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Journal of Bacteriology, February 2008, p. 1256-1266, Vol. 190, No. 4
0021-9193/08/$08.00+0     doi:10.1128/JB.01078-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Interaction between Two Putative Glycosyltransferases Is Required for Glycosylation of a Serine-Rich Streptococcal Adhesin{triangledown}

Su Bu,1 Yirong Li,1,{dagger} Meixian Zhou,1 Parastoo Azadin,3 Meiqin Zeng,1 Paula Fives-Taylor,4 and Hui Wu1,2*

Departments of Pediatric Dentistry,1 Microbiology, Schools of Dentistry and Medicine, University of Alabama at Birmingham, Birmingham, Alabama 35294,2 Complex Carbohydrate Research Center, University of Georgia at Athens, Athens, Georgia 30602,3 Department of Microbiology & Molecular Genetics, University of Vermont, Burlington, Vermont 054054

Received 8 July 2007/ Accepted 3 December 2007


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ABSTRACT
 
Fap1, a serine-rich glycoprotein, is essential for fimbrial biogenesis and biofilm formation of Streptococcus parasanguinis (formerly S. parasanguis). Fap1-like proteins are conserved in many streptococci and staphylococci and have been implicated in bacterial virulence. Fap1 contains two serine-rich repeat regions that are modified by O-linked glycosylation. A seven-gene cluster has been identified, and this cluster is implicated in Fap1 biogenesis. In this study, we investigated the initial step of Fap1 glycosylation by using a recombinant Fap1 as a model. This recombinant molecule has the same monosaccharide composition profile as the native Fap1 protein. Glycosyl linkage analyses indicated that N-acetylglucosamine (GlcNAc) is among the first group of sugar residues transferred to the Fap1 peptide. Two putative glycosyltransferases, Gtf1 and Gtf2, were essential for the glycosylation of Fap1 with GlcNAc-containing oligosaccharide(s) in both S. parasanguinis as well as in the Fap1 glycosylation system in Escherichia coli. Yeast two-hybrid analysis as well as in vitro and in vivo glutathione S-transferase pull-down assays demonstrated the two putative glycosyltransferases interacted with each other. The interaction domain was mapped to an N-terminal region of Gtf1 that was required for the Fap1 glycosylation. The data in this study suggested that the formation of the Gtf1 and Gtf2 complex was required for the initiation of the Fap1 glycosylation and that the N-terminal region of Gtf1 was necessary.


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INTRODUCTION
 
The Streptococcus sanguinis (formerly S. sanguis) streptococci are predominant microbes in the oral cavity. They colonize saliva-coated teeth and other oral surfaces and play an important role in the development of dental plaque (2, 12, 18). Adhesion of Streptococcus parasanguinis (formerly S. parasanguis) FW213 in an in vitro tooth model is mediated by fimbriae (9, 11). S. parasanguinis fimbriae are composed of multiple subunits of a serine-rich glycoprotein, Fap1 (35-37). Fap1 belongs to a growing family of serine-rich proteins. This family of proteins include GspB and Hsa of Streptococcus gordonii M99 (1) and Challis (29), SraP of Staphylococcus aureus (25), SrpA of Streptococcus sanguinis (21), PsrP of Streptococcus pneumoniae (20), and Srr-2 of Streptococcus agalactiae (23). Hsa, SraP, PsrP, and Srr-2 have been implicated in bacterial pathogenesis (20, 23, 25, 30). Fap1 is a large molecule comprised of 2,572 amino acid residues. It possesses two serine-rich repeat regions (RI and RII) and an acidic amino acid-rich, nonrepeat region. The RI region contains 28 copies of the dipeptide motif, whereas the RII region consists of 1,000 copies of the same dipeptide motif. A cell wall anchor domain at the carboxyl terminus is essential for Fap1 surface presentation, fimbrial biogenesis, and bacterial adhesion (35). Fap1 is heavily glycosylated with N-acetylglucosamine (GlcNAc) and glucose residues within the serine-rich repeat regions (27). Glycosylation of Fap1 is important for the biofilm formation of Streptococcus parasanguinis (38).

A seven-gene cluster downstream of fap1 has been identified and implicated in Fap1 glycosylation. One glycosyltransferase, Gtf1, is critical for Fap1 glycosylation (34), whereas two accessory secretion components, SecA2 and SecY2, are required for the biogenesis and secretion of mature Fap1 (3, 34). The serine-rich repeat regions are modified by O-linked glycan moieties (27). The compositions of the monosaccharides on Fap1 are known (27), but the order of transfer and the identity of the initial monosaccharide residue(s) transferred to Fap1 are not known. Other genes that are involved in the transfer and how they coordinate to initiate Fap1 glycosylation have not been investigated. O-linked glycan moieties are synthesized by glycosyltransferases in a stepwise manner (28). Eukaryotic glycosyltransferases and their related proteins interact with each other to form enzyme complexes in N-linked glycan biosynthetic pathways (6, 24). However, such interactions have not been identified in O-linked glycosylation systems (6). Both N- and O-linked glycosylation systems are found in a variety of bacterial species (28), including streptococci (8), and some bacterial and eukaryotic systems share a common transfer mechanism (33). However, whether bacteria have adopted protein-protein interaction schemes similar to eukaryotic cells to control protein glycosylation is unknown.

To address the above questions, we established an in vivo glycosylation model in Escherichia coli and studied the functional contribution of two putative glycosyltransferases, Gtf1 and Gtf2, to Fap1 glycosylation. Our experimental evidence suggested that GlcNAc was among the first group of sugar residues transferred to the Fap1 polypeptide, that Gtf1 interacted with Gtf2, and that this interaction was required for the transfer.


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MATERIALS AND METHODS
 
Bacterial strains and growth conditions. All bacterial and yeast (Saccharomyces cerevisiae) strains and plasmids used in this study are listed in Tables 1 and 2. S. parasanguinis parent strain FW213 and its derivatives were cultured using Todd-Hewitt broth (THB) with the appropriate antibiotics as described previously (38).


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  		TABLE 1. Strains used in this study


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  		TABLE 2. Plasmids used in this study

DNA manipulation and sequence analysis. Standard methods (22) were used for routine DNA manipulation. DNA-modifying enzymes were purchased from Promega (Madison, WI). Plasmid DNA was isolated using Qiaprep Spin Miniprep kit (Qiagen, Inc., Santa Clarita, CA). Genomic DNA from S. parasanguinis strains was isolated using the PureGene genomic DNA purification kit (Gentra Systems, Minneapolis, MN). The PCR primers used are listed in Table 3. Nucleotide sequences of amplified PCR products were determined by the dideoxy chain termination method with an ABI 1372A DNA sequencer (26). Homology comparison was carried out using ClustalW software (http://www.ebi.ac.uk/Tools/clustalw/).


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  		TABLE 3. Primers used in this study

Construction of gtf2 single mutant and gtf1 gtf2 double mutant in S. parasanguinis. The gtf2 gene with 500-bp flanking DNA sequence was PCR amplified from genomic DNA of S. parasanguinis using primers gtf2-5' and gtf2-3'. The PCR fragment was cloned into pGEM-T Easy vector to generate pAL60. An inverse-PCR strategy (19) was used to introduce a PstI site into pAL60. In brief, primers with embedded PstI sites, gtf2-PstIF and gtf2-PstIR, were used to inverse PCR amplify a 4.20-kb fragment from pAL60. The amplified fragment was digested with PstI and self-ligated to generate pAL61, in which the gtf2 gene was deleted. pAL61 was then digested with PstI and ligated with a nonpolar kanamycin resistance cassette, aphA3 (a gift of Allen Honeyman) (14), to form pAL62, in which gtf2 was interrupted by insertion of aphA3. The mutated gtf2 allele was excised from pAL62 by EcoRI digestion and then ligated into S. parasanguinis suicide vector pSF143 (32) to create pAL63.

A plasmid used to generate the gtf1 gtf2 double mutant was constructed in a similar manner. The gtf1 gtf2 fragment with 600-bp flanking sequences was amplified using the primer pair Gtf-Spara-FF and Orf4D2-Spara-RR and then cloned into pGEM-T Easy to generate plasmid pAL70. The majority of gtf1 and gtf2 open reading frame (ORF) sequences were deleted from pAL70 by inverse-PCR amplification with primer pairs gtf1-2-HindIII-IVS-RR and gtf1-2-HindIII-IVS-FF. The amplified fragment was digested with HindIII and self-ligated to generate pAL71. pAL71 was then digested with HindIII and ligated with the nonpolar kanamycin resistance cassette aphA3 to form pAL72, in which the gtf1 gtf2 fragment was replaced by aphA3. The allele replaced by aphA3 was excised from pAL72 by EcoRI digestion and then ligated into S. parasanguinis suicide vector pSF143 (32) to create pAL73. These plasmids, pAL63 and pAL73, were transformed into S. parasanguinis FW213 as described previously (10) to generate kanamycin-resistant and tetracycline-sensitive allelic replacement mutants. Insertional inactivation of gtf2 or gtf1 and gtf2 by a double-crossover recombination event was confirmed by PCR and Southern blot hybridization. Confirmed gtf2 and gtf1 gtf2 mutants AL63 and AL74 were used for further studies.

Complementation of gtf1, gtf2, and gtf1 gtf2 mutants. The ORFs of gtf1, gtf2, and the gtf1 gtf2 mutant were amplified from genomic DNA of S. parasanguinis with primer pairs SalI-gtf1-FF and BamHI-gtf1-RR, SalI-gtf2-FF and KpnI-gtf1-2-RR, and SalI-T7-gtf1-2-FF2 and KpnI-T7-gtf1-2-RR, respectively (Table 3). The PCR products were digested with KpnI and SalI or BamHI and SalI and then ligated into E. coli-Streptococcus shuttle vector GFP-pVPT (4). GFP-pVPT has a functional streptococcal promoter, Pmal. The desired plasmids were isolated from putative transformants and confirmed by restriction enzyme digestion and DNA sequencing. The control plasmid, GFP-pVPT, and the resulting plasmids, gtf1-pVPT, gtf2-GFP-pVPT, and T7-gtf1-2-GFP-pVPT, were transformed into the respective mutants. The ability of cloned gtf1, gtf2, and the gtf1 gtf2 fragment to restore the mature Fap1 expression in their corresponding mutants was examined by BactELISA assays (34) using monoclonal antibody (MAb) F51 (27). The positive transformants were further confirmed by Western blot analyses with peptide-specific MAb E42 and glycan-specific MAb F51 (27).

Construction of Fap1{Delta}RII in S. parasanguinis. pVT1175 (37) contains the full-length fap1 gene and an E. coli origin of replication. To construct Fap1{Delta}RII, An inverse-PCR strategy was used to delete the repeat region II from pVT1175 with fap1-1814KpnI and fap1-7080KpnI primers. The inverse-PCR product was digested with KpnI and self-ligated to generate plasmid pAL80. The resultant pAL80 was digested by HindIII and ligated with a HindIII fragment of pVA838 that contains a streptococcal origin of replication and an erythromycin resistance cassette to generate pAL81 (fap1{Delta}RII-pVA838-HindIII). pAL81 was transformed into the fap1 mutant VT1393 to generate AL81.

Expression, purification, and characterization of Fap1{Delta}RII. Culture supernatants of AL81 were collected by centrifugation and precipitated with 60% (NH4)2SO4. The precipitated proteins were subjected to buffer exchange against phosphate-buffered saline with Centricon YM-50 (Millipore), the concentrated samples were further purified with a MAb E42 immunoaffinity column which was packed with disuccinimidyl suberate (DSS)-cross-linked MAb E42 and protein G. The purification procedure was performed according to the instructions of the Seize X protein G immunoprecipitation kit (Pierce, Rockford, IL). The purity of the Fap1{Delta}RII was assessed by 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis and Coomassie brilliant blue staining.

Monosaccharide composition and glycosyl linkage analyses of Fap1{Delta}RII. The monosaccharide composition analysis procedure was carried out as previously described (15). In brief, methyl glycosides were prepared from 100 µg of the dried Fap1{Delta}II by methanolysis in 1 M HCl in methanol at 80°C for 18 h, followed by re-N-acetylation with pyridine and acetic anhydride in methanol (for detection of amino sugars). The samples were per-O-trimethylsilylated by treatment with Tri-Sil at 80°C for 20 min. Gas chromatography-mass spectrometry (GC-MS) analysis was performed on an HP5890 gas chromatograph interfaced to a 5970 MSD using a Supelco DB-1 fused-silica capillary column (30 m by 0.25-mm inside diameter).

Glycosyl linkage analysis of Fap1{Delta}II was performed with matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) MS. Purified Fap1{Delta}RII was subjected to β-elimination with 1 M NaBH4 in 50 mM NaOH to release O-linked glycan(s). The released glycans were permethylated by the method of Ciukanu and Kerk (7) and extracted with methylene chloride-water to remove any impurities, the resulting permethylated samples were suspended in methanol and mixed with MALDI matrix 2,5-dihydroxyl benzoic acid. MALDI-TOF MS was performed with a Voyager mass spectrometer operated in the positive-ion mode. Profiling of constituent glycans was monitored.

Coexpression of Fap1{Delta}RII with two glycosyltransferases and their derivatives. The gtf1 gtf2 fragment was amplified with BamHI- and XhoI-embedded primers listed in Table 3. The PCR product was digested and then ligated into pGEX6p1 to allow the in-frame fusion of glutathione S-transferase (GST) with gtf1 to generate pAL200. Two negative control plasmids were also constructed. One lacks gtf1, and the other lacks gtf2. To delete the majority of gtf1's ORF (amino acids [aa] 21 to ~485), we performed inverse PCR using pAL200 as a template and gtf1-KpnI-IVS-RR and gtf1-KpnI-IVS-FF as a primer pair. The inverse-PCR product was digested with KpnI and self-ligated to construct pAL201. A similar strategy was used to delete aa 15 to ~417 from gtf2's ORF in pAL200 using the primer pair gtf2-KpnI-IVS-RR and gtf2-KpnI-IVS-FF to construct pAL202. Substrate plasmid pAL81 (chloramphenicol resistant) was cotransformed with pGEX6p1; two negative control plasmids, pAL201 and pAL202; or a positive control plasmid, pAL200 (ampicillin resistant). The chloramphenicol- and ampicillin-resistant cotransformants were further analyzed by the presence of two distinct plasmids. The strains harboring correct plasmids were chosen for the subsequent experiments.

Analysis of Fap1{Delta}RII glycosylation by lectin blotting assays. Culture supernatants of S. parasanguinis were prepared as described previously (34). Cell pellets from 0.5-ml exponential-phase-grown E. coli cultures were harvested and subjected to electrophoresis on a 4 to 12% gradient gel. Separated proteins were transferred to nitrocellulose membranes and incubated with blocking buffer, which contains 1x Tris-buffered saline (TBS), 0.1% Tween 20, 1 mM CaCl2, 1 mM MnCl2, 1 mM MgCl2 and 2% polyvinyl alcohol (Sigma) for 1 h and then subjected to lectin blot analyses with horseradish peroxidase-conjugated succinyl wheat germ agglutinin (sWGA) or other conjugated lectins.

Yeast two-hybrid analysis. The procedure for yeast two-hybrid analysis was carried out according to the protocol of the MATCHMAKER system 3 (Clontech). DNA binding domain (BD) and activation domain (AD) fusion vectors, pGBKT7 (pBD) and pGADT7 (pAD), are designed for the identification and confirmation of protein interactions. Bait and prey proteins are expressed as GAL4 fusions with c-Myc and hemagglutinin epitope tags in these vectors, respectively. We first fused gtf1 and gtf2 with the DNA BD of pBD and DNA AD of pAD, respectively, using primer pairs listed in Table 3. The plasmid set pAD-gtf1 and pBD-gtf2 or plasmid set pAD-gtf2 and pBD-gtf1 was cotransformed into yeast strain AH109. Cotransformed yeast cells were streaked on a Leu/Trp/His/Ade amino acid-deficient SD agar plate to score protein-protein interactions.

In vitro GST pull-down assays. The in vitro GST pull-down assay can determine direct protein-protein interaction in vitro. We first constructed GST-Gtf1 and GST-Gtf2 fusion plasmids. pBD-gtf1 or -gtf2 was digested with BamHI and XhoI to release gtf1 and gtf2 ORFs, which were subcloned into pGEX5p1 vector (Amersham) to generate Gtf1- and Gtf2-GST in-frame fusion proteins, respectively. The GST fusion proteins were purified using glutathione Sepharose 4B beads (Amersham) following the manufacturer's instructions. In brief, 0.1 mM isopropyl-β-D-thiogalactopyranoside (IPTG)-induced cell pellets were lysed in the presence of 1 mM dithiothreitol and 1 mg/ml lysozyme by sonication. The cell lysates free of cell debris were mixed with 200 µl NETN buffer (20 mM Tris-HCl [pH 7.2], 0.1 M NaCl, 1 mM EDTA, 0.5% NP-40, and protease inhibitor cocktail [1:20, vol/vol])-equilibrated GST beads. The mixture was incubated at 4°C overnight. The beads bound with GST or GST fusion proteins were washed with NETN buffer five times. Concentrations of GST fusion proteins were analyzed by SDS-PAGE followed by Coomassie blue staining.

The c-Myc-tagged in vitro-translated protein products of the full-length gtf1 and gtf2 genes were generated using TnT Quick Coupled transcription/translation system (Promega). One microgram of supercoiled DNA from pGBK7, pGBK7-Gtf1, and pGBK7-Gtf2 was used in the reaction mixtures that have 40 mM methionine. Approximately 5 µg of GST or GST fusion proteins in NETN washing buffer was mixed with 5 µl of in vitro-translated c-Myc fusion proteins in a final volume of 200 µl NETN binding buffer and incubated on a rotary shaker at 4°C overnight. The beads were washed three times with 600 µl NETN washing buffer, and the bound proteins were eluted by boiling in the SDS sample loading buffer and subjected to Western blotting analysis with anti-c-Myc antibody.

Construction of transposon insertion mutants in the gtf1 fragment in E. coli. Plasmid gtf1-2-pGEX6p1 was used as a template to generate the construct. A T7 tag was fused into 3' end of gtf2 for the planned interaction study. There is an internal restriction enzyme site, BglII, near the 3' end of gtf2. Using the primer pair BglII-gtf2-FF and XhoI-gtf2-T7-RR, a partial gtf2 fragment with a T7 tag was amplified from pAL200. The amplified fragment was digested with BglII and XhoI and then ligated into pAL200 to generate pAL205, which has a T7 tag fused at the 3' end of gtf2. A similar strategy was used to generate a negative control plasmid, pAL206, with pAL201 as a template.

Construction of Tn5 transposon insertion mutants in the gtf1 fragment in E. coli was carried out using the EZ-Tn5 in-frame linker insertion kit (Epicentre). The kit is designed to rapidly produce random 57-bp (19 amino acids) insertions into a cloned DNA. Since the inserted 19 codons are designed to be readable in all three reading frames, the transposition generates nonpolar mutants.

As a NotI restriction enzyme digestion step is required to remove the kanamycin-resistant cassette from the insertional mutant constructs after transposition and there is a NotI site in the pGEX6p1 plasmid sequence, we first blunted the NotI site of the template plasmid pAL205 (gtf1-2-T7-pGEX6p1) by ligation of NotI-digested and T4 DNA polymerase-treated pAL205. The modified pAL205 was used to construct the transposon insertion mutants. The transposon mutants with Tn5-Km insertions were selected by their ability to resist to both ampicillin and kanamycin antibiotics. The insertion sites were identified by colony PCR using transposon- and Gtf1-specific primers NotI/KAN-3 RP-2 and pGEX6p1gtf1-F(910) (Table 3). The kanamycin-resistant cassette was removed by NotI digestion of plasmids prepared from selected transposon mutants. The exact transposition site was determined by sequencing. Four transposon insertion plasmids, pAL207, pAL208, pAL209, and pAL300 (Table 4), were chosen and used for cotransformation with pAL81 to generate AL408, -409, -410, and -411.


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  		TABLE 4. Characterization of Tn5 transposon insertion mutants in the gtf1 gene

LC/MS/MS spectrometric analyses. Tandem-MS (MS/MS) analyses were performed with a Q-Tof2 mass spectrometer (Micromass, Manchester, United Kingdom) using electrospray ionization. Excised proteins that had undergone a 16-h tryptic digestion at 37°C were purified using ZipTips to concentrate and desalt the samples. The samples were then analyzed by liquid chromatography-MS/MS (LC/MS/MS). LC was performed using an LC Packings Ultimate LC Switchos microcolumn switching unit and Famos autosampler (LC Packings, San Francisco, CA). The samples were concentrated on a 300-µm-inside diameter C18 precolumn at a flow rate of 10 µl/min with 0.1% formic acid and then flushed onto a 75-µm-inside diameter C18 column at 200 µl/min with a gradient of 5 to 100% acetonitrile (0.1% formic acid) in 30 min. The nano-1c interface was used to transfer the 1c eluent into the mass spectrometer. The Q-Tof2 mass spectrometer was operated in the automatic switching mode, whereby multiply charged ions were subjected to MS/MS if their intensities rose above 6 counts. The tandem mass spectra were processed with the MassLynx MaxEnt 3 software.


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RESULTS
 
Gtf1 and Gtf2 mutants exhibit a similar phenotype. Gtf1 is required for Fap1 glycosylation (34). Gtf2 shares some homology with other glycosyltransferases; however, its function in Fap1 glycosylation and its relationship with Gtf1 are not known. We inactivated the gtf2 gene and also constructed a double mutant of the gtf1 and gtf2 genes in S. parasanguinis and compared these two mutants with the previously characterized gtf1 mutant (34). As Fap1 and Fap1 precursors (32) have the same subcellular distribution pattern in S. parasanguinis (34), culture supernatant fractions were used to determine Fap1 expression. Wild-type S. parasanguinis FW213 expressed a mature 200-kDa Fap1 (Fig. 1A, lane 1). The gtf2 mutant and the double mutant (gtf1 gtf2) behaved like the gtf1 mutant and failed to express any mature 200-kDa protein. Instead, all of these mutants generated a high-molecular-mass Fap1 protein (360 kDa) that was detected by a peptide-specific MAb, E42 (Fig. 1A, lanes 3, 7, and 11). This high-molecular-mass Fap1 did not react with either of the glycan-specific MAbs F51 (Fig. 1B, lanes 3, 7, and 11) and D10 (data not shown), suggesting it has defects in Fap1 glycosylation.


Figure 1
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  		FIG. 1. Western blot analyses of Fap1 expression by gtf1, gtf2, and gtf1 gtf2 mutants and their derivatives. Culture supernatants were prepared from wild-type FW213 (lane 1), VT1393 (fap1 mutant; lane 2), VT1583 (gtf1 mutant; lane 3), AL51 (gtf1 mutant, pVPT; lane 4), AL52 (gtf1 mutant, pVPT-gtf2; lane 5), AL50 (gtf1 mutant, pVPT-gtf1; lane 6), AL63 (gtf2 mutant; lane 7), AL64 (gtf2 mutant, pVPT; lane 8), AL65 (gtf2 mutant, pVPT-gtf1; lane 9), AL66 (gtf2 mutant, pVPT-gtf2; lane 10), AL74 (gtf1 gtf2 mutant; lane 11), AL75 (gtf1 gtf2 mutant, pVPT-gtf1; lane 12), AL76 (gtf1 gtf2 mutant, pVPT-gtf2; lane 13), and AL77 (gtf1 gtf2 mutant, pVPT-gtf1 and -2; lane 14); separated by electrophoresis through 4 to 12% polyacrylamide gradient gels; and subjected to Western blot analyses with peptide-specific MAb E42 (A) and glycan-specific MAb F51 (B). The culture supernatants prepared from VT1393 (fap1 mutant; lane 1), wild-type FW213 (lane 2), VT1583 (gtf1 mutant; lane 3), and AL63 (gtf2 mutant; lane 4) were probed with lectin sWGA (C), carbohydrate staining (D), and MAb E42 (D), respectively. MW, molecular mass.

Furthermore, concurrent inactivation of both gtf1 and gtf2 did not result in a phenotype (Fig. 1A and B, lanes 11) that was different from that produced by the single-insertion mutants (Fig. 1A and B, lanes 3 and 7). Introduction of plasmid-borne gtf1, gtf2, or a gtf1 gtf2 fragment into their corresponding mutants restored the expression of the 200-kDa mature Fap1 protein (Fig. 1A and B, lanes 6, 10, and 14). Introduction of an empty plasmid, GFP-pVPT, or plasmids harboring the other gene (gtf2-pVPT into the gtf1 mutant or gtf1-pVPT into the gtf2 mutant) failed to restore Fap1 glycosylation (Fig. 1A and B, lanes 4, 5, 8, 9, 12, and 13). These results demonstrated that both Gtf1 and Gtf2 are required for Fap1 glycosylation and they cannot cross-complement each other, suggesting the two putative glycosyltransferases function in concert.

To confirm the high-molecular-mass Fap1 was not glycosylated, wild-type FW213, and gtf1 and gtf2 mutants were probed with lectin sWGA and a general carbohydrate stain. Only mature 200-kDa Fap1 in wild-type FW213 reacted with sWGA as well as the general carbohydrate stain (Fig. 1C and D, lane 2). The high-molecular-mass Fap1 species in the gtf1 and gtf2 mutants did not react with either reagent (Fig. 1C and D, lanes 3 and 4). In contrast, it reacted with the peptide-specific MAb E42 as a 360-kDa Fap1 precursor (Fig. 1E, lanes 3 and 4), suggesting that the high-molecular-mass Fap1 was not glycosylated.

The recombinant Fap1{Delta}RII is modified with GlcNAc-containing oligosaccharide(s). Fap1 is encoded by an 8-kb ORF and has two serine-rich repeat regions. These two repeat regions differ in size; however, they are identical in their amino acid compositions (35). It is difficult to work with full-length Fap1. Therefore, we engineered and expressed a new recombinant plasmid, pAL80, in which the repeat region II was deleted (Fig. 2A). The truncated Fap1, designated Fap1{Delta}RII, was expressed in recombinant strain AL81 and detected by MAb E42 and lectin sWGA (Fig. 2B, lanes 1). As sWGA is specific for GlcNAc, these data suggest Fap1{Delta}RII is glycosylated by GlcNAc-containing oligosaccharides. Fap1{Delta}RII was purified to near homogeneity by affinity chromatography (Fig. 2C, lane 2). Monosaccharide composition analyses of this protein revealed the presence of GlcNAc, glucose, galactose, and rhamnose at a ratio of 60:21:4:6. These sugar residues are also found in mature Fap1 (27), and the ratio is similar to that of full-length Fap1, indicating Fap1{Delta}RII was a useful model to study Fap1 glycosylation. Glycosyl linkage analyses of the purified Fap1{Delta}RII (Fig. 2D) revealed that the recombinant Fap1 was modified by mixed oligosaccharides and all the oligosaccharides started with GlcNAc, suggesting that the initial sugar residues attached to the peptide contain GlcNAc.


Figure 2
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  		FIG. 2. Characterization of the recombinant Fap1 (Fap1{Delta}RII) that contains only repeat region I. (A) Diagram of Fap1 and Fap1{Delta}RII constructs. The fap1{Delta}RII construct was generated by deletion of the entire RII region from the intact fap1. (B) Fap1{Delta}RII is glycosylated by GlcNAc-containing sugar residues. The fap1{Delta}RII plasmid pAL81 was transformed into the fap1-null mutant VT1393 to generate a recombinant strain, AL81. AL81 (lane 1) and VT1393 (lane 2) were analyzed by Western blotting with MAb E42 and lectin sWGA (B). (C) Purification of Fap1{Delta}RII. Fap1{Delta}RII was purified by MAb E42 affinity chromatography from culture supernatants of AL81 and analyzed by Coomassie blue staining. (D) Glycosyl linkage analysis of Fap1{Delta}RII. The purified Fap1{Delta}RII protein was subjected to β-elimination, and the released oligosaccharides were analyzed by MALDI-TOF MS.

Both Gtf1 and Gtf2 were essential for Fap1{Delta}RII glycosylation in a functional E. coli glycosylation system. The above experiments indicated that Fap1{Delta}RII was glycosylated with GlcNAc-containing oligosaccharides. Gtf1 and Gtf2 were essential for Fap1 glycosylation in S. parasanguinis. Therefore, we hypothesized Gtf1 and Gtf2 were the glycosyltransferases to transfer the GlcNAc-containing oligosaccharides to the recombinant Fap1{Delta}RII. To test this hypothesis, we developed an in vivo glycosylation system in E. coli using Fap1{Delta}RII as the substrate and Gtf1 and Gtf2 as enzymes. Coexpression of the two plasmids in E. coli led to the production of a Fap1{Delta}RII doublet that migrated at the 75- and 83-kDa positions when probed with MAb E42 (Fig. 3A, lane 4). The GlcNAc-specific lectin sWGA reacted with the two same protein bands (Fig. 3B, lane 4). However, the bands did not react with any other lectins tested, including concanavalin A, Dolichos biflorus agglutinin, peanut agglutinin, Ricinus communis agglutinin I, soybean agglutinin, and Ulex europaeus agglutinin I. As these lectins are specific for glucose, mannose, galactose, N-acetylgalactosamine, and fucose, respectively, and did not react with Fap1{Delta}RII, we concluded Fap1{Delta}RII was only glycosylated by GlcNAc-containing oligosaccharides in E. coli. Deletion of either one or both Gtfs mitigated the Fap1 glycosylation, as determined by sWGA blotting (Fig. 3B, lanes 1 to 3), and the protein bands recognized by MAb E42 migrated to lower-molecular-mass positions (Fig. 3A, lanes 1 to 3). These results demonstrated that both Gtf1 and Gtf2 were required for Fap1{Delta}RII glycosylation with GlcNAc-containing oligosaccharides in the E. coli glycosylation system, supporting our early conclusion that both Gtf1 and Gtf2 were required for Fap1 glycosylation in S. parasanguinis.


Figure 3
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  		FIG. 3. Analyses of Fap1{Delta}RII expression in an E. coli glycosylation system. E. coli strains carrying pAL81 were cotransformed with plasmids pGEX6p1 (lane 1), pAL201 (lane 2), pAL202 (lane 3), and pAL200 (lane 4) and subjected to immunoblot analysis with MAb E42 (A) and GlcNAc-reactive lectin sWGA (B). MW, molecular mass.

Interaction between Gtf1 and Gtf2 revealed by a yeast two-hybrid analysis. As gtf1 and gtf2 mutants have very similar phenotypes and gtf1 and gtf2 cannot cross-complement each other, it is likely that they are interdependent in the Fap1 biosynthetic pathway. We hypothesized that Gtf1 and Gtf2 formed an enzyme complex to modulate Fap1 glycosylation. To test this, we used the yeast two-hybrid analysis to detect protein-protein interactions between Gtf1 and Gtf2. Numerous colonies were observed when the plasmid pair pAD-gtf1 and pBD-gtf2 or pAD-gtf2 and pBD-gtf1 were cotransformed. Their growth was comparable to that of the positive control (pBD-p53+ and pAD-T), suggesting Gtf1 interacts with Gtf2. The cotransformants of pAD-gtf1 and pBD, pBD-gtf1 and pAD, pAD-gtf2 and pBD, and pBD-gtf2 and pAD, along with the negative control (pBD-lam+ and pAD-T), failed to grow on the selective media (Fig. 4A), indicating that there was no self-activation of Gtf1 or Gtf2. Thus, we concluded that Gtf1 and Gtf2 specifically interacted with each other.


Figure 4
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  		FIG. 4. Interaction between Gtf1 and Gtf2 determined by yeast two-hybrid and in vitro GST pull-down assays. (A) Yeast two-hybrid analysis of the interaction between Gtf1 and Gtf2. The full-length gtf1 and gtf2 genes were cloned into yeast two-hybrid reporter vectors pBD and pAD to construct pAD-gtf1, pBD-gtf1, pAD-gtf2, and pBD-gtf2, respectively. AH109 yeast was transformed with different combinations of bait and prey plasmids. The resulting growth of cotransformants was used to score protein-protein interactions. (B) In vitro GST pull-down assays for interactions between purified GST fusion proteins and in vitro-translated Gtf1 or Gtf2 products. The purified GST, GST-Gtf2, or GST-Gtf1 glutathione Sepharose beads were incubated with in vitro-translated c-Myc-Gtf1 (A) and c-Myc-Gtf2 (B) to pull down the interacting partners, and the captured protein complexes were subjected to Western blot analyses with c-Myc antibody. Inputs represented in vitro-translated c-Myc-Gtf1 (A) and c-Myc-Gtf2 (B) and serve as positive controls.

Direct interaction between Gtf1 and Gtf2 determined by in vitro GST pull-down assays. Yeast two-hybrid analysis demonstrated that Gtf1 interacts with Gtf2. To determine the direct interaction between Gtf1 and Gtf2, we performed in vitro GST pull-down assays. Gtf1 and Gtf2 were expressed as GST fusion proteins, and the purified fusion proteins were used to capture the in vitro-translated Gtf2- and Gtf1-c-Myc fusion proteins. GST-Gtf2 interacted strongly with c-Myc-Gtf1 (Fig. 4B, lane 2) and vice versa (Fig. 4B, lane 5), whereas GST did not interact with either of the c-Myc fusion proteins (lanes 3 and 6), demonstrating Gtf1 and Gtf2 directly interacted with each other in vitro.

Interactions between Gtf1 and Gtf2 in vivo in a functional Fap1{Delta}RII glycosylation system. Gtf1 and Gtf2 were required for Fap1{Delta}RII glycosylation in E. coli. We identified the interaction between Gtf1 and Gtf2 by yeast two-hybrid and GST pull-down analyses. To determine the interaction between Gtf1 and Gtf2 in vivo where the glycosylation occurs, we characterized the functional Fap1 glycosylation system that harbors Fap1{Delta}RII substrate (pAL81) and Gtf1-Gtf2 derivatives (pAL200, pAL201, or pAL202). In the presence of both Gtf1 and Gtf2, two proteins were copurified (Fig. 5A, lane 4): the top one migrated at an 86-kDa position corresponding to the predicted GST-Gtf1 fusion protein, and the bottom one migrated with a size of 50 kDa corresponding to the predicted Gtf2 protein. This 50-kDa protein band was absent when the gtf2 gene was deleted (Fig. 5A, lane 3). LC/MS/MS mass spectrometry analysis of this excised 50-kDa band identified this protein as Gtf2 (Fig. 5B). This 50-kDa band could not be copurified when we attempted to purify GST (Fig. 5A, lane 1) or a truncated GST-Gtf1 fusion protein (Fig. 5A, lane 2). These results suggested Gtf1 interacted with Gtf2 specifically in the in vivo Fap1{Delta}RII glycosylation system.


Figure 5
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  		FIG. 5. Gtf1 and Gtf2 interact with each other in the E. coli glycosylation system. (A) In vivo GST pull down. The recombinant Fap1 substrate plasmid pAL81 was cotransformed with various pGEX6p1 derivatives into E. coli to generate a series of recombinant strains, AL400/pGEX6p1 (lane 1), AL402/pGEX6p1-gtf1({Delta}21-485)-2 (lane 2), AL403/pGEX6p1-gtf1-2({Delta}15-417) (lane 3), and AL401/pGEX6p1-gtf1-2 (lane 4). GST fusion proteins were purified from these recombinant strains and stained with Coomassie blue. MW, molecular mass. (B) MS analysis of Gtf2. A 50-kDa protein was copurified with GST-Gtf1, excised, and subjected to LC/MS/MS (B). Six tryptic peptides identified by MS matched with predicted peptides from Gtf2. Numbers indicate the amino acid positions in Gtf2.

The N-terminal region of Gtf1 is important for the interaction and required for the glycosylation of Fap1{Delta}RII. Gtf1 interacted with Gtf2. In order to map the Gtf1 domain responsible for the interaction in detail, we randomly inserted 19 aa in Gtf1 using an in-frame transposon EZ-Tn5 insertion system. Four different mutants with the transposon elements inserted in the gtf1 region of pAL205 were identified and used to examine the effect of insertions on the interaction between Gtf1 and Gtf2. The plasmids harboring a nonpolar insertion at amino acid residues 31, 65, 159, and 416 of Gtf1 were designated as pAL207, pAL208, pAL209, and pAL300 (Table 4), respectively. These plasmids were cotransformed with the Fap1{Delta}RII substrate plasmid pAL81 to generate corresponding mutants AL408, AL409, AL410, and AL411 (Table 4). The expression and glycosylation profile of Fap1{Delta}RII of these mutants was determined by MAb E42 and lectin sWGA, respectively. Two mutants (AL408 and AL409) that have the insertions at the N-terminal region of Gtf1 failed to glycosylate Fap1{Delta}RII with GlcNAc-containing oligosaccharides (Fig. 6A, lanes 3 and 4). This phenotype was similar to that of the mutant AL411 (Fig. 6A, lane 6), which has the insertion at the C-terminal region of Gtf1. The Fap1{Delta}RII of these three mutants migrated at the same position as that of the negative control (Fig. 6B, lanes 3, 4, 6, and 2). Another mutant, AL410, which has the insertion at the amino acid residue 159 position, did not alter the modification pattern compared with the positive control AL406 (Fig. 6A and B, lanes 5 and 1), indicating the insertion in this position had no effect on Fap1{Delta}RII glycosylation.


Figure 6
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  		FIG. 6. The Gtf1 N-terminal region is critical for Gtf1 and Gtf2 interaction and Fap1 glycosylation. A variety of plasmids generated by using the EZ-Tn5 transposon system (Table 4) were cotransformed with pAL81 into E. coli to construct a positive control strain, AL406/pGEX6p1-gtf1-2T7 (lane 1); a negative control, AL407/pGEX6p1-gtf2T7 (lane 2); and several in-frame insertional mutants, AL408 gtf1::Tn31-gtf2 (lane 3), AL409 gtf1::Tn65-gtf2 (lane 4), AL410 gtf1::Tn159-gtf2 (lane 5), and AL411 gtf1::Tn416-gtf2 (lane 6). Cotransformants were subjected to immune blotting with lectin sWGA (A) and MAb E42 (B). GST fusion proteins were purified from those cotransformants and analyzed by Western blotting assays with anti-T7 antibody (C). MW, molecular mass.

A sequence similarity search revealed the C-terminal region of Gtf1 has a highly conserved glycosyltransferase catalytic domain; this may explain why insertion in this region (position 416) impaired Fap1 glycosylation. However, the function of the N-terminal domain of Gtf1 is not clear. The interaction between Gtf1 and Gtf2 is required for Fap1 glycosylation. Therefore, we hypothesized the N-terminal region of Gtf1 modulated its interaction with Gtf2. To test this, we performed in vivo pull-down experiments in the Fap1{Delta}RII glycosylation system. GST-Gtf1 fusion proteins were purified and adjusted to the same concentration semiquantitatively based on Coomassie blue staining of SDS-PAGE gel. We analyzed the copurified Gtf2 by Western blotting using T7 antibody and found the Gtf2-T7 fusion protein was absent from or dramatically reduced in AL408 and AL409 (Fig. 6C, lanes 3 and 4), suggesting that the interaction between Gtf1 and Gtf2 was abrogated in AL408 and significantly reduced in AL409. Notably, the interaction between Gtf1 and Gtf2 in AL408 was comparable to that in the negative control (Fig. 6C, lane 2), indicating the interaction was crucial for Fap1 glycosylation. Meanwhile, a putative catalytic mutant, AL411, failed to glycosylate Fap1{Delta}RII (Fig. 6B, lane 6), but the mutation did not alter the protein-protein interaction (Fig. 6C, lane 6). These data suggested that amino acid position 416 was critical for the Gtf1 enzyme activity but not required for the protein-protein interaction.

In conclusion, we identified two Gtf1 regions that were important for Fap1 glycosylation: the N-terminal region was critical for the enzyme complex formation, whereas the C-terminal domain was needed for the catalytic function of Gtf1.


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DISCUSSION
 
In this study, we determined (i) GlcNAc was among the first sugar residues transferred to the Fap1 peptide and (ii) the formation of a protein complex by two putative glycosyltransferases was required for the transfer. We engineered a recombinant Fap1 molecule that contains the serine-rich repeat region (RI) of Fap1 to study the glycosylation of Fap1. This recombinant molecule displayed a monosaccharide composition profile similar to mature Fap1. The predominant monosaccharide residues associated with the recombinant Fap1, N-acetylglucosamine and glucose, are also found in mature Fap1. Glycosyl linkage analyses suggested that all of the oligosaccharides were initiated from a GlcNAc residue, indicating that GlcNAc is the initial sugar transferred to the Fap1 peptide backbone.

To test the hypothesis that Gtf1 and Gtf2 are initial glycosyltransferases responsible for the transfer of GlcNAc-containing oligosaccharide(s), we established an in vivo Fap1 glycosylation system in E. coli. This system utilized the recombinant Fap1 (Fap1{Delta}RII) as a substrate and Gtf1 and Gtf2 as the only streptococcus-derived putative glycosyltransferases. This model excludes the effect of other streptococcal glycosyltransferases on Fap1 glycosylation and has allowed us to dissect the initial glycosylation step. Using this system, we confirmed that putative glycosyltransferases Gtf1 and Gtf2 are essential for production of the GlcNAc-containing oligosaccharides, supporting the notion that GlcNAc is among the first sugar molecules transferred to the peptide backbone. Interestingly, the oligosaccharides released from the recombinant Fap1 isolated from S. parasanguinis were not homogenous. The question arises as to why the recombinant Fap1 modification is homogenous in E. coli and heterogeneous in S. parasanguinis. A plausible explanation is that other putative glycosyltransferases identified from the Fap1 locus that are important for the subsequent Fap1 glycosylation in S. parasanguinis are missing in this recombinant E. coli strain. The GlcNAc residue-containing glycan moiety has also been identified in other serine-rich proteins, including GspB of S. gordonii (1) and SrpA of S. sanguinis (21). Therefore, it is possible that the transfer of GlcNAc-containing sugar residues to the serine-rich regions by Gtf1 and Gtf2 homologues may represent a common theme as the Gtf1 and Gtf2 molecules are highly conserved throughout streptococci and staphylococci that express Fap1-like proteins. Furthermore, Gtf1 and Gtf2 isolated from S. sanguinis and S. pneumoniae were able to glycosylate Fap1{Delta}RII with GlcNAc-containing oligosaccharides (data not shown), supporting the concept that Gtf1 and -2 homologues are capable of the transfer.

Genetic analyses of Gtf1 and Gtf2 mutants in both S. parasanguinis and E. coli systems suggested the two putative glycosyltransferases were required for Fap1 glycosylation. This result is consistent with a previous report that Gtf1 and -2 homologues are important for GspB glycosylation in S. gordonii (31). Protein glycosylation is modulated by glycosyltransferase-mediated initiation and elongation in a stepwise manner. Many glycosyltransferases of mammalian and eukaryotic cells are functionally and physically associated with each other to facilitate the glycosylation process (16). We have shown Gtf1 and Gtf2 formed a protein complex by yeast two-hybrid and in vitro GST pull-down assays. Furthermore, the interaction between two putative glycosyltransferases appears to be essential in the in vivo glycosylation system that simulates the initial Fap1 glycosylation reaction. All of these results suggest the formation of Gtf1 and Gtf2 complex was required for the initial glycosylation. We have identified two Gtf1 domains that are differentially involved in their interaction with Gtf2 and Fap1 glycosylation. A Gtf1 N-terminal domain directly interacted with Gtf2, thereby modulating Fap1 glycosylation. The C-terminal amino acid position 416 was not critical for this interaction; however, it was required for Fap1 glycosylation, suggesting it has a glycosyltransferase catalytic domain. Indeed, a domain search revealed the C-terminal region has a highly conserved glycosyltransferase activity motif.

GtfA and GtfB of S. gordonii, homologues of putative glycosyltransferases of S. parasanguinis, are critical for glycosylation of GspB (31). However, whether they interact with each other is not known. Given the amino acid sequence similarity observed among glycosyltransferases from a variety of streptococci and staphylococci, it is reasonable to assume the interaction between putative glycosyltransferases also exists in these streptococcal and staphylococcal species as well. It is not clear how the interaction between Gtf1 and Gtf2 mediates enzymatic activity that regulates Fap1 glycosylation. Conceivably, Gtf1 and Gtf2 are involved in sequential biosynthesis of Fap1 glycans and the interaction between Gtf1 and Gtf2 promotes the efficient sequential biosynthesis of Fap1 glycans. Alternatively, Gtf1 and Gtf2 may be an integral part of an enzyme complex. Perhaps Gtf2 functions like a chaperone and contributes a missing fold to Gtf1, making both essential for enzymatic activities. This hypothesis is in agreement with the fact that both gtf1 and gtf2 mutants showed very similar phenotypes: i.e., expression of a 360-kDa Fap1 precursor. Many proteins responsible for carbohydrate biosynthesis form heterocomplexes that facilitate glycan biogenesis. Biosynthesis of ganglioside precursors requires the assembly of distinct multiple enzyme complexes including glycosyltransferases (13). Two members of β1,3-N-acetylglucosaminyl-transferase family, β3Gn-T2 and -T8, interact with each other and can enhance corresponding enzymatic activities in vitro (24). However, whether the interaction between the two enzymes occurs in vivo has not been demonstrated. This is the first report that describes the importance of the interaction between two putative glycosyltransferases in the glycosylation of a bacterial adhesin. As many examples of protein-protein interaction between glycosyltransferases have been documented in eukaryotic systems, our findings indicate the formation of protein complexes by interactions between glycosyltransferases is a common theme in protein glycosylation, whether they are prokaryotic or eukaryotic in origin.


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ACKNOWLEDGMENTS
 
This work was supported by NIH grants DE014726 and DE 017954 (H. Wu) and DE11000 (P. Fives-Taylor) from the National Institute of Dental and Craniofacial Research.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Pediatric Dentistry, UAB Schools of Dentistry, Birmingham, AL 35294. Phone: (205) 996-2392. Fax: (205) 975-6251. E-mail: hwu{at}uab.edu Back

{triangledown} Published ahead of print on 14 December 2007. Back

{dagger} Present address: Department of Laboratory Medicine, Union Hospital, Institution of Immunology, Tongji Medical College, HuaZhong University of Science and Technology, Wuhan, Hubei, People's Republic of China. Back


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Journal of Bacteriology, February 2008, p. 1256-1266, Vol. 190, No. 4
0021-9193/08/$08.00+0     doi:10.1128/JB.01078-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.



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