Mutagenesis of plasmid R1-16 used the transposon delivery system pUT-miniTn5Cm (Nuk et al., 2011). A selection step requiring conjugative transfer of the R1-16 mutant derivatives was included to eliminate those with transposon insertions in the plasmid tra genes. One biofilm deficient mutant, R1-16miniTn5Cm E5, carried the transposon inserted at position 488 of parM (Accession Number X04268), effectively blocking transcription of parM and parR. Disruption of this locus did not lead to an immediate loss of plasmid from the population and donor cultures conjugated normally in a standard 30 min mating experiment (Nuk et al., 2011) Nonetheless, the poor biofilm formation and the complete R17 resistance of hosts carrying this mutant suggested that the parMRC locus could be involved in the assembly and function of the conjugation machinery. We first asked whether conjugative pili assembled in the absence of parMRC were defective for bacteriophage attachment. E. coli MS411 carrying R1-16 or R1-16miniTn5Cm E5 were combined with fluorescently labeled R17 and visualized microscopically (Figure 1A). Attachment of the labeled R17 to wild type pili was apparent but strikingly absent from hosts carrying the mutant. The attachment defect was complemented by expression of parM and parR in trans. These data suggest that pili assembled in the absence of parMRC have an abnormal conformation.

We then asked whether a defect in early stages of plasmid transfer could be detected. E. coli MS411 [R1-16miniTn5Cm E5] donors were combined with E. coli MS614 recipient cells in broth culture. Conjugation was interrupted after 3–15 min and transconjugants were selected on agar plates. After 3 min of coincubation, transfer of the R1-16miniTn5Cm E5 was detected, but at frequencies 2–3 log units lower than transfer of wild type R1-16 (Figure 1B). Complete complementation of transconjugant formation at this time point was observed by providing parM and parR in combination on an expression plasmid in trans. Presence of either parM or parR was not sufficient. Significantly lower transfer frequencies were also observed for R1-16miniTn5Cm E5 compared to wild type after 5 and 15 min of conjugation. The magnitude of this difference decreased with increasing time, however. In every case, addition of parM and parR in trans fully complemented mating efficiency to wild type levels. We conclude that E. coli carrying the parMRC transposon insertion in R1-16 exhibits a delayed initiation of transfer phenotype that is fully overcome when cultures are allowed to conjugate for longer than 15 min.

Transfer initiation requires the activity of the DNA processing relaxosome complex. We asked next whether the parMRC disruption influences nicking of R1-16 oriT by TraI in vivo. R1-16 plasmids express conjugative genes constitutively and therefore support continuous relaxosome assembly. Within the relaxosome TraI maintains an equilibrium of cleaving and resealing of the nick site, meaning that a fraction of R1-16 will be in a nicked state at any moment (Zechner et al., 1997). When host cells are lysed for plasmid DNA isolation, the population of nicked molecules covalently attached to TraI should be lost during phenol extraction, lowering the yield. In contrast, any condition that disrupts oriT cleavage would allow R1-16 to remain supercoiled, increasing DNA recovery. To validate this assay of relative plasmid yield we combined R1-16 or mutant derivatives in E. coli M1174 cells with a second independent replicon. The two plasmids were copurified by phenol extraction, linearized once with XbaI and applied to agarose gels to detect quantitative variation in the apparent copy number of the conjugative plasmid relative to the second replicon. Figure 2 illustrates changes in plasmid ratios obtained by controlled manipulation of oriT DNA processing (Δ Dtr) through deletion of traI. The band intensities of the recovered R1-16 derivatives were normalized to the internal control plasmid and compared (Figures 2A,B). Values obtained for R1-16 wild type plasmid were set to 1. Disruption of the traI gene resulted in a nearly four-fold higher relative yield compared to wild type R1-16 DNA. Induction of traI expression in trans restored nicking activity, resulting in a plasmid yield significantly lower than in the absence of traI. We have used a similar assay previously (Nuk et al., 2011) to test whether random insertions of transposon miniTn5 in plasmid R1-16 result in plasmid instability. Validation of the assay in that case relied on controlled manipulation of the plasmid R1 copy number control system. In the current study, the plasmid recovery assay was applied to E. coli cells carrying R1-16 wild type, R1-16miniTn5Cm E5 and, as positive control, an additional transposon insertion in the yjcA gene close to the kis/kid stability locus of plasmid R1. Due to the insertion in the parMRC locus, we predicted relatively low yields of R1-16miniTn5Cm E5 compared to the reference replicon (Jensen and Gerdes, 1999), yet surprisingly the parMRC mutant derivative was obtained in higher relative yields than wild type R1-16 (Figure 2C). In comparison the control mutant B4, carrying the miniTn5 at the kis/kid locus, was poorly recovered, as expected for a destabilized plasmid. A possible explanation for the unexpectedly high yield of the parMRC mutant is that the assay outcome reflects a stronger defect in relaxosome activity than in plasmid stability. In that case, the partitioning components of the ParMRC system appear to enhance oriT DNA processing in vivo.

To test whether this effect of the partitioning components directly involves TraI we purified ParR and ParM proteins and measured the impact of these effectors on known enzymatic activities of the TraI enzyme in vitro. A standard oriT DNA-cleavage assay used to monitor relaxase enzyme activity measures the conversion of supercoiled plasmid substrate to the open circular form using agarose gel electrophoresis (Lanka and Wilkins, 1995; Csitkovits et al., 2004). A supercoiled substrate plasmid (4 nM) carrying 420 bp of R1 oriT (pDE100) was preincubated with putative effector proteins ParM or ParR and the reaction started by the addition of 25 nM TraI (Figure 3). The percentage of oriT DNA captured in open circular form was significantly enhanced by the additional presence of ParM or ParR in a concentration dependent manner. Maximum stimulation (~three-fold, 5–16%) was observed when ParM was present in equimolar amounts (20–30 nM) relative to TraI. ParR alone (10 nM) stimulated TraI relaxase activity nearly four-fold (11–38%). At higher concentrations ParM and ParR failed to stimulate. Moreover, no superstimulation was observed when both factors were present. We then asked whether the Par proteins also stimulate truncated versions of TraI in this assay (not shown). N-terminal fragment TraI _1–308 (TraIN308) forms the minimal relaxase domain, and residues 1–992 (TraIN992) comprise the relaxase and the complete activation domain absolutely required for all T4SS activities we have tested thus far. Titration of either Par effector protein to the reactions containing truncated forms of TraI did not result in stimulation of oriT cleavage. We conclude therefore, that both ParM and ParR stimulate the oriT cleaving and joining activity of TraI independently. Stimulation was observed exclusively with the full length TraI protein.

Given that ParM and ParR increase TraI relaxase activity but do not bind to oriT DNA we next tested for direct protein-protein interactions. Par protein fusions were engineered with terminal FLAG-epitopes. Expression plasmids for the tagged fusion proteins were maintained in E. coli MS411 cells carrying either R1-16, or a second vector expressing a candidate tra gene. Following induction of fusion protein expression cells were lysed, protein complexes briefly cross-linked with formaldehyde and the Par proteins captured on FLAG-affinity beads. Bound proteins and their specific interaction partners were washed, eluted, and visualized by western immunoblotting. Based on the phenotypic and biochemical results, candidates for specific Par protein binding included TraI and the second key factor involved in pilus specific R17 phage infection, T4CP TraD, as well as the VirB4-like ATPase TraC, which is essential for assembly and function of R1 pili. Tra proteins retained by Par proteins on the FLAG affinity matrix were detected with polyclonal antibodies generated to specific transfer proteins. The amounts of Tra proteins detected in the whole cell lysates reflect native levels produced from R1-16. The Tra specific antibodies revealed that TraI, TraD and TraC were co-retained by affinity beads together with ParM_C−FLAG (Figure S2). The specificity of these interactions was confirmed with a par allele lacking FLAG. In contrast ParR retained very small amounts of TraI and TraC but only with the C- terminal FLAG epitope (Figure S2). To assess whether the observed Tra-Par binding interactions can occur in the absence of the other segregation and transfer components, we co-produced the proteins in a pairwise manner. As shown in Figure 6, TraI, TraD, and TraC were again co-purified with ParM_C−FLAG. Retention of low amounts of TraI by ParR was confirmed. No specific interaction with TraD or TraC was detected. Relaxase accessory factor TraM binds oriT, TraD, and the membrane (Schwab et al., 1991; Disque-Kochem and Dreiseikelmann, 1997; Lu and Frost, 2005), but antibodies to TraM did not detect co-retention of this protein with either ParM or ParR (not shown). The abundance of Par protein present in each cell extract and in the pull down fractions was quantitated using antibodies to the FLAG epitope (Figure S3). We conclude that ParR binds TraI less strongly than ParM. Moreover ParM protein in the cell binds not only TraI, but additionally the T4CP TraD and TraC. In every case, these interactions take place in the absence of an intact transfer machinery and filament formation. There is a possibility that ParM is bound by additional Tra proteins of plasmid R1 but currently we do not have antibodies available for the entire suite of purified components.

To test whether the additional binding partners, TraD and TraC, alter ParM ATPase activity, the purified proteins were assayed in combination in vitro. Our published (Mihajlovic et al., 2009) and unpublished (V.K.H. Rajendra and E.L.Zechner) observations show that the soluble form of TraD (lacking the N-terminal transmembrane domain; TraDΔN130) and full length TraC interact with several protein and DNA ligands in vitro, yet neither Tra protein increased ATP hydrolysis compared to reactions containing ParM alone (not shown).

We next asked whether the reciprocal activity, namely the direct transfer of either Par protein by the T4SS to recipient cells, could be detected. The cre gene was fused 5′ to each par gene and translocation of ParM and ParR was analyzed by CRAfT. No Cre-ParR transfer was detected (Figure 7B). Remarkably, however, Cre-ParM translocation was measured. The observed frequency was low compared to Cre-TraI transfer. To address whether Cre-ParM transfer is the result of specific recognition, we tested ParM variants with amino acid exchanges in residues exposed on outside loops and along the surface of ParM filaments (Salje and Lowe, 2008). These residues are important to ParR binding, and crucial to stable filament formation. Both mutant variants were transferred, but CreParMK123A and CreParMS39A secretion was significantly less efficient (24.5 and 59% of wild type levels, respectively). The impact of single residue exchanges on transfer efficiency strengthens the evidence for specific ParM recognition by the T4CP. We then asked whether the highly related conjugation system of plasmid F would also mediate transfer of ParM, despite the absence of parMRC on that plasmid. For this test CRAfT assays were performed with pOX38 (Figure 7C). Although Cre fused to F TraI was efficiently secreted, we measured no Cre-ParM transfer. We conclude that translocation of ParM protein is unique for plasmid R1-16.

All enzymes were used according to Manufacturer's recommendations.

Oligonucleotides are shown in Table 3.

Inserts for pMS_parM and derivatives were amplified with parMBamHIfw/parMEcoRIrev, for pMS_parR with parRBamHIfw/parREcoRIrev using pMS_parA, pJSC1-S39A, or pJSC1-K123A as templates. Amplicons were cut with BamHI/EcoRI and ligated with pMS119HE. N-terminal Cre fusions were constructed by inserting amplicons from templates pMS_parM, pMS_parR, or derivatives in CFB B Sm plasmid via KpnI/SalI. Primers for the parM inserts: ParM_SFW/ParM_SRev; for parR insert: ParR_FW/ParR_Rev. To generate the C-terminal CreParR fusion plasmid parR was amplified with parR_NheIFW/parR_NheIRev from template pMS_parA, and ligated to NheI cut CFP B. The parM PCR fragment from parMNdeI_fw/parMBamHI_rev and R1-16 template were cut with NdeI and BamHI and ligated to yield pet3A-parM. pASKIBA7PLUSTraC was generated by amplification of traC from R1-16 with primers StrepTraCEcorIFw and StrepTraCHindIIIR, cutting with EcoRI and HindIII and ligation in pASKIBA7PLUS.

pGZtraD was created by amplification of traD from R1-16 with primers SS01 and SS02, EcoRI and HindIII treated and inserted in pGZ119EH. To generate pGZtraC, traC was cut from R1-16 with EcoR1 and SmaI and ligated into pGZ119EH. pMS-CFLAGparM was constructed by amplification of parM with parM_CFLAG_EcoRI_rev/parMBamHI_fw, cutting with EcoRI and BamHI and ligation into pMS119EH. pMS-NFLAGparR and pMS-CLFAGparR were constructed by amplification of parR with primers parR_NFLAG_BamHI and parREcoRIrev or parRBamHIfw and parR_CFLAG_EcoRI_rev, respectively, restriction with BamHI/EcoRI and ligation with pMS119EH. pBlueparC was constructed by amplification of parC from R1-16 with primers parMRCFW and parCrev, restriction with EcoRI and ligation with pBluescript II KS(−).

To generate R1-16ΔparM, primers ParMloxFW/ParMloxRev were used to amplify a loxP-TetRA-loxP cassette from E. coli CSH26Cm::LTL. For R1-16ΔparR, FW_parR_TetRA/Rev_parR_TetRA were used to amplify a tetracycline resistance cassette from pAR183 (Reisner et al., 2006). For R1-16ΔparM R, FW_R1parM/Rev_R1parR were used to amplify a streptomycin resistance cassette from pAH144 (Haldimann and Wanner, 2001). The amplified fragments were introduced into E. coli DY330 [R1-16] and integrated via homologous recombination (Reisner et al., 2002). Introduction of the CFP B plasmid into strains carrying R1-16 mutants catalyzed a Cre/loxP mediated recombination reaction excising the tetRA cassette.

Alexa488 labeled R17 phage was prepared as described (Lang et al., 2014). E. coli MS411 carrying R1-derivatives with and without complementation plasmids were grown to an A₆₀₀ of 0.6–0.8, diluted in PBS to an A₆₀₀ of 0.5 and incubated with 0.01 vol. R17 phage for 10 min at RT. Five microliters were mounted on a glass slide, pictures were taken with an Eclipse Ti fluorescence microscope (Nikon).

For quantification of apparent copy number, plasmid yields of R1 derivatives (R1-16, R1-16::B4, R1-16miniTn5 E5, or R1-16miniTn5 B4) were determined and compared to the yields of an independent replicon (pGZ119EH) as described in Nuk et al. (2011). E. coli M1174 carrying the desired plasmid combinations were grown in 5 ml LB medium with antibiotics. One millimolar IPTG induced traI (pHP2) expression.

E. coli MS411 carrying the plasmids were grown overnight in LB media with antibiotics at 37°C. Hundred microliters of donor cells were centrifuged for 3 min at 3000 × g, resuspended in 1 ml 0.9% NaCl, subcultured in drug free LB for 1 h at 37°C and adjusted to A₆₀₀ 0.02. A 10-fold excess of recipient MS614 was added and the mixture was incubated for 3, 5, 15, or 30 min at 37°C without shaking. DNA transfer was stopped by vortexing for 1 min and rapid cooling on ice. Donors were selected on LB-plates with antibiotics (see Table 2) and transconjugants were selected on kanamycin (40 μg/ml) and streptomycin (25 μg/ml). Conjugation frequencies were calculated as transconjugants per donor.

CRAfT was performed as described previously (Lang et al., 2010). E. coli MS411 carrying the plasmids of interest and recipient CSH26Cm::LTL were used. Donors were selected on plates with antibiotics (Table 2). Transconjugants and recombinants were identified by plating serial dilutions on LB-Kan/Tc or chloramphenicol, respectively. Conjugation and protein translocation frequencies are calculated as transconjugants or recombinants per donor, respectively.

TraI, TraIN309, TraIN992, TraDΔN130 were purified as described (Csitkovits et al., 2004; Mihajlovic et al., 2009; Sut et al., 2009; Lang et al., 2011).

ParR purification: 2 l E. coli C41(DE3) [pJSC21] were grown in LB with 100 μg/ml ampicillin (LB-Amp) at 37°C with shaking to an OD600 of 0.6. One millimolar IPTG was used for induction. After 6 h, cells were harvested (10 min, 4000 × g, 4°C), resuspended in 20 ml buffer I (50 mM Tris-HCl pH 7, 100 mM KCl, 1 mM EDTA, 1 mM DTT, 0.001% PMSF) with proteinase inhibitor cocktail (cOmplete EDTA-free, Roche) and frozen at −80°C. Cells were lysed, incubated with DNAse I (AppliChem) for 20 min on ice and centrifuged (140,000 × g, 1 h, 4°C). The cytoplasmic fraction was filtered through a 0.45 μm PVDF syringe filter and loaded (1 ml/min) on HiTrap Heparin columns equilibrated with buffer I. After washing (2 column volumes buffer I), bound protein was eluted with buffer I and a linear gradient of 0–1 M NaCl. Partially pure ParR eluted at ~300–450 mM NaCl. These fractions were pooled and dialysed overnight against 100 × vol. buffer I. Dialyzed fractions were loaded (1 ml/min) on 2 serially connected 5 ml HiTrap SP HP columns equilibrated with buffer I. After washing (2 ml/min) with 10 ml of buffer I, protein was eluted with a linear gradient of buffer I + 0–1 M NaCl. ParR eluted at ~450–500 mM NaCl. Purity was confirmed by SDS-PAGE and fractions were pooled and dialyzed overnight [100 vol. buffer II (20 mM Tris-HCl pH 9, 50 mM KCl, 1 mM EDTA, 1 mM NaN₃)]. ParR was concentrated, adjusted to 20% glycerol and frozen at −80°C.

E. coli BL21 C41 (DE3) carrying pASKIBA7PLUSTraC were grown in 1l LB-Amp at 37°C to an OD₆₀₀ of 0.5. Overexpression was induced by addition of anhydrotetracyclin (AHT, 0.2 mg/l). Cells were harvested after 4 h shaking at 37°C, pellets were frozen at −80°C. Pellet was resuspended in 20 ml of buffer I (50 mM Tris-Cl pH 7.7, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, 1 tablet protease inhibitor (cOmplete, Roche) and lysed. The lysate was centrifuged for 45 min at 50,000 × g at 4°C. The supernatant was filtered (0.45 μm PVDF syringe filters) and loaded onto a Hitrap strep HP column pre-equilibrated with buffer I. After washing (5–10 column volumes) with buffer I, TraC was eluted with 30 ml buffer II (buffer I containing 2.5 mM desthiobiotin) in one step. Fractions containing TraC were pooled and concentrated (Amicon centrifugal filter, Millipore) and loaded onto a Hiload 16/60 Superdex 200 column. The protein was eluted with buffer III (25 mM Tris pH 7.6, 100 mM NaCl, 1 mM DTT, 1 mM MgCl₂, 1 mM PMSF). Pure TraC fractions were pooled, concentrated and frozen at −80°C (with 20% glycerol). Identity of TraC was confirmed by mass spectrometry and apparent molecular mass 99 kDa was determined (SDS-PAGE and Coomassie staining).

oriT specific cleavage activity was determined as described in Csitkovits et al. (2004). Indicated ParM and ParR concentrations were combined with 4 nM of pDE100 or pDE110 (negative control) independently or in combination, then the cleavage reaction was started by TraI (25 nM), TraIN308 (300 nM), or TraIN992 (100 nM). Statistical significance was determined using maximum stimulatory concentrations of ParM (15 nM) and ParR (15 nM) and 500 nM BSA as a negative control.

Construction of heteroduplex-substrates and the assay conditions were as described in Csitkovits et al. (2004) and Sut et al. (2009). Each heteroduplex substrate G2028 or IR (1 nM) was combined with effector proteins ParM and ParR independently or in combination in concentrations that were most stimulatory for TraI in the relaxase assay. Twenty-five nanomolar TraI was added to start the reaction. Resolution and quantitation of unwound DNA product were as described (Sut et al., 2009).

Enzyme activities were measured with the Malachite Green Assay Kit (Bioassay Systems). Briefly, different ParM concentrations (0–1 mM) with and without ParR (9 mM) and parC (pBlueparC, 17 nM) or DNA not containing parC (pDE110, 17 nM) were incubated (30 mM Tris-HCl pH 7.5, 50 mM KCl, 0.2 mM MgCl₂, 1 mM DTT, 0.1 mg/ml bovine serum albumin, 0.1 mM ATP; Jensen and Gerdes, 1997) at 30°C in a total volume of 25 μl. After 10 min, the reaction was stopped and color development after 30 min at RT was recorded at 595 nm. Stimulation of ParM ATPase activity by TraI, TraD, or TraC: 0.5 or 1 mM ParM were titrated with TraI, TraIN308, or TraIN992 (10–100 nM), TraD (20–500 nM), or TraC (0.5–8 mM), respectively, in the reaction buffer described above.

Impact of Par-Proteins on TraI ATP-hydrolysis: 1 fmol M13mp18 single-stranded phage DNA (New England BioLabs) was preincubated with ParR (0.5 mM) and ParM (9 mM) in buffer containing ATP (25 mM Tris HCl pH 7.5, 20 mM NaCl, 3 mM MgCl₂, 5 mM DTT, 2 mM ATP). TraI addition (0–10 nM) started the reaction. Basal TraI ATPase activity was 225,892 ± 83,485 mol/h/mol.

Oligos for fluorescence studies were reconstituted in 1 × STE (10 mM Tris pH 8.0, 50 mM NaCl, 1 mM EDTA). To create dsDNA substrates 3′-TAMRA labeled oligos were mixed in a 1:1 ratio with the unlabelled complementary strand, heated for 10 min, 96°C and re-hybridized at RT for 1 h. DNA binding by ParR was tested with ssDNA and dsDNA substrates. The consensus sequence of parC (Moller-Jensen et al., 2003) was created with Weblogo 3.4 (Crooks et al., 2004), a random DNA sequence was created with SMS (Stothard, 2000). Briefly, ds- or ssDNA parC^*, oriT^*, and randomseq^* (all 20 nM) were titrated with ParR in a total volume of 15 μl in 10 mM Tris HCl pH7.5, 50 mM NaCl, 0.005% NONIDET P-40, 1 mM DTT, 1 mM EDTA. The reaction was incubated for 20 min at 25°C, then 2 μl 87% glycerol were added. The products were resolved on 8% PAGE-gel without SDS at 15 V/cm for 30 min. Gels were scanned with a Typhoon 9400.

TraI affinity for 3′-TAMRA oriT DNA was measured on an ATF-105 fluorometer (Aviv Biomedical, Inc., Lakewood, NJ) as described in Dostal and Schildbach (2010). Briefly, 4 nM substrates (oriT17^*) with and without unlabelled competitor (50 nM 2 × G144C, Dostal et al., 2011) were preincubated with protein (ParR 10 nM, ParM 10 nM, 20 nM) at RT for 10 min in binding buffer (20 mM TrisHCl pH 7.5, 100 mM NaCl, 1 mM EDTA), the reaction was started with 0–100 nM TraI with a Microlab 500 titrator (Hamilton). A constant temperature of 25°C was maintained. Excitation wavelength was 520 nm, change in fluorescence intensity was followed at 580 nm. Equilibration time between each titration step was 3 min with mixing. Datapoints were averaged over 10 s. Volume correction and data fitting was done as described (Dostal and Schildbach, 2010).

The protocol was adapted from the TrIP assay (Cascales, Christie 2004). Hundred milliliters LB was inoculated with ONCs of E. coli MS411 strains carrying plasmid combinations. Thirty OD of exponentially growing (LB with antibiotics, 1 mM IPTG) cells were pelleted and resuspended in 45 ml 20 mM sodium phosphate buffer (pH = 6.8) with 1% formaldehyde and incubated for 15 min at RT. Five milliliters glycine (1.2 M in 20 mM sodium phosphate buffer pH = 6.8) were added, cells were pelleted and washed with 50 ml/30 OD of buffer A (50 mM Tris-HCl pH 6.8, 100 mM NaCl). For lysis, pellet was resuspended in 1 ml buffer B (10 mM Tris-HCl pH 8.0, 10 mM MgCl₂, 1 mg/ml lysozyme) and transferred to a 2 ml Eppendorf tube and frozen for 2 min with liquid N2, thawed on ice, frozen and thawed again. Each sample was sonicated for 10 s on ice. One, two milliliters of buffer C was added and adjusted with Triton X-100 to a final concentration of 4% and the lysate was incubated for 15 min with rotation at RT. Two hundred and thirty microliters cOmplete EDTA-free, Roche (1 tablet in 1 ml 25 mM MgCl2) was added and the mixture was incubated for 30 min at 37°C with shaking. 6.4 ml of buffer C (150 mM Tris-Hcl pH 8, 0.5 M sucrose, 10 mM EDTA) were added and insoluble material was removed by centrifugation (14,000 × g, 15 min). At this point, 200 μl aliquots of the supernatant were saved and stored at −80°C (whole cell lysate fraction, WCL). Remaining supernatant was transferred to tubes with Anti-FLAG affinity gel (90 μl/sample, A2220, Sigma) and incubated over night at 4°C. Beads were pelleted at 5000 × g, supernatant was removed. Beads were washed once with 950 μl buffer C supplemented with 1% Triton X-100 and twice with 950 μl buffer C supplemented with 0.1% Triton X-100. Immunoprecipitates (IP) were eluted by incubation of the beads for 30 min with FLAG-peptide (F3290, Sigma) at RT [80 μl FLAG-peptide (1 mg/ml in buffer C)/40 μl beads]. Beads were pelleted and the supernatant was collected (IP fraction).

A₆₀₀ 0.015–0.03 equivalents of the WCL and IP fractions were mixed with sample buffer containing DTT and SDS and resolved on SDS-PAGE (10%). Gels were blotted for 1.5 h on PVDF membranes. Blocking was done for 2 h (3% milk in TST). Detection of TraI and TraD -proteins was done with rabbit-antisera and α-rabbit HRP-conjugated antibody (7074S, Cellsignalling). Affinity purified TraC antibodies raised against TraC were produced by immunoGlobe. FLAG-tagged proteins were detected by HRP-conjugated α-FLAG antibody (A8592, Sigma). After washing (3 × 5 min) with 1 × TST, secondary antibody was incubated for 1 h. Blot development was done with ECL (Clarity Western, Bio-Rad).

ImageJ (Schneider et al., 2012) was used to quantify bands in gel-electrophoresis assays and SigmaPlot 12.2.0.45 and Qtiplot 0.9.8.3.3 were used to plot data.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.