (Zeilstra-Ryalls et al., 1991), was found to be associated with recombinant Gap3 only when Gap1 was absent, indicating that it is important to have a molecular chaperone to assist expression of Gap3. It is likely that when the specific chaperone (Gap1) is not available in the recombinant E. coli cells, Gap3 interacts with a less preferred general chaperone GroEL, which helps to generate limited amount of properly folded recombinant Gap3. Indeed, no detectable GroEL was co-purified when Gap3 was co-expressed with a preferred chaperone Gap1. Furthermore, Gap1-specific monoclonal antibody reacted with the GroEL protein co-purified with Gap3. Together these data support that Gap1 functions as a molecular chaperone. Molecular chaperones are a large group of unrelated protein families that assist protein folding and assembly, stabilize unfolded proteins and assist them for translocation across membranes or degradation (Hendrick & Hartl, 1995; Houry, 2001; Betiku, 2006; Ellis, 1990). The hallmark of molecular chaperones is their ability to interact with and to stabilize their partners. In fact, Gap1 and its homologues from other Gram-positive bacteria exhibit such activities and are thus likely to function as special chaperones for their interacting partners. They may exhibit higher affinity for Gap3 and its homologues than GroEL, which explains why they can stabilize Gap3 and its homologues and prevent them from proteolytic degradation, while GroEL cannot do the same. Furthermore, we demonstrated that the interaction between Gap1 and Gap3 mediates the stability of Gap3. N-terminal deletion mutants that inhibited the interaction concurrently reduced the stability of Gap3 and production of mature Fap1. Interestingly, the Δ11–20 Gap1 mutant still produces considerable amount of mature Fap1 compared with other two mutants, Δ1–10 and Δ29–45, which may suggest that the 1–10 and 29–45 domains are more important than the 11–20 region in some unknown mechanisms. Protein-protein interactions have been recognized as an important factor in stabilizing protein complexes. For instance, the chaperone SseA interacts with type III secretion translocon (TTSS) components (SseB and SseD) directly (Ruiz-Albert et al., 2003), which regulates the stability of SseB and SseD and prevents their premature interactions with SPI-2 TTSS translocon of Salmonella. SseA deficiency prevents the assembly of functional SPI-2 TTSS (Ruiz-Albert et al., 2003; Zurawski & Stein, 2003). SseA also has discrete domains required for SseB stabilization and export (Zurawski & Stein, 2004). A different chaperone SsaE also interacts with SseB and a putative TTSS-associated ATPase, SsaN (Miki et al., 2009) to support the assembly of TTSS in Salmonella, thereby contributing to bacterial virulence.

How the interaction between Gap 1 and Gap3 mediates the stability of Gap3 is unknown. It is possible that the interaction situates Gap3 in a stable protein complex where it maintains a native conformation that is resistant to protein degradation by certain protease(s). As Gap3 is unstable in both E. coli and streptococcal recombinant systems, it is reasonable to speculate that a well conserved protease activity is involved in the process. Such a conserved protease complex exists. ClpP, a well conserved protease subunit, plays an important role in intracellular protein degradation in both Gram-negative and Gram-positive bacteria (Thomsen et al., 2002; Chastanet & Msadek, 2003; Lemos & Burne, 2002). Indeed, our data suggested that the ClpP protease could degrade Gap3 in the gap1 mutant of S. parasanguinis. As a bacterial protease complex, the ClpP system may mimic the eukaryotic proteasome, which functions in protein degradation and quality control (Porankiewicz et al., 1999). In this regard, the Gap1 and Gap3 complex and ClpP protease may constitute a bacterial proteasome, engaging in quality control of Fap1 biogenesis.

In addition, Gap1 may contribute to the stability of Gap3 by modifying Gap3, thereby stabilizing Gap3. Gap1 is predicted to be a glycosytlransferase as it has a domain of unknown function (DUF1975), which is conserved in hundreds of known glycosyltransfersaes. Furthermore, Gap1 affects glycosylation and maturation of Fap1 (Li et al., 2008). Whether Gap1 functions as a glycosylatransferase using Gap3 as a substrate is unknown. It has been shown that O-fucosyltransferase 1 plays an important role in Notch signaling via its distinct chaperone and glycosyltransferase activity by different domains (Okajima et al., 2005; Okajima et al., 2008). Thus, it is not surprising that Gap1 may have a dual function, the chaperone activity through the interaction domain and the glycosyltransferase activity through other domains. Many posttranslational modifications stabilize interacting partners (Ju et al., 2011; Arnesen et al., 2010). Therefore, it is equally plausible that Gap1 may modify Gap3 by other means of posttranslational modification.

Why is maintaining the stability of Gap3 so important in Fap1 biogenesis? We have demonstrated that Gap3 is critical for glycosylation and stability of Fap1 (Peng et al., 2008a; Peng et al., 2008b). A recent study suggested that Asp3, a Gap3 homologue from S. gordonii, is a key scaffolding protein that organizes the accessory Sec complex (Seepersaud et al., 2010), thereby affecting biogenesis of GspB, a Fap1 homologue. Thus, destabilization of Gap3, an Asp3 homologue, may also lead to the formation of a faulty accessory Sec complex, which may render bacteria fail to form mature Fap1.

Asp1 from S. agalactiae and S. aureus, Gap1 homologues, also interact with and stabilize Asp3, Gap3 homologues, suggesting that the molecular mechanism of the interaction between Gap1 and Gap3, and the chaperone activity exhibited by Gap1 and its homologues are common in the biogenesis of SRRPs. Furthermore, numerous Gap1 and Gap3 homologues are present in many other pathogenic bacteria such as Streptococcus pnemoniae and S. epidermidis. Our finding of Gap1 as a molecular chaperone not only sheds new insights into the biosynthetic mechanisms of SRRPs, but also provides us with opportunities to design potential therapeutics targeting this unique protein-protein interaction mechanism to control Gram-positive bacterial infection that is dependent on SRRPs.

Experimental procedures

Bacterial strains, plasmids and growth conditions

Bacterial strains and plasmids used in this study are listed in Table 1. Streptococcal strains and S. aureus COL were cultivated statically in 5% CO2 in Todd–Hewitt (TH) broth or on TH agar plates at 37°C. E. coli strains were grown in Luria–Bertani (LB) broth at 37°C with shaking or on LB agar plates at 37°C. Culture medium was supplemented with antibiotics at the following concentrations: E. coli, ampicillin and kanamycin (50 µg ml−1) and erythromycin (300 µg ml−1); S. parasanguinis, erythromycin (10 µg ml−1) and kanamycin (125 µg ml−1).

Table 1.

Strains and plasmids used in this study.

Strains/Plasmids Relevant Properties Source
Strains
E. coli Top10 Host for propagation of the recombinant plasmids Invitrogen
E. coli BLR(DE3) pET system host strain Invitrogen
S. parasanguinis FW213 Wide type (Cole et al., 1976)
    gap1− Wild type; gap1::aphA3; Kanr (Li et al., 2008)
    gap2− Wild type; gap2::aphA3; Kanr Wu et al. unpublished
    gap3− Wild type; gap3::aphA3; Kanr (Li et al., 2008)
    VT324 Chemically mutagenized gap1 mutant (Zhou et al., 2008b)
    Clp− Wild type; clp::aphA3; Kanr This study
    VT324/clp− VT324; clp::aphA3; Kanr This study
S. agalactiae J48 wild type Brady
S. aureus COL wild type (Gill et al., 2005)
Plasmids
BD::Gap3 gap3 from FW213 cloned in BD vector; Kanr (Li et al., 2008)
BD::Asp1 asp1 from J48 cloned in BD vector; Kanr This study
BD::Asp3 asp3 from J48 cloned in BD vector; Kanr This study
pET28-SUMO SUMO fusion protein expression vector; Kanr Ma
pET28-SUMO-Gap1Δ21–28 Gap1Δ21–28 cloned in pET28-SUMO; Kanr This study
pET28-SUMO-Gap3 gap3 from FW213 cloned in pET28-SUMO; Kanr This study
pET28-SUMO-Gap1-3 gap1 and gap3 from gap2− cloned in pET28-SUMO; Kanr This study
pGEM-ClpP clpP and its upstream 730bp flanking region from FW213 cloned in pGEM-T Easy This study
pGEM::ΔclpP-aphA3 ΔclpP-aphA3 cloned in pGEM-T Easy This study
pGEX-5X-1 GST fusion protein expression vector; Ampr Amersham
pGEX-Gap1 gap1 from FW213 cloned in pGEX-5X-1; Ampr This study
pGEX-Asp1 (J48) asp1 from J48 cloned in pGEX-5X-1; Ampr This study
pGEX-Asp3 (J48) asp3 from J48 cloned in pGEX-5X-1; Ampr This study
pGEX-Asp1 (COL) asp1 from COL cloned in pGEX-5X-1; Ampr This study
pGEX-Gap1Δ1–10 aa 1–10 of Gap1 deleted from pGEX-Gap1; Ampr This study
pGEX-Gap1Δ11–20 aa 11–20 of Gap1 deleted from pGEX-Gap1; Ampr This study
pGEX-Gap1Δ21–28 aa 21–28 of Gap1 deleted from pGEX-Gap1; Ampr This study
pGEX-Gap1Δ29–45 aa 29–45 of Gap1 deleted from pGEX-Gap1; Ampr This study
pVT1666 E. coli-Streptococci shuttle vector; Ermr (Chen et al., 2006)
pVPT-Gap1 gap1 from FW213 cloned in pVT1666; Ermr (Zhou et al., 2008b)
pVPT-CHSV E. coli-Streptococci shuttle vector; Ermr (Zhou et al., 2008a)
pVPT-Gap3-CHSV gap1 from FW213 cloned in pVPT-CHSV; Ermr This study
pVPT-Gap1-3-CHSV gap1 and gap3 from gap2− cloned in pVPT-CHSV; Ermr This study
pVPT-Asp3-CHSV (J48) asp3 from J48 cloned in pVPT-CHSV; Ermr This study
pVPT-Asp3-CHSV (COL) asp3 from COL cloned in pVPT-CHSV; Ermr This study
pVPT-Gap1Δ1–10 Gap1Δ1–10 cloned in pVT1666; Ermr This study
pVPT-Gap1Δ11–20 Gap1Δ11–20 cloned in pVT1666; Ermr This study
pVPT-Gap1Δ21–28 Gap1Δ21–28 cloned in pVT1666; Ermr This study
pVPT-Gap1Δ29–45 Gap1Δ29–45 cloned in pVT1666; Ermr This study
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DNA manipulation

Extraction of genomic DNA from S. parasanguinis, S. agalactiae J48 and S. aureus COL was carried out using the PUREGENE® DNA Isolation Kit (Gentra Systems). PCR amplification was performed using KOD Hot Start DNA polymerase (Novagen) or Taq DNA Polymerase (Promega). PCR Primers are listed in Table 2.

Table 2.

Primers used in this study.

Table Sequences
FimA-1F ATGAAAAAAATCGCTTCTGTCC
FimA-930R TTACTGACTCAATCCTTCTGC
Gap1-Notl-1F AAGGAAAAAAGCGGCCGCATGTTTTATTTTGTACCTTCTTGG
Gap1-SalI-F1 CGTCAGTCGACATGTTTTATTTTGTACCTTC
Gap1-SalI-F2 ACGCGTCGACGTATGTTTTATTTTGTACCTTCTTGG
Gap1-KpnI-R GATCAGGTACCTTTCTTTTTTAGCATACCTTTC
Gap1-Xhol-R ACCGCTCGAGTTATTTCTTTTTTAGCATACC
Gap3-SalI-F1 CGGCCGTCGACATGACTAAACAGTTAATTTC
Gap3-SalI-F2 ACGCGTCGACGTATGACTAAACAGTTAATTTCTG
Gap3-KpnI-R GATCAGGTACCAATATATTCTATTAAATTTTTCAC
Gap3-XhoI-R ACCGCTCGAGTTAAATATATTCTATTAAATTTTTC
ClpP-F ACATCCACTCATTCAACAC
ClpP-R CTATCCTTCTGTGACCACAG
ClpP-HindIII-F GATCAAAGCTTCTTACGATATTTATTCCCGTCTGTTG
ClpP-HindIII-R GATCAAAGCTTTTGTTCAATAACTACTGGAATCAT
Asp1-J48-SalI-F GATCAGTCGACATGTTTTATTTTATTCCTTCG
Asp1-J48-BamHI-F GATCAGGATCCCCATGTTTTATTTTATTCCTTCG
Asp1-J48-BamHI-R GATCAGGATCCTTATTCTTTTTCTAATAATTTTCG
Asp1-Col-SalI-F GATCAGTCGACATGAAATACTTTATTCCAGC
Asp1-Col-BamHI-F GATCAGGATCCATGAAATACTTTATTCCAGC
Asp1-Col- XhoI -R GATCAGGATCCTTACGTGGCATCATTTTCACC
Asp3-J48-BamHI-F GATCAGGATCCCCATGATTTTGGGAGAGTGTTTAG
Asp3-J48-XhoI-R TGTGTCTCGAGTCATTTTTTATCCTTAGA
Asp3-J48-BamHI-R GATCAGGATCCTCATTTTTTATCCTTAGA
Asp3-J48-SalI-F GATCAGTCGACATGATTAAAAAAAGAATACAG
Asp3-J48-KpnI-R GATCA GGTACC TTTTTTATCCTTAGAAAATG
Asp3-Col-SalI-F GATCAGTCGACATGCTGAAAAACAAAACATTTAAAG
Asp3-Col-KpnI-R GATCAGGTACCTGTTTCATTTACTTCCCC
Gap1-EcoRI-F(Δ1–10) GATCAGAATTCGGTCAGAGGCAATGGTATTATG
Gap1-SalI-F(Δ1–10) GATCAGTCGACATGGGTCAGAGGCAATGGTATTATG
Gap1-HindIII-F(Δ11–20) GATCAAAGCTTTGGTGGTTTCGAGTAAAAAATAG
Gap1-HindIII-R(Δ11–20) GATCAAAGCTTATTATACCAAGAAGGTAC
Gap1-HindIII-F(Δ21–28) GATCAAAGCTTATGACCTTTGATGATAGTG
Gap1-HindIII-R(Δ21–28) GATCAAAGCTTCGGCGTATCATAATACCATTG
Gap1-HindIII-F(Δ29–45) GATCAAAGCTTGAAGAAATAGGATTAATG
Gap1-HindIII-R(Δ29–45) GATCAAAGCTTTCTATTTTTTACTCGAAAC
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Production of Gap1 and Gap3 in gap1 and gap3 mutants

To examine production of Gap1 and Gap3 in the gap1 and gap3 mutants, 1 ml of mutant cells was grown to exponential growth phase (OD470 = 0.6), harvested by centrifugation and lysed in NETN buffer with LambdaSa2 lysin (Li et al., 2008). The cell lysates were subjected to Western blot analysis using Gap1 and Gap3 polyclonal antibody. Gap1 and Gap3 polyclonal antibodies were produced by immunizing rabbits with the recombinant proteins (Cocalico). Antisera against Gap1 and Gap3 were affinity purified using the highly purified recombinant proteins as described (Chen et al., 2006). Production of FimA in S. parasanguinis was determined by anti-FimA antibody(Oligino & Fives-Taylor, 1993) and used as a loading control.

RT-PCR to evaluate gene expression

To examine expression of gap3 by the gap1 mutant, total RNA was extracted from S. parasanguinis wild type and the gap1 mutant strains using TRIzol® Reagent (Invitrogen). RQ1 DNase (Promega) treated RNA samples were used for RT-PCR. Briefly, 10 µg of RNA were mixed with 1 µl of 5 µM gap1 reverse primer (Gap1-KpnI-R), denatured at 70°C for 5 min, and quickly chilled on ice. Four µl of M-MLV 5X Reverse Transcriptase Buffer (Promega), 1 µl of 10 mM dNTP, 0.5 µl of 40 U RNasin® (Promega) and 1 µl of 200 U M-MLV reverse transcriptase (Promega) were added to the above RNA-primer mixture to a total volume of 20 µl, which was incubated at 42°C for 1 h and heated at 94°C for 5 min to terminate the reaction. cDNA from the reverse-transcribed products was subsequently used as a template for PCR amplification using a Gap1 primer set, Gap1-SalI-F1 and Gap1-KpnI-R. Expression of fimA was also detected by RT-PCR with FimA-1F and FimA-930, and served as a positive control.

Construction of recombinant strains to evaluate protein stability

Plasmid pVPT-gap3-CHSV and pVPT-gap1/3-CHSV were constructed to evaluate Gap3 stability in the presence or absence of Gap1 in recombinant E. coli and S. parasanguinis strains. A full-length gap3 gene was amplified from genomic DNA of S. parasanguinis FW213 by PCR with the Gap3-SalI-F1/Gap3-KpnI-R primer set. Full-length gap1 and gap3 was amplified from the genomic DNA of a gap2 mutant (Wu et al., unpublished) with the Gap1-SalI-F1/Gap3-KpnI-R primer set. The amplified PCR fragments were digested by SalI and KpnI and ligated into pVPT-CHSV to generate pVPT-gap3-CHSV and pVPT-gap1/3-CHSV, respectively.

Plasmid pVPT-asp3-CHSV (S. agalactiae J48 or S. aureus COL) and pGEX-asp1 (J48 or COL) were constructed to examine the stability of Gap3 homologues from S. agalactiae J48 and S. aureus COL in the presence or absence of corresponding Asp1 homologues of Gap1, respectively. Full-length asp3 (gap3 homologues from J48 and COL) was amplified from genomic DNA of S. agalactiae J48 and S. aureus COL by PCR with primer sets Asp3-J48/COL-SalI-F and Asp3-J48/COL-KpnI-R, respectively. The amplified PCR fragments were digested by SalI and KpnI, and ligated into pVPT-CHSV to yield pVPT-asp3-CHSV (J48 or COL). Full-length asp1 from J48 and COL was amplified from their respective genomic DNA by PCR with primer sets Asp1-J48-BamHI-F/Asp1-J48-BamHI-R and Asp1-COL-BamHI-F/ Asp1-COL-XhoI-R, respectively. The PCR fragments were digested by BamHI for J48 asp1 and by BamHI and XhoI for COL asp1, and ligated into pGEX-5X-1 to yield pGEX-asp1 (J48 or COL), respectively.

Protein stability assays

Recombinant E. coli strains carrying pVPT-gap3-CHSV and pVPT-gap1/3-CHSV plasmids or pVPT-asp3-CHSV (J48/COL) and pGEX-asp1 (J48/COL) plasmids were used to evaluate the stability of Gap3 or Asp3 by a well-established stability assay (Aldridge et al., 2003). The recombinant E. coli strains were grown to exponential growth phase (OD600 = 0.6) and treated with chloramphenicol (200 µg/ml) to stop nascent protein synthesis and then monitor protein stability. One ml of chloramphenicol-treated cells was harvested at different time points, mixed with SDS sample buffer and subjected to Western blot analysis. The stability of Gap3-HSV fusion protein in recombinant E. coli was evaluated using anti-Gap3 polyclonal antibody and anti-HSV monoclonal antibody (Novagen). The stability of Asp3-HSV fusion protein in recombinant E. coli was evaluated using anti-HSV monoclonal antibody. For Gap3 stability in S. parasanguinis, pVPT-gap3-CHSV and pVPT-gap1/3-CHSV constructs were each transformed into a gap1 mutant. The transformed S. parasanguinis cells were grown to OD470 = 0.6 and treated with chloramphenicol at 250 µg/ml to stop protein synthesis. One ml of chloramphenicol-treated cells was harvested 0–60 min after the treatment and lysed in NETN buffer with LambdaSa2 lysin. The cell lysates were subjected to Western blot analysis using anti-Gap3, anti-HSV and anti-FimA antibody, respectively.

Construction of the gap1 deletion mutants

A series of gap1 deletions in pGEX-5X-1 and E. coli-streptococcus shuttle vector pVT1666 (Chen et al., 2006) were constructed as follows. The DNA fragment corresponding to the full-length gap1 with a 10-amino acid deletion (1–10aa) at the N terminus was amplified from genomic DNA of FW213 using primer sets Gap1-EcoRI-F(Δ1–10)/Gap1-XhoI-R and Gap1-SalI-F(Δ1–10)/ Gap1-KpnI-R, respectively. The PCR products were digested by EcoRI/XhoI and SalI/KpnI, respectively, and ligated into pGEX-5X-1 and pVT1666 to generate pGEX-Gap1Δ1–10 and pVPT-Gap1Δ1–10, respectively. Other constructs were generated in a similar fashion. In brief, PCR fragments were amplified from pGEX-Gap1 with the following primer sets, Gap1-HindIII-F/R(Δ11–20), Gap1-HindIII-F/R(Δ21–28), Gap1-HindIII-F/R(Δ29–45) respectively. The resulting PCR products were digested with HindIII and self-ligated to generate six gap1 deletion constructs in pGEX. These gap1 deletion constructs were PCR amplified using the Gap1-SalI-F1/Gap1-KpnI-R primer set. The resulting PCR fragments were digested with SalI and KpnI, and ligated into pVT1666 to obtain corresponding gap1 deletion constructs in pVT1666.

GST pull-down assays to analyze protein-protein interactions

GST pull-down assays described previously (Li et al., 2008) were used to determine protein-protein interactions between Gap1 and Gap3, and their homologues.

To examine the in vitro interaction between Gap1 variants and Gap3, pGEX-5X-1 and pGEX-Gap1/Gap1Δ1–10/Gap1Δ11–20/ Gap1Δ21–28/Gap1Δ29–45 were used to express GST, GST-Gap1 and GST-Gap1 mutant fusion proteins, and pBD-Gap3 was used to produce c-Myc tagged Gap3 in the in vitro translation system. The GST fusion proteins were purified by glutathione Sepharose 4B beads (Amersham). The in vitro translated products with c-Myc tag were synthesized using the TNT® Quick Coupled Transcription/Translation System (Promega). Five micrograms of glutathione Sepharose-bound GST fusion proteins were mixed with 5 µl of the c-Myc-tagged proteins in NETN buffer (20 mM Tris-HCl, pH7.2, 100 mM NaCl, 1 mM EDTA, 0.2% NP40) and incubated overnight on a rotary shaker at 4°C. The beads were washed 3 times with 600 µl NETN buffer, the proteins were eluted with SDS-PAGE sample buffer, and subjected to Western blot analysis with anti-c-Myc antibody.

Interactions between Asp1 and Asp3 from S. agalactiae J48 and S. aureus COL was investigated as follows. pGEX-Asp1 (J48/COL) and pGEX-Asp3 (J48) were used, respectively, to express GST-Asp1 (J48/COL) and GST-Asp3 (J48) fusion proteins. pBD-Asp1 (J48) and pBD-Asp3 (J48) were constructed and used to generate in vitro translated Asp1 (J48) and Asp3 (J48) proteins, and pVPT-Asp3-CHSV (COL) was used to express HSV-tagged Asp3. The asp3 gene was amplified from S. agalactiae J48 by PCR using the Asp3- J48-BamHI-F/Asp3- J48-XhoI-R primer set. The PCR products were digested by BamHI and XhoI, and ligated to pGEX-5X-1 to construct pGEX- Asp3 (J48). To construct plasmids for in vitro translation, asp1 and asp3 genes were amplified from S. agalactiae J48 by PCR using primer sets Asp1- J48-BamHI-F/Asp1- J48-BamHI-R and Asp3- J48-BamHI-F/Asp3- J48-BamHI-R, respectively. The PCR products were digested with BamHI and then ligated to pGBKT7 to create pBD-Asp1 (J48) and pBD-Asp3 (J48).

To investigate the interaction between Asp1 (COL) and Asp3 (COL), pGEX-Asp1 (COL) and pVPT-Asp3-CHSV (COL) were co-transformed into E. coli Top10. Proteins purified using glutathione Sepharose 4B beads from the constructed recombinant E. coli strain were subjected to Western blot analysis with anti-HSV monoclonal antibody. A recombinant E. coli strain carrying both pGEX-5X-1 and pVPT-Asp3-CHSV (COL) was used as a GST control for the pull-down assays.

Complementation of a gap1 mutant by different gap1 variants

Gap1Δ1–10, Gap1Δ11–20, Gap1Δ21–28, Gap1Δ29–45 and the positive control pVPT-gap1 were each transformed into a gap1 mutant via electroporation. Transformants were selected on TH agar plates containing kanamycin and erythromycin. The ability of the different Gap1 deletion variants to alter production of mature Fap1 was examined by Western blotting analysis using mature Fap1-specific antibody F51.

Expression and purification of His-SUMO-tagged fusion proteins

Three fusion plasmids were constructed to express His-SUMO (small ubiquitin-related modifier) -tagged Gap1Δ21–28, Gap3 and Gap1/3 complex, respectively. In brief, the full-length gap1 with a deletion of DNA fragment coding for amino acid residues 21–28 was amplified from pGEX-Gap1Δ21–28 using the Gap1-SalI-F2/ Gap1-XhoI-R primer set digested with SalI and XhoI, and then ligated into pET-SUMO to generate pET28-SUMO-Gap1Δ21–28. The full-length gap3 sequence was amplified from the genomic FW213 DNA using the Gap3-SalI-F2/Gap3-XhoI-R primer set, digested with SalI and XhoI, and cloned into pET-SUMO to produce pET28-SUMO-Gap3. The full-length gap1 and gap3 was amplified from genomic DNA of the gap2 mutant using the Gap1-NotI-1F/Gap3-XhoI-R primer set, digested by NotI and XhoI, and ligated into pET-SUMO to construct pET28-SUMO-Gap1/3. The constructed plasmids were verified by DNA sequence analysis and then transformed into E. coli BL21 (DE3).

Production of 6xHis-SUMO-tagged fusion proteins was induced with 0.1 mM isopropyl β-D-thiogalactoside (IPTG) at 18°C overnight, and the induced proteins were purified by Ni-NTA resin (Novagen). The 6xHis-SUMO tag was removed from the fusion proteins with SUMO protease (Peroutka Iii et al., 2011) at 4°C overnight. The Gap1Δ21–28, Gap3 and Gap1/3 complex were further purified by Ni-NTA resin. The concentration of the purified proteins was determined by the Bradford method (Bio-Rad Protein Assay). The highly purified recombinant Gap1Δ21–28 protein was used to immunize in mice to produce monoclonal antibody against Gap1 using a standard protocol (Elder & Fives-Taylor, 1986).

Mass spectrometry

The purified 6xHis-SUMO-Gap3 fusion protein was eluted by cleavage of the tag with SUMO protease, subjected to 12% SDS-PAGE analysis, and stained with Coomassie brilliant blue. A stained protein band was excised from the gel and digested with trypsin. The digested peptide fragments were analyzed by liquid chromatography-tandem mass spectrometry (LC/MS/MS) as described previously for protein identification ((Wu et al., 2010)).

Construction of a clpP mutant and a gap1/clpP double mutant

A clpP mutant and a gap1/clpP double mutant were constructed by insertional mutagenesis with a nonpolar kanamycin resistance cassette (aphA3) as described previously (Wu et al., 1998). A full-length clpP gene and its upstream 730bp flanking region were amplified from genomic DNA of S. parasanguinis FW213 by PCR with a primer set of ClpP-F and ClpP-R. The amplified 1.3kb fragments were cloned into pGEM-T Easy vector to yield pGEM-ClpP. A 28bp clpP internal fragment was deleted from pGEM-ClpP by inverse PCR with a primer pair of ClpP-HindIII-F and ClpP-HindIII-R. The inverse PCR product was digested with HindIII and then ligated in-frame with an 830bp nonpolar kanamycin resistance cassette (aphA3) to construct pGEM::ΔclpP-aphA3. The correct constructs were confirmed by PCR and DNA sequencing, and then transformed into S. parasanguinis FW213 and VT324, a gap1 mutant (Gap1Δ513–525)(Zhou et al., 2008b) by electroporation to produce a clpP mutant and a gap1/clpP double mutant (VT324/clpP−), respectively. The resulting mutants were selected by their ability to resist to kanamycin on TH agar plates and further verified by PCR and sequencing analysis. The confirmed mutants were used in this study.

In vitro proteolytic assays

Recombinant Gap1, Gap3 and the Gap1/3 complex were purified and used for proteolytic assays. Briefly, the purified 6xHis-SUMO fusion proteins, Gap1, Gap3 and Gap1/3 complex were treated with SUMO protease to remove the His-SUMO tag, and then digested with protease Endoproteinase Glu-C or Trypsin from Proti-Ace Kit (Hampton research) according to the manufacturer’s protocols. The proteolytic reaction was stopped at various time points (0–60 min) by the addition of Halt Protease Inhibitor Cocktail (Thermo Fisher Scientific). These samples were then subjected to SDS-PAGE analysis and stained with Coomassie brilliant blue.

Acknowledgments

This study was supported by R01DE017954 (H. Wu) from the National Institute of Dental and Craniofacial Research, by grants 30900034 (M. Zhou) and 30970060 (Yirong Li) from the National Natural Science Foundation of China. We thank Dr. Jeannine Brady from University of Florida for providing us with the clinical isolate S. agalactiae J48. We also thank Dr. Mary Ann Accavitti-Loper from UAB for providing help with the production of the Gap1 mAb.

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