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Department of Biological Sciences, CW-405 Biological Sciences Building, University of Alberta, Edmonton, AB T6G 2E9, Canada
Correspondence
Laura S. Frost
laura.frost{at}ualberta.ca
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ABSTRACT |
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Present address: Department of Chemistry, 456 Chemistry Building, York University, 4700 Keele St, Toronto, ON M3J 1P3, Canada.
Present address: National Center for Biotechnology Information/National Institutes of Health, 6th Floor, 45 Center Drive, Bethesda, MD 20892, USA.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Wilkins & Frost, 2001). Although conjugative plasmids have been identified in several bacterial species, the F plasmid from Escherichia coli remains a paradigm for the study of bacterial conjugation (Lawley et al., 2004). The 100 kb F plasmid encodes all genes necessary for conjugation in the 33.3 kb transfer (tra) region, including genes for pilus synthesis, mating pair formation (Mpf) and DNA transfer (Frost et al., 1994). Conjugative systems have recently been classified as a subset of the type IV secretion system (T4SS), due to the considerable similarity between the genes involved in both processes (Lawley et al., 2003).
The conjugative transfer of DNA is a plasmid-driven process that involves an intimate association between a donor and recipient cell, termed a mating pair (Lederberg, 1956; Achtman, 1975; Durrenberger et al., 1991). This is followed by the transfer of plasmid ssDNA through a membrane-associated, supramolecular structure known as the Mpf complex. In F-like systems, mating pair stabilization (Mps) allows cells to mate more efficiently in liquid media, and to more readily resist disaggregation by mechanical forces (Bradley et al., 1980; Manning et al., 1981). Initially, Mps was thought to be mediated through the F pilus alone. However, further analysis has shown that, whereas the F pilus is important for initiating Mpf, the TraN and TraG proteins are responsible for the phenotypes of resistance to SDS and shear forces (Achtman et al., 1977). Kingsman & Willetts (1978) have shown that mutations in traN and traG allow the initiation of conjugative DNA replication in the donor, but not the transfer of radiolabelled F plasmid DNA to the recipient cell. Another F protein, TraU, has been shown to be involved in DNA transfer, but not pilus synthesis, and is tentatively grouped with the Mps proteins (Moore et al., 1990; Lawley et al., 2003). F TraN is a cysteine-rich, outer-membrane protein that requires OmpA in the outer membrane of the recipient cell for efficient mating in liquid cultures (Klimke & Frost, 1998; Klimke et al., 2005). TraN has also been suggested to interact with TraG, although this remains unproven (Firth & Skurray, 1992; Klimke et al., 2005). A counterpoint to Mps is provided by two processes involved in the prevention of redundant DNA transfer between donor cells. The first is surface exclusion (Sfx), encoded by traT, which impedes contact between donor cells carrying the F plasmid (Achtman et al., 1977; Harrison et al., 1992; Klimke et al., 2005). The second process is entry exclusion (Eex), encoded by traS, which prevents the transfer of DNA between equivalent donor cells (Kingsman & Willetts, 1978). The Eex systems of the closely related plasmids F and R100 are plasmid-specific, whereby the transfer apparatus in the donor cell recognizes, via TraG, the cognate TraS in the recipient cell (Anthony et al., 1999). Recently, the integrating conjugative elements SXT and R391 from Vibrio cholerae and Providencia rettgeri, respectively, have been shown to behave in a similar fashion to TraG, recognizing its cognate TraS during Eex (Marrero & Waldor, 2005).
At 102.5 kDa, F TraG is one of the largest proteins encoded by the tra region of the F plasmid (Frost et al., 1994). Mutations that map to the N-terminal half of F TraG (e.g. traG106, frameshift) affect pilus assembly, whereas mutations in the C-terminal half of F TraG, such as traG101 (frameshift) and traG79 (amber), affect Mps but permit pilus synthesis, as monitored by phage infection (Willetts & Achtman, 1972; Fig. 1). Antibodies directed towards the C-terminal domain of F TraG detect two products: full-length F TraG and, within the periplasm, a 50 kDa product named TraG* (Firth & Skurray, 1992). This is not to be confused with the TraGp* fusion product that results from in-frame cloning of the proximal end of traG to a protein coding sequence in pRS29 via an EcoRI site (Manning et al., 1981). Sequence analysis of TraG predicts the presence of a signal peptidase I cleavage site after residue 451, generating a 52.7 kDa product that could be TraG*, which may be released into the periplasmic space and contribute to Mps (Firth & Skurray, 1992).
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METHODS |
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), as has ED24 (SpR, F– Lac–) (Achtman et al., 1971). Vectors included pBAD24 and pBAD33 (Guzman et al., 1995), pCR-Blunt-TOPO (Invitrogen), pALTER-Ex2 (Promega) and pET26b (Novagen). Conjugative plasmids used included FlactraG79, FlactraG106, FlactraG101 (Achtman et al., 1972) and pOX38 : : Tc (Anthony et al., 1994). pOX38 : : Tc
GKm was constructed by replacing the PstI/PstI fragment in traG of pOX38 : : Tc with a Km resistance cassette using recombination, as described by Klimke et al. (2005).
ISphoA/in insertions into F traG.
A 3.0 kb fragment containing F traG was subcloned into pBSII SK+, which also eliminated the BamHI site in the multiple cloning site of the vector. This construct, named pBK20, was used to insert ISphoA/in into traG, according to previously described procedures (Manoil & Bailey, 1997; Klimke et al., 2005). DNA isolated from ISphoA/in derivatives of traG was treated with BamHI, and the larger DNA fragment was religated to give mutants of traG with 31 aa epitopes fused in-frame in the traG coding sequence.
Construction of ppelBTraG*.
nt 1357–1717 of the coding region of TraG* (aa 452–938) were amplified by PCR with Vent polymerase (New England Biolabs), using forward primer LFR193 (5'-CCATGGCAGGCAGTGTGG-3') containing an NcoI site at the 5' end, and reverse primer G21120 (5'-GCAGTGACTCTGTCACTGG-3'), and cloned into pCR-Blunt-TOPO. After NcoI and SmaI digestion, the 5' proximal portion (front) of traG was moved into a pBAD24 construct carrying the back half (SmaI to 3' end; nt 1718–2814) of traG to produce pTraG*. The traG* fragment of this construct was cloned using NcoI and HindIII into pET26b immediately downstream of the pelB leader sequence. The pelB leader sequence and the TraG* encoding sequence were then recloned into pBAD24, using XbaI and HindIII, generating the plasmid ppelBTraG*.
Generation of traG point mutations.
The PstI/PstI fragment from the F traG gene was cloned into pALTER-Ex2. A mutation was induced in this fragment using the Altered Sites II in vitro Mutagenesis Systems kit (Promega), according to the manufacturer's instructions. The base at position 1353 was changed from G to C, resulting in an amino acid change from alanine to proline at residue 452. Additional mutations at A449 (A449D) and A451 (A451E) were similarly constructed. The altered DNA was reintroduced into full-length traG in pBAD24 by restriction digest and ligation, and called pTraGA452P, pTraGA451E and pTraGA449D, respectively.
Construction of chimeric F and R100 TraG and TraS clones.
R100 and F traG chimeras were constructed using conserved restriction sites in the two genes (Fig. 1
). The chimeras were named by referring to the source of the DNA in the 5' gene fragment (nt 1–1599), middle section (nt 1600–2160) and the 3' end (nt 2161–2814), using F and R for the F and R100 plasmids, respectively. All constructs with F sequences in the 5' region were in the pBAD24 vector, while chimeras starting with the R100 sequence were in pT7.3.
Construction of traS vectors.
pTraS-F was prepared by amplification of F traS using forward primer LFR183 (5'-CCATGGGTAGGGTATGGAG-3') containing a 5' NcoI site, and reverse primer LFR184 (5'-AAGCTTTTATTTTACTCTTGATAAC-3') with a 5' HindIII site. The PCR product was ligated into pCR-Blunt-TOPO, digested from this vector with NcoI and HindIII, and ligated into pBAD24 digested with the same enzymes. pTraS-R100 was prepared in a similar manner using the forward primer LFR191 (5'-CCATGGCTGTTTTTGGGAGATG-3') with a 5' NcoI site, and reverse primer LFR192 (5'-AAGCTTAAAACGTCATAATATAC-3') containing a 5' HindIII site. A six-histidine sequence (His6 tag), which was fused to the N or C terminus of F TraS, was similarly constructed, but with the coding sequence for the six histidines included in the primers used for amplification.
Membrane separation by density flotation.
Density flotation of cellular membranes was done according to the procedure of Grahn et al. (2000), and isolated cell membranes were resuspended in 2 ml 55 % sucrose. All sucrose solutions (w/v) were made up in 10 mM Tris/HCl, pH 7.5, and 5 mM EDTA; gradients were prepared in a stepwise fashion with layers of 0.5 ml 60 % sucrose, 2 ml sample, 2 ml 50 %, 3 ml 45 %, 3 ml 40 %, 1 ml 35 % and 0.5 ml 30 % sucrose. Gradients were centrifuged in a Beckman SW41 rotor at 200 000 g for 72 h. Fractions of 750 µl were collected from the top, the protein content of the samples was measured using a Bradford assay (data not shown), and 15 µl aliquots of each fraction were analysed by Western blotting for TraG, TraD, OmpA and CpxA, as follows. Proteins from SDS-PAGE were transferred to PVDF membranes, according to the manufacturer's instructions. Identification of TraG, TraD, CpxA and OmpA was done using standard procedures and employed rabbit polyclonal primary antibodies at the following dilutions: anti-TraG, 1 : 5000; anti-TraD, 1 : 15 000; anti-OmpA, 1 : 100 000; and anti-CpxA, 1 : 5000. Chemiluminescent detection employed a goat anti-rabbit horseradish peroxidase-conjugated secondary antibody (Amersham) at a dilution of 1 : 20 000.
Detection of periplasmic TraG.
pBAD24 constructs containing traG, traGA452P, pelBtraG* and a vector control were transformed into XK1200/pOX38 : : TcGKm. Cultures (2 ml) of these transformants were grown in LB broth to mid-exponential phase, and induced with 0.1 % arabinose for 1 h at 37 °C. Periplasmic shock preparation proceeded as follows. Cell pellets from 1.5 ml chilled, induced cultured cells were washed twice in 0.8 ml chilled 10 mM Tris/HCl (pH 7.5) and 30 mM NaCl. The washed cells were pelleted and resuspended in 20 % sucrose and 33 mM Tris (pH 7.5), and then EDTA was added to a concentration of 1 mM. After incubation at room temperature with gentle rocking for 10 min, the cells were pelleted and resuspended in a small amount of supernatant, and chilled. The resuspended cells were introduced into 10 ml cold 0.5 mM MgCl2, containing Complete Mini Protease Inhibitors–EDTA (Roche), and stirred gently for 10 min. The resulting shock fluid was cleared of cells by centrifugation, and 0.5 ml aliquots were TCA-precipitated for SDS-PAGE and Western blotting analysis, which proceeded as described above. Samples of induced whole cells (OD600 0.1) were also included for comparison.
Mating assays.
Mating assays were performed as described by Anthony et al. (1994). Briefly, 2 ml cultures were grown to mid- to late-exponential phase in LB broth with appropriate antibiotics. pBAD constructs were induced with 0.1 % arabinose for 1 h at 37 °C. Cells were pelleted, washed once with LB broth to remove antibiotics, and resuspended in an equal volume of LB broth. One hundred microlitres each of donor and recipient cells (typically 1x107 to 4x108 cells ml–1 were used in each mating assay) were mixed with 0.8 ml LB broth and 0.1 % arabinose for pBAD constructs, and allowed to mate for 45 min at 37 °C. The mating mixtures were vortexed vigorously and put on ice to prevent further mating. Mating cultures were serially diluted 10-fold in 4 °C SSC buffer (150 mM sodium chloride, 15 mM sodium citrate, pH 7.0) five times, and 10 µl of each dilution was spot-plated separately on agar plates selective for donors, recipients or transconjugants. Colonies were counted after overnight incubation at 37 °C, and the number of transconjugants per 100 donor cells was calculated.
Bioinformatics software.
Sequence alignments of F and R100 TraG and TraS were performed using CLUSTALW (Thompson et al., 1994). Figs. 4 and 5 were prepared with minor manual editing of the CLUSTALW alignments. Transmembrane helices were predicted using TMHMM (Krogh et al., 2001; http://www.cbs.dtu.dk/services/TMHMM-2.0/).
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RESULTS |
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). TraG is predicted to contain three (aa 54–73, 327–385 and 407–447; Firth & Skurray, 1992) or five (aa 54–73, 327–348, 364–385, 407–426 and 428–447; Frost et al., 1994) transmembrane 
-helices, as well as a putative C-terminal fragment (TraG*, aa 452–938) in the periplasm (Achtman et al., 1972; Manning et al., 1981; Firth & Skurray, 1992). TraG was confirmed to localize to the inner membrane, using density flotation gradients to separate inner and outer membranes. The fractions were analysed by immunoblotting using antibodies to TraG, as well as known inner (CpxA, TraD; Weber & Silverman, 1988; Panicker & Minkley, 1992) and outer membrane proteins (OmpA; Sonntag et al., 1978) (Fig. 2).
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) was an intriguing idea that merited further exploration, since it would explain the need for the C-terminal region of TraG for Mps. TraG was detected as a faint band in single-copy F+ cells, but TraG* could have been below the limits of detection (data not shown). The existence of TraG* was further assessed in periplasmic fractions from cells carrying pBAD24 derivatives of either wild-type TraG or a signal peptidase-resistant mutant (TraGA452P) (Fig. 3). These were compared to pelBTraG*, which is a fusion of the putative TraG* sequence to the PelB signal sequence to ensure transport to the periplasm. In cells expressing pelBTraG*, the TraG-derived polypeptide was present in the periplasmic fraction, and acted as a size marker for the putative TraG* protein (52.7 kDa). When cellular lysates or periplasmic shock fractions were probed with an anti-TraG antibody, full-length TraG and associated degradation products were found in cell lysates containing TraG or the TraGA452P mutant, whereas no TraG-derived bands were found in the periplasm. We conclude that TraG* does not exist under normal circumstances. Immunoblots of mating cells did not reveal a band that could be interpreted as TraG* (data not shown), suggesting that it is not generated during Mpf.
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; Frost et al., 1994) was examined using ISphoA/in insertions that can be converted to in-frame 31 aa epitope fusions, as described by Manoil & Bailey (1997). Twelve separate ISphoA/in insertions into TraG were obtained in four unique sites in TraG [aa D159 (one insertion), N163 (six insertions), L716 (one insertion) and E739 (four insertions)]. D159 and N163 resided in the first predicted periplasmic loop (aa 74–326), and L716 and E739 were within the C-terminal domain of TraG that extended into the periplasm (Fig. 4). No inserts were isolated for the second predicted periplasmic domain (aa 386–406), possibly because the target was too small (60 bp), or because there are three, not five, transmembrane helices. Only the insert at codon E739 (pBKG739) could be converted to a stable 31 aa fusion. Using a complementation assay with pOX38 : : TcGKm, pBKG739 had a mating efficiency that was 15 % of that of the wild-type, suggesting that insertion of the 31 aa in the periplasmic domains greatly affected TraG function.
The nature of the mutations originally described by Willetts & Achtman (1972) was characterized by sequencing clones of each of the mutant traG genes. The mutations in traG106 and traG101 were +1 frameshifts in runs of Gs or Cs starting at nt 19795 and 21606 (aa T131 and T735), respectively, according to the sequence presented by Frost et al. (1994) (Fig. 4). The mutation in traG79 was a C to T transition at nt 21341, resulting in a glutamine (CAG, Q647) changing to an amber codon (Fig. 4).
Mps requires full-length TraG
We asked whether TraG* alone, expressed as pelBtraG* (ppelBTraG*), could complement a frame-shift mutant capable of synthesizing pili (FlactraG101), with pOX38 : : TcGKm acting as a control. Using mating experiments, only clones expressing full-length TraG could complement both pOX38 : : TcGKm and FlactraG101 donors (Table 1). ppelBTraG* partially complemented FlactraG101 at very low levels in some assays, which was thought to be due to reversion of the frameshift mutation in FlactraG101.
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), mutations that are predicted to severely alter peptidase activity were introduced into TraG by site-directed mutagenesis at positions –3, –1 and +1 of the predicted cleavage site [residues 449–452 (ASA-A); G*, Figs 1 and 4]. Thus +1 alanine was converted to proline (A452P), –1 alanine to glutamic acid (A551E), and the –3 alanine to aspartic acid (A449D). These mutations were introduced into pTraG and used in complementation assays with pOX38 : : TcGKm and FlactraG101, which synthesizes F pili (Table 1). The lack of complementation by pTraGA452P suggested either that the proline altered the conformation of TraG or that A452 was indeed important for TraG function. However, two other point mutations, TraGA449D and TraGA451E, complemented pOX38 : : TcGKm and gave 40–80 % of wild-type results for pOX38 : : TcGKm/pTraG (Table 1). We conclude that there is no signal peptidase I cleavage site in the middle of TraG, and that the proline at position 452 likely introduced a kink into the periplasmic domain that affected TraG function in Mps. Thus, full-length TraG is required for Mps with both the N- and C-terminal regions participating in establishing stable mating junctions.
Eex involves plasmid-specific recognition between TraG and TraS
Eex is a plasmid-specific process, defined by TraS in the recipient cell, which blocks DNA transport between cells carrying the same conjugative plasmids (Ou, 1975; Kingsman & Willetts, 1978) by recognizing its cognate TraG in the donor cell (Anthony et al., 1999; Marrero & Waldor, 2005). While the F and R100 TraS clones were being constructed, we realized that the sequences originally presented for R100 TraS (Ogata et al., 1982) and F TraS (Jalajakumari et al., 1987) were incorrect. The correct accession numbers are NP_061479 (F TraS; gi398520) and NP_052977 (R100 TraS; gi5036330), based on more recent sequence data (Fig. 5). TraS was fused to a His6 tag at both the N and C termini, and tested for Eex using mating efficiency assays. Both fusions gave Eex comparable to wild-type TraS (Table 2). Using anti-His6 antibodies, the presence of TraS, expressed from a pBAD24 clone that was induced with 0.05 % arabinose, was confirmed to be in the inner membrane by flotation density gradient analysis (data not shown).
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) that were assessed for their ability to restore Eex in traG-deficient donor cells (pOX38 : : TcGKm; Table 3). When native TraG (TraGFFF or TraGRRR) was assessed, only the corresponding TraS (F or R100, respectively) in the recipient cell gave Eex. This was narrowed down to the middle region of TraG (aa 533–717 in F), since this region was required for recognition of the cognate TraS in the recipient cell (Table 3). All other TraG chimeras had low to undetectable Eex. With the exception of a single amino acid difference at position 558 (S to T), the only differences in the sequences of aa 533–717 in F and R100 TraG were within aa 610–673 (Fig. 4), which further delimits the region of TraG involved in Eex.
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). Neither TraG nor TraG* interfered with Eex, suggesting that TraG and TraS interacts in trans between donor and recipient cells. Thus, TraG must be delivered to the recipient cell inner membrane by a process that defines the phenomenon of Mps. |
DISCUSSION |
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). Sequence homology has been noted between the N-terminal domain of TraG and VirB6, a polytopic inner-membrane protein of the Ti plasmid of Agrobacterium tumefaciens also involved in T pilus synthesis and T-DNA transport (Lawley et al., 2003). However, unlike Ti VirB6, which is predicted to contain between four and nine transmembrane helices (Christie, 2001; Judd et al., 2005), TraG is predicted by hydropathy plot analysis (Firth & Skurray, 1992; Frost et al., 1994) to span the inner membrane three to five times, with large periplasmic regions at approximately aa 75–325 and 445–938 (C terminus). Our limited number of ISphoA/in insertions confirmed these results, and suggested that there were at least three transmembrane domains. Interestingly, the only ISphoA/in insertion that could be converted to a stable 31 aa epitope fusion (at E739) is very close to the traG101 frameshift mutation at T735 that affects Mps but not pilus synthesis. No evidence was observed for TraG*, a periplasmic fragment suggested to be involved in Mps (Firth & Skurray, 1992). Degradation products in the lysate of cells overproducing TraG were approximately correct in size (Fig. 3), but none of these products was found in the periplasm. The observation that TraG, but not TraG*, can fully complement FlactraG101 suggests that whole TraG is required for biological activity, which also argues against the presence of TraG*.
Examination of the ability of F or R100 transfer proteins to complement F tra mutations and affect phage sensitivity or Eex suggests that TraG in the donor cell is specific for its cognate TraS in the recipient cell (Anthony et al., 1999). A comparison of the sequences of TraG from the F and R100 plasmids indicates that they share an overall sequence identity of 93 %. However, the region between residues 610 and 673 of TraG shows the most dissimilarity, with an overall sequence identity of 55.7 % (Fig. 4). This region is within the portion of TraG in the chimeric plasmids (aa 533–717) that exhibits specificity for TraS, and is probably responsible for this phenomenon.
Unlike TraG, TraS from F and R100 plasmids were highly distinct, exhibiting only 17 % sequence identity (Fig. 5). The orientations of F and R100 TraS in the membrane were difficult to predict and could be quite different for the two proteins. Using the TMHMM algorithm, three or four transmembrane helices were predicted for R100 and F TraS, respectively, but in significantly different locations. Thus, there is no limited region of dissimilarity in TraS to aid in predicting the features that are recognized by TraG. Marrero & Waldor (2005), who have studied Eex specificity in the more closely related SXT and R391 systems, have been able to narrow the specificity region of TraG to three amino acids, aa 606–608, which approximates the location of the dissimilar region in F and R100 TraG (aa 610–673). They have also been able to identify the C-terminal half of TraS as being responsible for Eex specificity. Our attempts to construct stable TraS chimeras were unsuccessful, possibly due to incorrect insertion of the chimeras in the membrane.
TraG is involved in pilus synthesis as well as Mps; therefore, it would be expected to interact with other proteins in the donor cell, although none have so far been identified. There has been speculation that TraG might form a transenvelope bridge and contact TraN in the outer membrane (Firth & Skurray, 1992); however, this interaction has not been detected using cross-linking (Klimke et al., 2005). We are currently exploring the possibility that TraG interacts with TraU, which is a putative Mps protein, and is known to be part of an interaction complex that is involved in pilus synthesis, but is not itself required for this process (Moore et al., 1990; Harris & Silverman, 2004).
This report further defines a third function of TraG, that of Eex through interaction with TraS in the recipient cell. This function is plasmid-specific and appears to involve aa 610–673 of TraG. This TraG–TraS interaction occurs in trans between a donor (TraG) and recipient cell (TraS); supplying TraG or TraG* in the recipient cell did not affect mating or Eex. As both TraG and TraS are inner-membrane proteins, the mechanism by which they recognize each other is puzzling. Since Mps involves close contact between mating cells that is difficult to disrupt (Achtman, 1975), a process can be envisioned whereby F TraN binds OmpA in the recipient outer membrane and triggers pore formation. TraG would then be thrust into the periplasmic space to establish contact with the inner membrane of the recipient cell. If TraS interferes with this signalling mechanism, conjugative DNA synthesis and transfer are blocked. Attempts to isolate a TraG–TraS complex from mating cells by cross-linking or immunoprecipitation, or to demonstrate interaction using a bacterial two-hybrid system, have been unsuccessful. This appears to be due to the small percentage of the total amount of TraG or TraS involved in a single mating bridge, and possibly a requirement for higher-order multimeric structures for correct interaction between TraS and TraG.
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ACKNOWLEDGEMENTS |
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Edited by: L. Jannière
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Received 28 August 2006;
revised 30 October 2006;
accepted 2 November 2006.
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