Use of Chimeric Type IV Secretion Systems to Define Contributions of Outer Membrane Subassemblies for Contact-Dependent Translocation

SUMMARY

Recent studies have shown that conjugation systems of Gram-negative bacteria are composed of distinct inner and outer membrane core complexes (IMCs and OMCCs, respectively). Here, we functionally characterized the OMCC, focusing first on a cap domain that forms a channel across the outer membrane. Strikingly, the OMCC caps of the Escherichia coli pKM101 Tra and Agrobacterium tumefaciens VirB/VirD4 systems are completely dispensable for substrate transfer, but required for formation of conjugative pili. The pKM101 OMCC cap and extended pilus also are dispensable for activation of a Pseudomonas aeruginosa type VI secretion system (T6SS). Chimeric conjugation systems composed of the IMC_pKM101 joined to OMCCs from the A. tumefaciens VirB/VirD4, E. coli R388 Trw, and Bordetella pertussis Ptl systems support conjugative DNA transfer in E. coli and trigger P. aeruginosa T6SS killing, but not pilus production. The A. tumefaciens VirB/VirD4 OMCC, solved by transmission electron microscopy, adopts a cage structure similar to the pKM101 OMCC. Our findings establish that OMCCs are highly structurally and functionally conserved - but also intrinsically conformationally flexible - scaffolds for translocation channels. Furthermore, the OMCC cap and a pilus tip protein coregulate pilus extension but are not required for channel assembly or function.

Graphical Abstract

INTRODUCTION

The bacterial type IV secretion systems (T4SSs) are a highly versatile superfamily of macromolecular transporters (Costa et al., 2015, Christie, 2016). An early review highlighted this versatility by comparing three members of a subfamily of the T4SSs now termed the P-type or type IVa systems: i) the Escherichia coli pKM101 Tra system, which mediates conjugative DNA transfer between bacteria, ii) the Agrobacterium tumefaciens VirB/VirD4 system responsible for delivery of oncogenic T-DNA and effector proteins to plant cells, and iii) the Bordetella pertussis Ptl system that drives export of the multisubunit pertussis toxin (PT) across the B. pertussis outer membrane to the milieu (Winans et al., 1996). Each of these systems shares a common ancestry, subunit composition and, most likely, overall architecture (Christie & Vogel, 2000, Christie, 2016, Chandran Darbari & Waksman, 2015). In view of these commonalities, it is intriguing to ask how each of these systems was adapted during evolution to achieve such striking functional diversity.

Recent studies have provided a structural blueprint for the type IVa subfamily. The E. coli pKM101 Tra (designated Tra_pKM101) and closely related R388-encoded Trw (Trw_R388) conjugation machines are composed of large (~1 Megadalton; MDa), similarly cage-shaped outer membrane (OM) core complexes (the OMCC) (Fronzes et al., 2009, Chandran et al., 2009, Low et al., 2014). These OMCCs are composed of 14 copies each of VirB7- and VirB9-like subunits and of the C-terminal halves of VirB10-like subunits (we use here the A. tumefaciens VirB/VirD4 system, VirB/VirD4_At, as a unifying nomenclature). The N-terminal halves of the VirB10 subunits extend to the inner membrane, where they form part of an even larger inner membrane complex (IMC; 2.5 MDa). Besides this portion of VirB10, the IMC is composed of 24 copies of VirB6 homologs, and 12 copies each of VirB3, VirB4 ATPase, VirB5 and VirB8 homologs (Low et al., 2014). Missing from the Trw_R388 structure, which is designated as T4SS_3–10 because it is composed of the VirB3 through VirB10 homologs, are VirB2-like pilins, the conjugative pilus, and the VirD4- and VirB11-like ATPases. Recently, however, structural studies established that the F plasmid-encoded conjugative pilus is composed of a 5-start helical assembly. Strikingly, the F pilus consists of a stoichiometric complex of the TraA pilin subunit bound to a phospholipid (Costa et al., 2016), but it remains unknown how the pilus is mounted onto the T4SS IMC and OMCC.

At the distal end of the OMCC is an interesting feature termed the cap domain. An X-ray structure of the outer layer (O-layer) of the OMCC from Tra_pKM101 revealed that this cap domain is composed of a helix-loop-helix extension of VirB10, termed the antennae projection (AP). The AP helices from the 14 TraF monomers come together to form the cap with a diameter of ~100 Å and a central channel of ~10 – 30 Å (Fronzes et al., 2009, Chandran et al., 2009). The cap domain spans the OM where it is envisioned to participate in various surface phenomena including pilus biogenesis, substrate transfer across the OM, and formation of target cell contacts.

Here, we report results of mutational and domain swapping studies of the Tra_pKM101 and VirB/VirD4_At model systems establishing the requirement for the OMCC cap for assembly of the conjugative pilus, but not productive mating junctions or even contact-dependent activation of a type VI secretion system (T6SS). We also show that chimeric T4SSs composed of the IMC_pKM101 coupled to heterologous OMCCs from the Trw_R388, VirB/VirD4_At, and Ptl_Bp systems support conjugative DNA transfer in E. coli and activate T6SS killing, but fail to elaborate detectable pili. Finally, we solved the structure of the A. tumefaciens VirB/VirD4 OMCC by transmission electron microscopy and negative staining, enabling structural comparisons with the other solved OMCCs. We discuss our findings in the context of a model for conjugation machines functioning in Gram-negative species in which the distal end of the OMCC, together with a pilus tip protein, coordinates a late-stage morphogenetic switch that alternatively directs pilus extension or intercellular substrate transfer.

RESULTS

Genetic requirements for assembly of the pKM101 Tra T4SS

We initiated studies of the Tra_pKM101 T4SS by constructing a set of individual tra gene deletion mutations to define genetic requirements for machine assembly (Fig. S1). Eight of the 11 tra genes were precisely deleted from pKM101 by recombineering, but for unknown reasons we were unable to delete traL, traN, and traO. We therefore created miniaturized versions of pKM101 consisting of the entire tra gene cluster and upstream regulatory sequences introduced into ColE1 or pACYC184 plasmid vectors or joined to pKM101’s oriV replication region along with a selectable Kan^r gene. These mini-pKM101 plasmids encode fully functional Tra T4SSs (see below) and served as templates for further genetic manipulations of the tra cluster, including the construction of a complete set of Δtra mutations.

E. coli donors conjugatively transfer pKM101 at frequencies approaching 1 transconjugant per donor (Tc/D) in 2 h matings on solid-surfaces (Winans & Walker, 1985). In contrast to previous reports that IncN plasmids typically transfer poorly in liquid matings (Bradley et al., 1980, Jorgensen & Stenderup, 1982), MG1655(pKM101) cells delivered the plasmid to recipients at frequencies of approximately 5 × 10⁻⁴ and 10⁻¹ Tc’s/D in 2 h and overnight liquid matings (under constant agitation), respectively (Fig. S1). A donor carrying mini-pKM101 pRP100 mobilized transfer of pJG142, a plasmid that carries the entire traK/traJ/traI/oriT mobilization (mob) region from pKM101 (Paterson et al., 1999), at frequencies slightly higher than observed for pKM101 in both solid surface and liquid matings (Fig. 1).

FIG. 1.

E. coli pKM101 tra gene deletion and complementation analyses. The E. coli pKM101 tra and A. tumefaciens virB loci are similar in gene composition and order, as shown by color-coding of genes encoding homologs of the T4SS subunits. The pKM101 tra genes expressed from pRP100 encode a fully functional Tra T4SS, as shown by efficient conjugative DNA transfer and IKe bacteriophage sensitivity. The schematic depicts effects of individual Δtra mutations (histogram, upper bars) and results of complementation studies (histogram, lower bars) in which corresponding genes were trans-expressed from the P_BAD promoter (black arrow) on conjugative transfer. Matings (2 h) were carried out on solid-surface (solid bars) and in liquid (stippled bars); pRP100-carrying donors also were mated overnight in liquid with constant agitation (light stippled bars). *, denotes transfer frequencies below the threshold of detection (<10⁻⁸ Tc’s/D). Transfer frequencies are presented as transconjugants/donor (Tc’s/D). IKe phage sensitivity (S, sensitive, R, resistant) for Δtra mutants and complemented strains is shown at the right.

Most of the Δtra mutations abolished Tra T4SS function (Figs. 1 & S1), in agreement with previous findings for the A. tumefaciens VirB/VirD4 T4SS and the E. coli Trw_R388 conjugation system (Berger & Christie, 1994, Larrea et al., 2013). Furthermore, strains harboring the pKM101Δtra or pRP100Δtra mutant plasmids were complemented by trans-expression of the corresponding tra gene from the P_BAD promoter (Figs. 1 &S1). In general, complementation did not fully restore plasmid transfer to wild-type (WT) levels, possibly due to negative effects accompanying nonstoichiometric production on machine assembly or slight polar effects of the deletion mutations on downstream tra gene expression. Regardless, results of the complementation studies confirmed that 9 of the 11 Tra proteins are essential for elaboration of a functional Tra_pKM101 T4SS.

virB1-like traL codes for a peptidoglycan (PG) hydrolase and its deletion had no discernible effect on plasmid transfer in either solid-surface or liquid matings (Fig. 1). This finding contrasts with previous reports documenting attenuating effects of PG hydrolase deletions on substrate transfer through related T4SSs (Berger & Christie, 1994, Bayer et al., 1995, Zahrl et al., 2005). Complementation of the ΔtraL mutation by a P_BAD::traL expression plasmid conferred an apparent ^~10-fold increase in transfer frequencies (Fig. 1). These cells, however, exhibited growth defects and enhanced cell lysis compared with cells expressing traL from the native promoter (data not shown), as observed previously for the P19 hydrolase associated with the plasmid R1-encoded conjugation system (Bayer et al., 2001). The apparent increase in mating frequency (in Tc’s/D) is therefore likely due to a reduction in donor cell viability accompanying TraL overproduction.

The last tra gene, virB5-like traC, is postulated to encode a pilus tip protein by extrapolation of results presented for homologs in the VirB/VirD4_At and Helicobacter pylori Cag T4SSs (Aly & Baron, 2007, Shaffer et al., 2011). Interestingly, ΔtraC mutant donors transferred DNA substrates at frequencies of 5 × 10⁻⁵ Tc’s/D on solid surfaces, but were completely defective for plasmid transfer in liquid matings (Figs 1& S1). In the F plasmid transfer system, F pili extend to initiate distant contacts and then retract to promote formation of mating junctions, and it is thought that these dynamic rounds of extension and retraction account for the observed high-frequency transfer of F plasmids in liquid matings (Clarke et al., 2008). There is no evidence that the Tra_pKM101 pilus dynamically extends and retracts, yet its production might still facilitate target cell contacts enabling fairly efficient transfer in liquid. pKM101-encoded pili are difficult to visualize microscopically, but pilus production can be assessed by susceptibility of plasmid-carrying cells to IKe, an M13-like filamentous phage that uses the pKM101 pilus as a receptor (Bradley, 1979). Among the Δtra strains, only the ΔtraL mutant exhibited IKe sensitivity suggesting that TraL is dispensable for pilus production. TraC, however, was required for IKe infection and plasmid transfer in liquid - but not solid-surface – matings. These findings support the notion that TraC plays an important role in pilus biogenesis but not elaboration of the translocation channel.

The outer membrane core complex (OMCC) cap is dispensable for substrate transfer but required for pilus biogenesis

With a pKM101 molecular ‘toolkit’ in hand, we sought to define the T4SS machine requirements for productive engagement with recipient cells. Many T4SSs of Gram-negative bacteria elaborate conjugative pili, which clearly play a role in initiating donor-target cell interactions. It is noteworthy, however, that mutations have been isolated that block detectable pilus production (Pil⁻) without affecting substrate transfer (Tra⁺) (Sagulenko et al., 2001, Jakubowski et al., 2003, Jakubowski et al., 2005, Haase et al., 1995). In the absence of extended pili, these systems must therefore elaborate a surface feature capable of promoting donor-target cell contacts and formation of productive mating junctions.

We sought to define the functional importance of the OMCC cap, which to date is the only identified surface-exposed domain of the type IVa T4SSs other than the pilus. As mentioned above, this cap is assembled from the 14 AP domains of VirB10-like subunits. In the Tra_pKM101 O-layer structure, the α2 and α3 helices of the AP domains of TraF form the OM-spanning channel while the intervening AP loops (APLs) project from the cell surface (Fig. 2A) (Chandran et al., 2009). We introduced deletion or substitution mutations in the AP of TraF_pKM101, which spans residues 307–355. Interestingly, substitution of the APL with 5 Gly residues (5xGly) or introduction of a FLAG epitope in TraF’s APL had no detectable effects on TraF protein accumulation (Fig. 2C) or donor-directed plasmid transfer (Fig. 2B). Deletion of the entire AP domain also did not affect TraF abundance and conferred only a modest reduction in DNA transfer in solid-surface matings compared with donors producing native TraF (Figs. 2B,C). The ΔAP mutant donors, however, failed to transfer the plasmid substrate at detectable levels in liquid matings (see below).

FIG. 2.

Substitution and deletion mutational analysis of the outer membrane cap of the pKM101 Tra T4SS. A) Ribbon diagram of the O-layer of the pKM101 outer membrane core complex (OMCC). VirB7-like TraN and VirB9-like TraO are color-coded magenta and cyan, respectively. The α-helical antennae projection (AP) forming the OM-spanning cap and the C-terminal (CT) domain of TraF are color-coded red, and the β-barrel domain of TraF is color-coded yellow. At right, ribbon diagram of a TraF monomer depicting the β-barrel, AP, and CT domains in same color-coding. Domain junctions (residues from N terminus) and positions of deletion or substitution mutations are indicated. B) Schematic depicts TraF domain architecture with junctions (in residues) indicated. Mutations in the AP or CT are listed at left, and effects of the mutations on plasmid transfer (transconjugants per donor, Tc’s/D) and IKe phage infection (S, sensitive; R, resistant). C) Levels of His-TraF and mutant proteins in total cell extracts, as monitored by immunostaining with α-His antibodies. RNA polymerase β-subunit (α-RNAP) served as a loading control.

To evaluate the generality of these findings, we inserted duplicate (2X) or triplicate (3X) FLAG tags at several positions within the AP domain of A. tumefaciens VirB10 (Fig. S2A). These insertions did not affect VirB10 protein accumulation (Fig. S2C) or function as deduced by the appearance of morphologically wild-type plant tumors resulting from A. tumefaciens-mediated delivery of oncogenic T-DNA into plant cells (Fig. S2B). The AP domains of TraF and VirB10 share only 26 % identity (Figs. 3A, S3), yet reciprocal swaps of these domains yielded stable (Figs. 3C, S2C) and fully functional TraF/AP_VirB10 and VirB10/AP_TraF chimeric proteins as shown by robust transfer of a pKM101 substrate in E. coli matings (Fig. 3B) and oncogenic T-DNA in A. tumefaciens infection assays (Fig. S2B). Together, these findings support a conclusion that the OMCC caps of the Tra_pKM101 and VirB/VirD4_At T4SSs enhance, but are not required for, substrate transfer to target cells.

FIG. 3.

Domain swapping reveals compositional flexibility of TraF’s β-barrel, antennae projection (AP,) and C terminus (CT). A) Sequence alignment of the AP and C-terminal (CT) domains of TraF and VirB10, with identical (red) and nonidentical (black) residues shown. Numbers correspond to domain junctions (residues from N terminus). Sequences comprising the α2 - loop (APL) - α3 regions of AP domains and the highly-conserved RDLF motifs are highlighted. B) Schematics depicting domains of TraF and VirB10, with junctions (residues from N terminus) indicated: Cyto, cytoplasmic; TM, transmembrane domain; Pro-Rich, proline-rich-region; β-Barrel; AP, antennae projection; CT, C-terminal domain. Schematics of the TraF/VirB10 chimeras depict the VirB10 domain(s) swapped for the equivalent domain(s) of TraF. Strains producing the TraF/VirB10 chimeras supported plasmid transfer in 2 h solid-surface matings at the frequencies shown in transconjugants per donor (Tc’s/D), and exhibited sensitivity (S) or resistance (R) to IKe infection. C) Levels of His-TraF and chimeric proteins in total cell extracts, as monitored by immunostaining with α-His antibodies. RNA polymerase β-subunit (α-RNAP) served as a loading control.

Complete deletions of the APs from TraF and VirB10, however, abolished pilus production, as evidenced in E. coli by a failure of the ΔAP mutant to transfer the pKM101 substrate in liquid and their resistance to IKe phage infection (Fig. 2B). In A. tumefaciens, the ΔAP mutant lacked detectable VirB2 pilin on the cell surface, which serves as a convenient assay for T pilus production by the VirB/VirD4_At T4SS (Figs. S2B) (Sagulenko et al., 2001, Kerr & Christie, 2010). Further mutational analyses established the importance of the AP membrane-spanning α-helices, but not the APLs, for pilus biogenesis. For example, strains producing TraF with a 5xGly replacement of the APL, or a FLAG insertion in this domain, transferred their plasmid substrates in liquid matings and were susceptible to infection by IKe (Fig. 2B). Similarly, A. tumefaciens strains producing VirB10 variants with FLAG insertions in the APL accumulated abundant amounts of VirB2 pilin on the cell surface, whereas strains with FLAG insertions in VirB10’s α2 or α3 helices had comparatively lower levels of surface pilin (Fig. S2B). It is also noteworthy that reciprocal swaps of TraF’s and VirB10’s AP domains supported pilus production in E. coli and A. tumefaciens (Fig. 3B, S2B). These findings suggest that the elaboration of an α-helical channel across the OM, regardless of its sequence composition, fulfills a minimal requirement for pilus production by the Tra_pKM101 and VirB/VirD4_At systems.

The C-terminal (CT) domain, but not the lever arm, is critical for TraF function

The C termini of TraF and VirB10 are highly related (72 % identity), and both domains possess an RDLDF motif that is also highly conserved among the VirB10 family members (Figs. 3A & S3C) (Jakubowski et al., 2009). As shown by the Tra_pKM101 O-layer structure, TraF’s C-terminal (CT) domain forms a β-strand that extends along the β-barrel domain (see Figs. 2A, S3) (Chandran et al., 2009). Deletions of TraF’s AP and CT domains or just the CT domain were strongly destabilizing, as evidenced by low abundance of the mutant proteins and accumulation of presumptive proteolytic breakdown products (Fig. 2C). Deletion of the RDLDF motif or of the 9 C-terminal residues did not affect TraF’s abundance, but completely eliminated function as monitored by substrate transfer and IKe phage infection (Fig. 2B, C). Next, we substituted VirB10_At’s AP and CT domains or just the CT domain for those of TraF. The respective TraF/AP-CT_VirB10 and TraF/CT_VirB10 chimeric proteins supported plasmid transfer on solid surfaces, but not transfer in liquid or IKe phage infection (Fig. 3B). Interestingly, VirB10’s CT domain extends 11 residues beyond that of TraF, and a chimera (TraF/CTΔ11_B10) deleted of this extension was stably produced and supported substrate transfer and IKe phage infection (Figs. 3B, C). In A. tumefaciens, a VirB10 chimera bearing TraF’s AP-CT (VirB10/AP-CT_TraF) also supported WT levels of substrate transfer to plants but not detectable T pilus production (Fig. S2). The CT domains thus promote stabilization of the VirB10 proteins and also mediate intra- or intersubunit contacts necessary for channel formation and pilus production. The 11 C-terminal residues of VirB10 are required for T pilus production by the VirB/VirD4_At T4SS, but appear to poison pilus production when appended to TraF in the Tra_pKM101 system. Interestingly, however, both TraF and VirB10 accommodate C-terminal epitope tags without effects on function (Fig. 2B, C, and see below).

In the Tra_pKM101 O-layer crystal structure, a domain of TraF designated as the lever arm extends laterally from one TraF monomer to form a network of contacts with 3 adjacent TraF monomers. This results in a tetradecameric complex in which the 14 lever arms form a continuous inner shelf at the base of the OMCC (Chandran et al., 2009). Notably, TraF’s CT domain, and more specifically β-strand β7c containing the RDLDF motif, interacts with β-strand β_n1 in the lever arm of an adjacent TraF monomer (Fig. 2, S3) (Chandran et al., 2009). To evaluate the functional importance of this putative CT domain - lever arm interaction, we constructed a variant of TraF deleted of the lever arm. In contrast to native TraF, the TraFΔlever mutant migrated in SDS-polyacrylamide gels as multiple, presumptive degradation products (Fig. S4A). Strikingly, however, donors producing the TraFΔlever mutant protein were proficient for DNA transfer and also exhibited IKe phage sensitivity (Fig. S4B). Thus, despite an apparent stabilizing effect of TraF’s lever arm, the network of lateral contacts forming the lever arm shelf at the base of the OMCC is dispensable for elaboration of the translocation channel and pilus.

TraF chimeras with substituted OM core complexes support substrate transfer

VirB10 subunits are unique among known Gram-negative bacterial proteins in spanning the entire cell envelope (Jakubowski et al., 2009, Chandran et al., 2009). To determine if TraF could tolerate substitutions of domains other than the AP and CT, we constructed additional TraF/VirB10 chimeras (Fig. 3B). Chimeras in which VirB10’s cytoplasmic (Cyto), transmembrane (TM), or periplasmic Proline-Rich Region (PRR) were substituted for the equivalent regions of TraF accumulated at detectable levels but failed to support substrate transfer or pilus production (Figs. 3B, C). These findings are in general agreement with results of previous studies showing that the N-proximal regions of the VirB10-like proteins form extensive interactions with cognate IMC components and VirD4 coupling proteins (Das & Xie, 2000, Atmakuri et al., 2004, Llosa et al., 2003, Rivera-Calzada et al., 2013).

By contrast, chimeras consisting of TraF’s N-terminal half joined to VirB10’s β-barrel domain with or without swaps of the AP and CT domains accumulated at low levels yet supported substrate transfer at frequencies of 10⁻⁵ – 10⁻⁶ Tc’s/D (Fig. 3B). The functionality of these chimeras was particularly striking in view of the low sequence identities of TraF and VirB10 across these domains (~19 %; Figs. 3A, S2) and the Tra_pKM101 O-layer X-ray structure, which shows that TraF’s β-barrel forms extensive contacts with its partner subunit TraO (Fig. S3C) (Chandran et al., 2009). The OMCCs are intrinsically stable subassemblies (Fronzes et al., 2009), prompting a test of whether the TraF/βB-CT_VirB10 chimera would function more efficiently if the VirB7 and VirB9 subunits of the VirB/VirD4_At OMCC also were substituted for their Tra_pKM101 counterparts, essentially creating an IMC_Tra::OMCC_VirB chimera (designated Tra::VirB; Fig. 4A). To ensure temporal and stoichiometric synthesis of the IMC and OMCC subassemblies, we substituted codon-optimized virB7, virB9 and the traF/βB-CT_B10 chimera for traN, traO, and traF within the tra region of the functional mini-pKM101 plasmid pCGR108 (see Fig. 4B). Very interestingly, donors producing the Tra::VirB chimeric T4SS transferred the plasmid substrate in solid-surface matings, although at frequencies comparable to donors producing the TraF/βB-CT_VirB10 chimeric protein (compare Figs. 3B & 4B). The Tra::VirB-producing donors were transfer deficient in liquid matings and were insensitive to IKe phage infection, indicative of a lack of pilus production.

FIG. 4.

Chimeric T4SSs support conjugative DNA transfer and activate T6SS killing. A) Sequences encoding the outer membrane core complex (OMCC) subunits TraN, TraO, and the C-terminal half (residues 194–386) of TraF were replaced with corresponding genes or gene fragments from the A. tumefaciens VirB, E. coli R388 Trw, or B. pertussis Ptl systems on mini-pKM101 plasmid pCGR108. The chimeric T4SSs composed of the inner membrane complex (IMC) of pKM101 (yellow) joined to the OMCCs from the VirB, Trw, or Ptl systems (color-coded) are modeled on the R388 T4SS_3–10 structure (Low et al., 2014). B) E. coli donors carrying pCGR108 derivatives encoding the only the Tra_pKM101 IMC or OMCC, or the IMC::OMCC chimeras transferred the mobilizable plasmid pJG42 at the frequencies shown in transconjugants per donor (Tc’s/D) in solid-surface (histogram, solid bars) or liquid (stippled bars) matings, and were resistant to IKe phage infection (S, sensitive; R, resistant). C) E. coli survival when cultivated in the absence or presence of P. aeruginosa PAO1. E. coli DH5α cells lacked or produced intact or variant forms of the Tra_pKM101 T4SS depicted. Statistical significance is shown based on a Student’s t test corresponding to the values of plasmid-free DH5α or growth in the absence of P. aeruginosa (NS, not significant; *P < 0.05; **P < 0.01). For panels B and C, data presented are mean +/− SD, n = 3 independent replicates.

We also engineered two more chimeric T4SSs, the first bearing an OMCC from the E. coli Trw_R388 conjugation machine. The Trw_R388 system is closely related phylogenetically and functionally to the Tra_pKM101, and evidence has been presented for the interchangeability of some constituent subunits between these systems (Llosa et al., 2003, de Paz et al., 2005). As might be expected, the chimera composed of pKM101’s IMC and the OMCC from Trw_R388 supported transfer of the pKM101 substrate at a moderately high frequency of 10⁻³ Tc’s/D (Fig. 4B). The Ptl_Bp T4SS, by contrast, is highly divergent from the E. coli conjugation systems both in primary sequence (Fig. S3B) and in its function as a contact-independent PT export system (Locht et al., 2011). Interestingly, however, the Tra::Ptl chimera also supported transfer albeit at a lower level (^~10⁻⁷ Tc’s/D) than either the Tra::VirB or Tra::Trw systems. (Fig. 4B). Together, these findings confirm the modular and interchangeable nature of OMCCs from phylogenetically diverse type IVa systems.

T4SS requirements for triggering of Type VI-mediated killing

One striking outcome from the above studies is that OMCCs from the A. tumefaciens and B. pertussis systems mediate formation of productive mating junctions with E. coli recipients, in spite of the fact that the former delivers substrates to plant cells and the latter functions as a contact-independent toxin exporter. These findings suggest that surface-exposed structures elaborated by the T4SSs are capable of directing the formation of cell contacts with diverse target cells even in the absence of pilus production or possibly even substrate transfer. We sought to further define the nature of T4SS surface structures mediating cell-cell contacts, and to this end we employed a T6SS killing assay. It was shown previously that pKM101-carrying E. coli cells convey a signal across the P. aeruginosa cell envelope that triggers production of the H1-T6SS. In turn, P. aeruginosa cells kill the activating E. coli cells through transfer of toxic effectors (Ho et al., 2013). To identify T4SS surface features responsible for propagating the contact-dependent signal, we incubated E. coli donors producing WT or mutant T4SSs with P. aeruginosa strain PAO-1 and assayed for T6SS-mediating killing by serial dilution on media selective for E. coli.

E. coli cells lacking pKM101 exhibit comparable growth in the presence or absence of P. aeruginosa (Fig. 4C), whereas pKM101-carrying cells exhibit a ~2-log reduction in colony forming units (CFUs) when incubated in the presence vs the absence of P. aeruginosa PAO-1 or the presence of a T6SS⁻ (vipA) mutant (Fig. 4C) (Ho et al., 2013). Strains separately producing the IMC_pKM101 or OMCC _pKM101 subassemblies were not killed in the presence of P. aeruginosa, confirming the importance of an intact Tra_pKM101 T4SS for activation of the T6SS. Interestingly, however, E. coli strains engineered to produce any of the chimeric T4SSs (Tra::Trw, Tra::VirB, Tra::Ptl) triggered T6SS killing at levels comparable to the native Tra_pKM101 T4SS (Fig. 4C). These chimeric systems thus efficiently transmit an activating signal to P. aeruginosa target cells, despite wide variations in substrate transfer and a failure to support pilus production (Fig. 4B).

As expected from the above findings and our earlier phenotypic analyses (see Fig. 1), the ΔtraL mutant activated the P. aeruginosa T6SS while other Δtra mutants failed to trigger killing (Fig. S5). All complemented strains potentiated the killing response (Fig. S5), confirming the importance of an intact T4SS for signal transmission to P. aeruginosa. To further evaluate the requirements for contact-dependent signaling, E. coli strains harboring TraF mutations in the AP or CT domains were analyzed for T6SS activation. Interestingly, strains harboring TraF AP deletion (confers a Tra⁺,Pil⁻ phenotype) or CT mutations (Tra⁻,Pil⁻) efficiently triggered T6SS killing (Fig. 5A). These findings establish that the Tra_pKM101 T4SS retains the capacity to transduce a potentiating signal to target cells even in the absence of DNA transfer, elaboration of the OMCC cap, or formation of the WT pilus.

FIG. 5.

Requirements for activation of T6SS killing by P. aeruginosa PAO1. E. coli DH5α lacking or producing the Tra_pKM101 T4SS composed of the His₆-TraF variants shown; in each case, the traF allele was substituted for wild-type traF by incorporation into the pKM101 tra locus on plasmid pCGR108. Statistical significance is shown based on a Student’s t test corresponding to the values of plasmid-free DH5α or growth in the absence of P. aeruginosa (NS, not significant; *P < 0.05; **P < 0.01). Data presented are mean +/− SD, n = 3 independent replicates. Lower panel: Levels of His-TraF variants in total cell extracts, as monitored by immunostaining with α-His antibodies. RNA polymerase β-subunit (α-RNAP) served as a loading control.

Structure of the A. tumefaciens VirB/VirD4 OM core complex and comparisons with the pKM101 OMC

Finally, to allow for further structural comparisons of OMCC subassemblies shown here to be functionally interchangeable, we purified and solved the A. tumefaciens VirB/VirD4 OMCC structure by negative-stain electron microscopy (see Fig. S6). This OMCC has dimensions of 180 Å in diameter and 155 Å in height, closely resembling the pKM101- and R388-encoded OMCCs (Fronzes et al., 2009, Chandran et al., 2009, Low et al., 2014). The VirB OMCC also has 14-fold symmetry with openings at its proximal and distal ends whose dimensions are similar to those of the other OMCCs (Fig. 6). The VirB OMCC is more cylindrically-shaped than its Tra_pKM101 and Trw_R388 counterparts, and its OM-spanning cap is also broader with a notable cup presumptively exposed to the extracellular milieu. It is not yet possible to assess the significance of these structural differences due to the low resolution achieved for the VirB OMCC by negative staining. Nevertheless, the VirB structure adds to an accumulating body of evidence that OMCCs from the type IVa T4SSs exhibit strong similarities in their overall architectures. These findings also provide a structural basis for understanding the interchangeability of the heterologous OMCCs in supporting conjugative DNA transfer through the Tra_pKM101.

FIG. 6.

Negative-stain EM structure of the A. tumefaciens outer-membrane core complex (OMCC) and comparison with the NS-EM structure of the OMCC (EMDB-5032) encoded by E.coli pKM101. A) A. tumefaciens OMCC side view (left) and cut-away side view (right). B) E.coli pKM101 OMCC side view (left) and cut-away side view (right). C) Representation of the cut-away side view of the overlay of A. tumefaciens and E.coli pKM101 OMCC’s. D) Cross-section of overlaid A. tumefaciens and E.coli pKM101 OMCC complexes. Dashed line S in panel C indicates the level of the cross section shown in panel D.

DISCUSSION

E. coli pKM101 is widely known for its mutagenic and protective properties in cells exposed to UV irradiation and other DNA-damaging agents (Ames et al., 1975). pKM101 also encodes a highly-efficient T4SS (Winans & Walker, 1985, Paterson et al., 1999). In this study, we capitalized on the construction of a pKM101 ‘molecular toolkit’ to address a central question in the type IV secretion field, namely, how do these machines establish productive contacts with target cells? Guided in part by solved structures of T4SS subassemblies, we evaluated the contributions of the pKM101 Tra subunits and OMCC domains of special interest to pilus production, formation of donor-target cell contacts, and intercellular substrate transfer. Importantly, we determined that T4SSs lacking identified surface features, including the OM-spanning cap domain and the TraC pilus tip protein, display the Tra⁺, Pil⁻ “uncoupling” phenotype. Results of these mutational studies strongly implicate the OMCC cap and TraC as pilus-specificity factors and support a model in which the Tra_pKM101 T4SS alternatively functions as a pilus-assembly machine or an active translocation channel. We also determined that heterologous OMCC subassemblies could be substituted for the Tra_pKM101 OMCC. From a structural perspective, these latter findings underscore the broad importance of a conserved OMCC scaffold for elaboration of T4SS translocation channels. From a mechanistic perspective, however, the functionality of these chimeric T4SSs is remarkable, first, because two of the swapped OMCCs were derived from systems adapted for substrate trafficking to eukaryotic cells either by contact-dependent or -independent mechanisms. Second, as discussed further below, T4SSs are known to initiate substrate transfer in response to transduction of intracellular and extracellular signals across the cell envelope. The Tra_pKM101 IMC must therefore not only physically interact with heterologous OMCCs, but also convey energizing signals to the OMCC for channel activation.

We defined the functional importance of TraC and the other Tra subunits through systematic tra deletion/complementation analyses using native pKM101 and functional mini-pKM101 plasmids that were more amenable to genetic manipulation (Figs. 1 & S1). These studies established the essentiality of 9 of the 11 tra genes and also confirmed an early report that TraC is not required for pKM101 transfer on solid surfaces (Winans & Walker, 1985). Interestingly, however, we also found that pKM101 transfers at moderate frequencies (5×10⁻⁴ Tc’s/D) in 2 h liquid matings and that TraC is required for such transfer events. These traC mating phenotypes, coupled with the requirement of TraC for infection by the male-specific phage IKe (Figs. 1 & S1) (Yeo et al., 2003), strongly indicate that TraC is critical for pilus production but not a functional transfer channel. In their early study, Winans and Walker (1985) supplied genetic evidence that TraC is surface-exposed and might even be transmitted intercellularly. In a phenomenon termed ‘extracellular complementation’, a TraC-producing (but non-conjugative) ‘helper’ strain restores mating proficiency of a ΔtraC mutant when the two strains are mixed with a third, plasmid-free recipient strain. These investigators postulated that the ‘helper’ strain delivers surface-localized TraC to the ΔtraC mutant to fulfill a requisite function for pilus assembly and mating pair formation. Studies have since confirmed that TraC is indeed surface-localized even on cells lacking the Tra_pKM101 T4SS (Schmidt-Eisenlohr et al., 1999), although there is still no direct evidence for its cell-to-cell transmission. Further supporting the notion that the VirB5-like subunits are selectively involved in pilus production, B. pertussis Ptl_Bp system functions as an exporter even in the absence of a VirB5 homolog and detectable pilus production (Farizo et al., 2000, Locht et al., 2011). Mutations of virB5/traC-like genes do, however, abolish substrate transfer by some T4SSs (Berger & Christie, 1994, Fischer et al., 2001, de Paz et al., 2005, Larrea et al., 2013). In such systems, the VirB5-like subunits appear to have appropriated novel functions relating to target cell recognition. In H. pylori, for example, an RGD (Arg-Gly-Asp) motif and other motifs carried on VirB5-like CagL mediate binding of the Cag T4SS to β integrin receptors on human host cells (Kwok et al., 2007, Barden & Niemann, 2015). CagL also displays extensive sequence variation, thought to arise from evolutionary selection pressures within the human host, which allows for immune evasion or altered host cell binding by infecting H. pylori strains (Olbermann et al., 2010, Gorrell et al., 2016). In A. tumefaciens, VirB5 also carries an RGD motif (Backert et al., 2008), which similarly might contribute to establishment of productive contacts with susceptible plant cells (Lacroix & Citovsky, 2011). Finally, in Bartonella henselae, variant forms of surface-located VirB5 subunits are thought to determine host-specificity of erythrocyte parasitism (Dehio, 2008). CagL_Hp and VirB5_At localize at the tips of pili produced by the respective T4SSs, further supporting the idea that VirB5 subunits have evolved multiple functions relating to pilus nucleation and target cell binding (Aly & Baron, 2007, Kwok et al., 2007, Barden & Niemann, 2015).

The OM-spanning caps of the Tra_pKM101 and VirB/VirD4_At T4SSs were surprisingly permissive to mutation with respect to substrate transfer (Figs. 2, S2), suggesting that the translocation channel assembles across the OM even without an intact cap domain. The nature of this channel is not yet defined, but a growing body of evidence suggests that it consists of pilin monomers most probably in the form of a short pilus extending from the inner membrane to the cell surface. In earlier crosslinking studies, we showed that the VirB2_At pilin forms formaldehyde (FA)-crosslinkable contacts with DNA substrates during their transit through the A. tumefaciens VirB/VirD4 T4SS (Cascales & Christie, 2004b). VirB2_At - DNA crosslinking also was observed with variant channels harboring Tra⁺, Pil⁻ “uncoupling” mutations, but not among T pili isolated from the cell surface by shearing (Jakubowski et al., 2005, Cascales & Christie, 2004b). Very recently, structural studies of the F pilus revealed the striking finding that the inner lumen is composed of phospholipids derived from the inner membrane and in stoichiometric association with the TraA_F pilin subunit (Costa et al., 2016). In line with early models describing the dynamics of F pilus assembly and retraction (Manchak et al., 2002), these findings suggest that TraA_F pilin - phospholipid complexes comprise the building blocks for polymerization of the F pilus from an inner membrane platform (Costa et al., 2016). Some evidence also has been presented for the capacity of extended F pili to mediate substrate transfer in the apparent absence of direct donor-recipient cell contacts (Babic et al., 2008, Harrington & Rogerson, 1990). Such transfer events are rare, in line with extensive biochemical and some ultrastructural evidence that efficient conjugative transfer instead requires direct cell-to-cell contact (Samuels et al., 2000, Lawley et al., 2002, Arutyunov & Frost, 2013). Nevertheless, the capacity of extended pili to mediate substrate transfer is consistent with a model in which a pilus polymer extending across the donor cell envelope, and potentially beyond, functions as a conduit for substrate passage.

Our findings that the OMCC caps of the Tra_pKM101 and VirB/VirD4_At T4SSs are essential for pilus production (Figs. 2, S2) confirms and extends results of an earlier study showing that an A. tumefaciens VirB10ΔAP mutation confers low levels of surface-exposed VirB2 pilin and defects in pilus polymerization (Jakubowski et al., 2009). In that study, however, the AP boundaries (residues 308–337) were assigned on the basis of a crystal structure of VirB10-like ComB10 associated with a H. pylori competence system (Terradot et al., 2005). With the availability of the X-ray structure for the Tra_pKM101O-layer (Chandran et al., 2009), we reassigned the AP boundaries of both TraF and VirB10 so that the latter spans residues 282–335. The VirB10 AP mutant analyzed here (VirB10Δ282–335) completely blocked detection of surface-exposed VirB2 pilin (Fig. S2), while the corresponding TraF mutant (TraFΔ307–355) similarly blocked pilus production as monitored by IKe uptake and transfer in liquid (Fig. 2). Further reinforcing the notion that the distal end of the OMCC regulates pilus biogenesis, we previously showed that an Arg substitution for a highly-conserved Gly residue in VirB10_At that maps in the OMCC’s interior chamber and near the OM-spanning cap also blocks T pilus production without affecting substrate transfer to plant cells (Banta et al., 2011).

We incorporate our findings into a working model for conjugation systems in Gram-negative bacteria in which the default pathway is pilus production (Fig. 7). These pili function in a ‘mate-seeking’ mode either through dynamic rounds of extension and retraction as shown for F pili (Clarke et al., 2008) or via a mechanism(s) in which adhesive pili accumulate abundantly in the milieu to promote formation of mating aggregates (Samuels et al., 2000). Upon establishment of donor - recipient cell contacts, the T4SS ceases pilus production and transitions to the ‘mating’ mode. Various signals regulate the pilus-to-channel morphogenetic switch, including those propagated by the recipient cell (Frost & Koraimann, 2010, Arutyunov & Frost, 2013) and those from within the donor cell associated with substrate docking with VirD4-like receptors, engagement of the VirD4 receptor with the IMC, and ATP hydrolysis by the T4SS ATPases (Cascales & Christie, 2004a; de la Cruz et al., 2010; Lang et al., 2011; Lang & Zechner, 2012; Cascales et al., 2013). Our model is reminiscent of earlier models invoking a late-stage bifurcation in the T4SS assembly pathway for production either of the pilus or translocation channel (Christie et al., 2005, Trokter et al., 2014), but highlights the importance of the pilus tip protein and the distal end of the OMCC specifically for the pilus extension mode (Fig. 7). Accordingly, we propose that the pilus tip protein is recruited to and forms specific contacts with the OMCC as a prerequisite for pilus extension from the cell surface. Recruitment may occur at the extracellular surface, in view of evidence that VirB5-like subunits are exported across the OM independently of the T4SS (Schmidt-Eisenlohr et al., 1999). However, pilus tip proteins also have been reported to interact with IMC components (Yuan et al., 2005, Villamil Giraldo et al., 2012), raising the alternative possibility of their engagement with the T4SS in the periplasm. Regardless of the entry point, a central feature of our model is that the TraC/VirB5-like subunit engages - probably dynamically - with the distal end of the OMCC to drive extension of pilus from the cell surface. Extracellular and intracellular ‘mating’ signals thus might regulate the pilus-to-channel switch by blocking formation of productive contacts between the pilus tip protein and the OMCC cap or the major pilin subunit.

FIG. 7.

Working model for biogenesis of Type IVa secretion systems highlighting the importance of the postulated OMCC checkpoint in regulating pilus extension. Steps in the assembly pathway of the T4SS include (A) formation of the stable T4SS_3–10 substructure (Low et al., 2014) and (B) elaboration of a short pilus that extends from an inner membrane platform to the cell surface by a mechanism requiring TraB/VirB4- and TraG/VirB11-type ATPases. Next, (C) the pilus extends from the cell surface in a mate-seeking mode by a mechanism activated by recruitment of surface-exposed TraC to the distal end of the OMCC (denoted by yellow lightning bolt). TraC alternatively might be recruited to the T4SS via a periplasmic location (red-dashed line, ?). Finally, (D) upon pilus-mediated or direct contact with a recipient cell, a mating signal is transduced across the donor cell envelope resulting in recruitment of the TraJ/VirD4 substrate receptor, substrate docking and ATPase hydrolysis. These signals (denoted by lightning bolts) activate the morphogenetic switch to the T4SS ‘mating’ mode. The assembly intermediate depicted in (B) may bypass the pilus assembly (mate-seeking) mode (C) if presented with signals, e.g., recipient cell contact, required for activation of the substrate transfer (mating) mode (D), as could occur when donors and recipients grow in dense biofilm (solid-surface) communities. Abbreviations: OM, outer membrane; IM, inner membrane; P, peptidoglycan; OMCC, outer membrane complex; IMC, inner membrane complex; GSP, general secretory pathway. The pKM101 Tra proteins and their VirB counterparts required for each step of the assembly pathway are denoted.

Our studies also supplied important new insights into the OMCC domain requirements for T4SS function. For example, TraF’s CT domain (residues 355–386) is important for protein stability, but the conserved RDLDF motif within this domain is critical for TraF function (Fig. 2B). On the basis of the X-ray structure for the Tra_pKM101 O-layer (Chandran et al., 2009), we had envisioned that CT domain - lever arm contacts among adjacent TraF subunits (see Fig. S2C) might be important for OMCC assembly or stability. Indeed, the CT and lever arm deletions did impact TraF stability (Figs. 2 & S4), yet the Δlever mutant retained near WT function with respect to substrate transfer and pilus production. These findings establish that the lateral intersubunit contacts formed by the lever arm in the lateral shelf are not essential for assembly of a functional OMCC scaffold. Analyses of the TraF β-barrel domain swaps supplied further evidence for conformational flexibility of the OMCC. The functionality of the TraF/βB-CT_VirB10 chimera (Fig. 3), and of equivalent chimeras composed of the β-barrel domains from the TrwE_R388 and PtlG_Bp homologs (Fig. 5 & data not shown), was particularly surprising in view of the low overall sequence relatedness of the VirB10 homologs (Fig. S3). Furthermore, TraF’s β-barrel extensively interacts with VirB9-like TraO in the Tra_pKM101 O-layer crystal structure (Fig. S3C) (Chandran et al., 2009), and only a few of the residues comprising the TraF-TraO subunit interfaces are conserved among the β-barrel domains of the TraF homologs (Fig. S3C). These findings suggest that TraF’s network of intra- and intersubunit contacts do not structurally lock the Tra_pKM101 OMCC, which is in line with evidence that VirB10_At undergoes a conformational change in response to substrate docking and ATP energy signals, as well as unspecified extracellular signals, to regulate substrate passage.

The functionality of the chimeric T4SSs (Tra::VirB, Tra::Trw, Tra::Ptl) confirmed the modular nature of the IMC and OMCC subassemblies and supplied further evidence that these machines are intrinsically adaptive and flexible. While prior studies have shown that certain subunits of the OMCC (Llosa et al., 2003, de Paz et al., 2005) or to a limited degree the VirB8 subunit of the IMC (Paschos et al., 2006, Bourg et al., 2009) are exchangeable between closely related T4SSs, this is the first demonstration of the functionality of chimeric systems built from phylogenetically diverse IMC and OMCC subassemblies. At one level, the observed architectural similarities of the OMCCs solved previously (Fronzes et al., 2009, Chandran et al., 2009, Low et al., 2014) and here for the VirB OMCC (Fig. 6) provide a structural basis for understanding how heterologous OMCCs might substitute for the Tra_pKM101 subassembly. Moreover, given the widespread phylogenetic distribution of the VirB7, VirB9, and VirB10 homologs among the type IVa systems, and the recent identification of ring-shaped OMCCs associated with type IVb systems (represented by the Legionella pneumophila Dot/Icm T4SS) (Kubori et al., 2014, Kubori & Nagai, 2015), it is reasonable to predict that the OMCC structures solved to date are paradigmatic for T4SSs associated with Gram-negative species. However, the OMCC also must physically and functionally interact with the IMC to build the translocation channel and the pilus, and to regulate their dynamic activities. A complex network of contacts involving the VirB9- and VirB10-like OMCC subunits and the VirB6- and VirB8-like IMC subunits are required for elaboration of these structures (Hapfelmeier et al., 2000, Das & Xie, 2000, Krall et al., 2002, Jakubowski et al., 2003, Jakubowski et al., 2004, Baron, 2006). Contacts between the VirB10-like subunits and VirD4-like subset receptors also are implicated in transduction of the aforementioned intracellular (substrate docking/ATP energy) and extracellular (target cell binding) signals necessary for transitioning to the ‘mating’ mode (Llosa et al., 2003, Atmakuri et al., 2004, de Paz et al., 2005, Cascales & Christie, 2004a, Mihajlovic et al., 2009, Lang et al., 2011, Cascales et al., 2013, Arutyunov & Frost, 2013). Our use of chimeric TraF proteins facilitated productive coupling of pKM101’s IMC with the heterologous OMCCs, however, the functionality of the chimeric T4SSs almost certainly requires formation of other IMC-OMCC contacts as well as signal-activated conformational changes. Further studies of these and other chimeric T4SSs should reveal additional structure-function relationships between these two subassemblies.

Finally, our studies employing the P. aeruginosa T6SS killing assay confirmed that neither the Tra_pKM101 OMCC cap nor the extended pilus was required for establishment of potentiating donor - target cell contacts. Studies with the pKM101Δtra mutants supplied evidence that an intact Tra_pKM101 T4SS is essential for T6SS activation (Fig. S5), which agrees with earlier findings for the RP4 conjugation system (Ho et al., 2013). Strikingly, however, we also found that the chimeric T4SSs (Fig. 4), as well as Tra_pKM101 cap deletion and other Pil⁻ and Tra⁻ mutant strains (Figs. 5), triggered this killing system. That the Tra::Ptl chimera efficiently activated P. aeruoginosa T6SS killing was particularly surprising since the native Ptl system supports neither pilus biogenesis nor binding of B. pertussis to eukaryotic target cells (Fig. 5) (Burns, 2003, Locht et al., 2011). These findings establish that in absence of the OMCC cap or the extended pilus, another pKM101-encoded surface feature(s) must be capable of contacting and transmitting a potentiating signal to the P. aeruginosa cell envelope. Such a feature might correspond to i) a motif of the OMCC that becomes surface-exposed only upon target cell sensing ii) a short, surface-exposed pilus structure that was not detectable by our available assays, or iii) another surface-exposed protein that is not encoded by the pKM101 tra cluster but physically or functionally interacts with the OMCC. Future studies utilizing the T6SS killing assay should continue to refine our understanding of the T4SS surface features required for initiation of donor-recipient cell contacts and mating junction formation.

EXPERIMENTAL PROCEDURES

Strains and growth conditions

E. coli DH5α (GIBCO-BRL) was used for plasmid constructions and the type VI secretion system (T6SS) killing assay. E. coli MG1655 (E. coli Genetic Stock Center) served as donors in the conjugation assays and for the phage infection assays. E. coli CAG18477 served as recipients in the conjugation assays (Singer et al., 1989). E. coli HME45 (Thomason et al., 2014) was used for construction of tra gene deletions from native pKM101. Pseudomonas aeruginosa PAO1 (Holloway, 1955) containing an ISphoA insertion in the retS locus was used for the T6SS killing assay (Pseudomonas Transposon Mutant Collection, University of Washington Genome Sciences). E. coli strains were grown in Luria Broth (LB) at 37°C with shaking. E. coli strains were cultured in the following antibiotics: carbenicillin (50μg ml⁻¹), kanamycin (50μg ml⁻¹), spectinomycin (100μg ml⁻¹), chloramphenicol (20μg ml⁻¹), tetracycline (20μg ml⁻¹), and rifampicin (50μg ml⁻¹). P. aeruginosa PAO1 was grown in LB without antibiotic selection.

Plasmid constructions

Plasmids and oligonucleotides used in these studies are listed in Tables S1 and S2, respectively.

Vectors

pBAD24Spc was created by isolation of the spc^r gene as a SmaI fragment from pHP45Ω and inserting it into the ScaI site within the crb^r gene on pBAD24.

pKM101Δtra mutant plasmids

Eight of the 11 tra genes were deleted from pKM101 by recombineering (Thomason et al., 2014). Briefly, pKM101 or pKM101Spc^r were transferred by conjugation into E. coli strain HME45, which contains the bacteriophage λ red system under the control of the cI857 repressor. For construction of each tra gene deletion, the kan^r cassette from plasmid pUC4K was PCR amplified so that it carried flanking NcoI sites and 35 basepairs (bps) of 5′ and 3′ sequences that were complementary to regions immediately upstream and downstream of a tra gene of interest. HME45(pKM101Crb^r) or HME45(pKM101Spc^r) cells were temperature-induced for expression of the red-gam genes, and the kan^r amplicons were introduced by electroporation with Kan^r selection for transformants. Because pKM101 is a multicopy plasmid, we eliminated plasmids lacking the integrated kan^r cassette by subculturing the Kan^r transformants for 4 days in LB broth containing kanamycin (200 μg ml⁻¹). Isolated plasmids were digested with NcoI and religated to delete the kan^r cassette, and ligation mixes were introduced into DH5α with selection for Crb^r or Spc^r. Transformants were screened for Kan sensitivity, and tra deletion mutations were confirmed by sequencing across the deletion junction.

Mini-pKM101 plasmids

We constructed 2 mini-pKM101 plasmids with a goal of simplifying genetic manipulations of the tra gene cluster. pCGR108 was generated by introduction of the tra region from pKM101 into pBAD24. We amplified a ~10-kilobase region of pKM101 encompassing the upstream regulatory region and tra promoter through traG. This fragment was amplified with primers pKM101_2700NcoI_F and pKM101_13500XbaI_R and the resulting PCR product was introduced into pBAD24 using NcoI and XbaI restriction sites. The second mini-pKM101 plasmid, pRP100, was constructed by joining three PCR products: i) the tra gene cluster extending from the 3′ end of kikA through the end of traG, ii) the pKM101 oriV replication origin, and iii) an nptII gene encoding Kan^r. The ~10-kb tra gene cluster was amplified from pKM101 with primers pKM101_1921Nco1_F and pKM101_13500Xba_R, a ~3-kb region encompassing the replication origin was amplified with primers RSP007 and RSP008, and the nptII gene was amplified with primers RSP005 and RSP006 using plasmid pUC4K as a template. The replication origin and nptII gene were joined together using overlapping PCR, digested with NcoI and XbaI, and the resulting fragment was ligated to the tra gene cluster. The resulting circularized product was transformed into E. coli DH5α with Kan^r as a selection for self-replicating pRP100. Transformants were screened for plasmids bearing the three PCR fragments followed by sequence analysis of the PCR fragment junctions. We also confirmed that each of the mini-pKM101 plasmids encodes a fully functional Tra T4SS (see Results).

pRP100Δtra and pCGR108Δtra variants

We precisely deleted each of the tra genes from pRP100 by inverse PCR using the 5′phosphorylated primers listed in Table S1 and pRP100 as a template. The resulting plasmids, designated pRP101-pRP111, sustain deletions of traL through traG, respectively. We also deleted traF and traN-traF from pCGR108 to create pJG125 and pJG143, respectively, using a similar inverse PCR protocol, except that SacI and XhoI restriction sites were incorporated at the 5′ and 3′ ends of the deletion junctions.

pKM101 mob plasmid

We constructed a mobilizable plasmid bearing the pKM101 origin-of-transfer (oriT) sequences and adjacent traK, traJ, and traI genes. These genes code for the oriT processing proteins, relaxase TraI and accessory factor TraK, and the coupling protein TraJ. A PCR fragment spanning the oriT-traI region was generated with primers oriT_NcoI_F and TraI_HindIII_R and pKM101 as a template, and then introduced as a blunt-ended fragment into a blunt-ended HindIII site on the pSC101 derivative pGB2 to make pJG142.

tra gene expression plasmids

Plasmids pMS1 through pMS11 express the pKM101 traL through traG genes, respectively, from the P_BAD promoter. Each tra gene was PCR amplified using primers listed in Table S2, and pKM101 as a template. The resulting PCR fragments were digested with NcoI and KpnI for introduction into NcoI/KpnI-digested pBAD24Kan. Plasmids pJG59 and pJG62 express native and his₆-tagged traF, respectively, from the P_BAD promoter on pBAD24Spc. They were constructed by PCR amplification of traF using primers TraF_FWD_NcoI or TraF_NT_His_FWD NcoI and TraF_RVS_XhoI with pKM101 as a template, digestion of the PCR fragments with NcoI and XhoI, and introduction of the resulting fragments into NcoI/SalI-digested pBAD24Spc. Plasmid pJG103 expresses P_BAD::traF-CT_FLAG, producing C-terminally FLAG-tagged TraF. It was constructed by amplifying traF using primers TraF_FWD_NcoI and TraF_FLAG_CT_RVS_XhoI with pKM101 as a template. The resulting PCR fragment was digested with NcoI and XhoI and introduced into a NcoI/SalI digested pBAD24Spc

traF mutant plasmids

The following plasmids expressing traF mutant alleles from the P_BAD promoter were constructed by PCR amplification of gene fragments of interest using primers listed in Table S2 and pKM101 as a template, digestion of the final products with NcoI and XhoI, and introduction of the digested fragments into NcoI/SalI-digested pBAD24Spc. Plasmids: pJG95 produces TraFΔAP-CT from P_BAD::traF1-301 (numbers correspond to traF codons); pJG96 produces TraFΔCT from P_BAD::traF1-353; pJG76 produces TraFΔAP from P_BAD::traFΔ307-354 (traF1-307 and traF355-386 were amplified and joined by overlapping PCR, and cloned as above); pJG61 produces TraFAPL-5xGly from P_BAD::traFAPL-5xGly (traF1-322 and traF346-386 were amplified to carry a 5xGly residues at their 3′ and 5′ ends, respectively, and then joined by overlapping PCR); pJG64 produces His₆-TraF-FLAG330 from P_BAD::his₆-traF-FLAG330 (traF1-330 and 331-386 were amplified to carry a FLAG tag at their 3′ and 5′ ends, respectively, and then joined by overlapping PCR); JG101 produces TraFΔRDLDF from P_BAD::traFΔ373-377; pJG97 produces TraFΔCT9 from P_BAD::traFΔCT9. pJG92 produces TraFΔlever from pBAD::traFΔlever (traF1-170 and traF200-386 were amplified, and joined by overlapping PCR).

traF/virB10 chimera plasmids

The following plasmids expressing traF/virB10 chimeric genes were constructed by PCR amplification of gene fragments of interest using primers listed in Table S2 and pKM101 or traF fragments or pKVD10 for virB10 fragments. The amplification products (listed in parantheses) were joined by overlapping PCR, and the resulting fragments were digested with NcoI and XhoI for introduction into NcoI/SalI digested pBAD24Spc. Plasmids: pJG68 produces TraF/NT_VirB10 from P_BAD::traF/NT_virB10 (virB10.1-29 and traF40-386); pJG69 produces TraF/TMD_VirB10 from P_BAD::traF/TM_virB10 (traF1-40, virB10.29-50, traF60-386); pJG2005 produces TraF/TMD-CT_VirB10 from P_BAD::traF/TM-CT_virB10 (traF1-40, virB10.29-377); pJG70 produces TraF/PRR_VirB10 from P_BAD::traF/PRR_virB10 (traF.1-60, virB10.51-172, traF194-386); pJG150 produces TraF/βB_VirB10 from P_BAD::traF/βB_virB10 (traF1-193, virB10.173-286, traF307-386); pJG134 produces TraF/βB-AP_VirB10 from P_BAD::traF/βB-AP_virB10 (traF1-193 and virB10.173-335, traF356-386); pJG57 produces TraF/AP-CT_VirB10 from P_BAD::traF/AP-CT_virB10 (traF1-307, virB10.286-377); pJG58 produces TraF/βB-AP-CT_VirB10 from P_BAD::traF/βB-CT_virB10 (traF1-193, virB10.173-377). pJG60 produces TraF/AP_VirB10 from P_BAD::traF/AP_virB10 (traF1-307, virB10.286-335, traF356-386); pJG65 produces TraF/APL_VirB10 from P_BAD::traF/APL_virB10 (traF1-322, virB10.301-325, traF364-386); pJG66 produces TraF/CT_VirB10 from P_BAD::traF/CT_virB10 (traF1-354, virB10.335-377); pJG201 produces TraF/βB-CTΔ11_VirB10 from P_BAD::traF/βB-CTΔ11_virB10 (traF1-354, virB10.335-366).

Chimeric tra operons

Plasmid pJG145 expresses the chimeric gene cluster tra::trw. A DNA fragment encoding trwH-traE-trwF-traF/βB-CT_trwE was generated by overlapping PCR (trwH, traE, trwF, traF1-193, trwE197-395) using primers listed in Table S2 and pSU1443 and pKM101 as templates, and the resulting amplicon was digested with SacI and XhoI for introduction into pJG143. Plasmid pJG143 contains pKM101 traL-traD-SacI/XhoI-traG; it was derived from pCGR108 by inverse PCR. Plasmid pJG144 expresses the chimeric gene cluster tra::virB. A DNA fragment encoding virB7-traE-virB9-traF/βB-CT_virB10 was designed with codon-optimization for expression in E. coli, and synthesized by Genewiz Inc. The DNA fragment was isolated from pJG202 by digestion with SacI and XhoI and introduced into similarly-digested pJG143. Plasmid pJG151 expresses the chimeric gene cluster tra::ptl. A DNA fragment encoding ptlI-traE-ptlF-traF1-172/βB-CT_ptlG (ptlG160-374) was designed with codon-optimization for expression in E. coli, synthesized by Genewiz Inc. The DNA fragment was isolated from pJG202 and introduced into pJG143 as described above.

Mini-pKM101 plasmids with traF variants

Plasmid pJG125 carries the pKM101 tra genes except that XhoI and SacI sites were substituted for traF. It was constructed by inverse PCR using 5′ phosphorylated primers listed in Table S2 and pCGR125 as a template. pJG125 derivatives expressing different traF alleles were constructed by introduction of PCR fragments generated with primers and templates listed in Table S2 into SacI/XhoI-digested pJG125. Plasmids: pJG152 produces N-terminally FLAG-tagged TraF; pJG158 produces FLAG-TraFΔAP (deleted of codons 307–354); pJG154 produces FLAG_TraFΔRDLDF (deleted of codons 373–377); pJG153 produces FLAG-TraFΔCT9 (deleted of 9 codons at the 3′ end); pJG155 produces FLAG-TraF/AP-CT_VirB10 (traF1-307, virB10.286-377); pJG157 produces FLAG-TraF/AP-CT_TrwE (traF1-307/trwE303-395); pJG156 produces FLAG-TraF/AP-CT_PtlG (traF1-307/ptlG294-374).

A. tumefaciens virB10 expression plasmids

We incorporated a Strep-tag (St) sequence at the 3′ end of virB10 on plasmid pTiA6NC of strain A348 (Garfinkel et al., 1981). virB10-St was amplified with 500 basepairs (bps) of 5′ and 3′ flanking sequences using overlapping PCR and primers listed in Table S2. We then cloned this fragment into pBB50 for introduction into the ΔvirB10 derivative PC1010 by marker-exchange eviction mutagenesis, as previously described (Berger & Christie, 1994). The resulting strain, A348virB10-St, carrying the incorporated virB10-St gene was used for purification and structural characterization of the A. tumefaciens VirB/VirD4 OMCC.

We introduced the following plasmids expressing virB10 alleles into strain PC1010. Plasmid pKVD10 produces native VirB10 from P_lac::virB10 and fully complements the ΔvirB10 mutation (Jakubowski et al., 2009). Plasmids pSJ510, pSJ511, and pSJ512 were constructed by inserting an SphI restriction site at codons 298, 329, and 332, respectively, by inverse PCR using primers listed in Table S2 and pKVD10 as a template. We then PCR amplified 3×FLAG tag sequences, each with flanking SphI sites, using primers listed in Table S2 and pSJ503 as a template. Plasmid pSJ503 contains a 3×FLAG tag and was constructed by annealing oligonucleotides listed in Table S2, digesting the product with NcoI and BamHI, and inserting the digested fragment into similarly-digested pBSIISK⁺.NdeI. We then digested the amplified 3× FLAG tag sequences with SphI for insertion at codons 298, 329, and 332, creating plasmids pJG40, pJG42, and pJG44, respectively. Plasmid pJG52 producing VirB10 with a 2xFLAG tag inserted at codon 310 was constructed by two-step overlapping PCR using primers listed in Table S2 and pKVD10 as a template. Plasmid pSJ500 producing VirB10 with a C-terminal FLAG tag was constructed by amplification of virB10 with a 3′ terminal FLAG sequence using oligonucleotides listed in Table S2 and pKVD10 as a template.

Plasmid pSJ504 producing VirB10ΔAP (deleted of residues 288 – 337) was constructed by inverse PCR using primers listed in Table S2 and pKVD10 as a template. Plasmids pSJ501 and pSJ502 producing VirB10/AP_TraF and VirB10/AP-CT_TraF, respectively, were constructed by overlapping PCR using the primers listed in Table S2 and pKVD10 or pKM101 as templates. Following amplification, the virB10/AP_traF and virB10/AP-CT_traF products were digested with NdeI-XhoI for insertion into pBSIISK+.NdeI.

All ColE1 plasmids expressing the virB10 alleles were ligated to broad-host-range plasmids pXZ151 or pBBR-Kan for introduction into A. tumefaciens (Berger & Christie, 1994).

Conjugation assays

E. coli conjugation assays on solid surfaces were carried out essentially as previously described (Whitaker et al., 2016). Briefly, overnight cultures of donors and recipients were diluted 1:100 in antibiotic-free media and incubated for 1 h with shaking at 37°C. For induction from the P_BAD promoter, arabinose was added (0.2 % final concentration) followed by incubation for 1 h with shaking at 37°C. Donors and recipients (2.5 μl) were mixed on a nitrocellulose filter on LB media containing 0.2% arabinose and the mating mix was incubated for 2 h at 37°C. For broth matings, induced donors were mixed with recipients at a 1:1 volumetric ratio and incubated at 37°C for 2 h. Filter and broth mating mixtures were serially diluted and plated on media selective for transconjugants and donors. Frequency of transfer was calculated as the number of transconjugants per donor (Tcs/D). Experiments were performed at least three times in duplicate or triplicate, and results are reported as the mean frequency of transfer.

Phage infection assays

IKe bacteriophage was propagated as described previously for R17 (Lang et al., 2011). Strains carrying plasmids of interest were grown and assayed for susceptibility to IKe infection as previously described with slight modifications (Cellini et al., 1997). Briefly, cells were induced with arabinose as described above for the conjugation assays. Fifty microliters of cells at a concentration of ~10⁸ ml⁻¹ were spread on an LB plate containing appropriate antibiotics and arabinose, and allowed to dry. Five microliters of IKe (10⁶ pfu, final concentration) was spotted onto the lawns of cells, and plates were incubated overnight at 37°C.

Type VI killing assay

T4SS-mediated killing of E. coli by the Pseudomonas aeruginosa type VI secretion system (T6SS) was carried out as previously described (Ho et al., 2013). Briefly, 2ml of E. coli DH5α donors and a P. aeruginosa PAO1retS were incubated overnight with shaking at 37°C, then resuspended in 2ml of antibiotic-free LB followed by a 1:100 dilution in 5ml of antibiotic-free LB. Cells were then incubated with shaking at 37°C for 2 h, pelleted and resuspended in 100 μl LB. P. aeruginosa (17 μl) were mixed with E. coli (3μl) on filters placed on LB plates and incubated for 3 h at 37°C. Cells were resuspended in 1 ml of LB and serial dilutions were spotted onto plates containing spectinomycin (300μg/ml) and rifampicin (100μg/ml) to select for growth of E. coli. T6SS killing of E. coli is presented as E. coli cell viability in CFU per ml.

Protein detection

E. coli strains were grown and induced for expression of His₆- or FLAG-tagged TraF variants of interest in LB media, harvested, and normalized to equivalent optical densities (OD₆₀₀). Total protein extracts were subjected to SDS-PAGE, proteins were transferred to nitrocellulose membranes, and blots were developed with α-His or α-FLAG primary antibodies and HRP-conjugated secondary antibodies for detection of the TraF proteins by chemiluminescence (Whitaker et al., 2016). For VirB10 detection in A. tumefaciens, cells were grown and induced for expression of the vir genes (see below), and total cell extracts were analyzed by SDS-PAGE and immunostaining of western blots with α-VirB10 antibodies (Jakubowski et al., 2009).

Extracellular VirB2 blot assay

Surface-exposed VirB2 was detected by colony immunoblotting using α-VirB2 antibodies as described previously (Kerr & Christie, 2010).

A. tumefaciens outer-membrane complex expression and purification

A. tumefaciens strain A348virB10-St was inoculated in 100ml of MG/L media supplemented with 100ug/ml of Kanamycin. After overnight incubation at 26°C, 10 ml of culture pellet was inoculated into 200 ml of fresh MG/L media and incubated with shaking to an OD₆₀₀ of 0.5–0.8. The culture was further harvested by centrifugation and re-suspended in 6 L of ABIM media (supplemented with 100 mM of acetosyringone for vir genes expression) to an OD₆₀₀ of 0.1–0.2. After 12–14 h of incubation at 23°C, the cultures were harvested by centrifugation and re-suspended in cooled 50 mM Tris-HCl pH 8.0, treated with DNase I, lysosyme and EDTA-free protease inhibitor tablets, and sonicated on ice. After cell disruption, 1 mM EDTA was added and the lysate was clarified by centrifugation at 38000xg for 20 min. The membrane fraction was then collected by centrifugation at 98000xg for 45 min and membrane pellets were mechanically homogenized and solubilized in 50 mM of Tris-HCl pH 8.0, 200 mM NaCl, 1 mM EDTA, 0.5 % w/v Digitonin (Sigma), 0.75 % w/v DM-NPG (Anatrace), 0.5 % w/v DDM (Anatrace) and 1 mM DTT for 1h at 4°C. The suspension was clarified by centrifugation at 98000xg for 20 min and the supernatant was loaded onto a 5 ml Strep Trap HP (GE Healthcare) column and washed with 50mM of Tris-HCl pH 8.0, 200 mM NaCl, 1 mM EDTA, 0.1 % w/v Digitonin, 0.05 % w/v DM-NPG and 1 mM DTT at 4°C. The outer-membrane complex was eluted with the equivalent wash buffer supplemented with 2.5mM of desthiobiotin. The single peak fractions were pooled and loaded onto a Superose 6 10/300 column (GE Healthcare) equilibrated with the same buffer without desthiobiotin. The sample eluting as a peak after the column void was used immediately for the preparation of negative stain EM grids.

Electron microscopy and image processing

Sample preparation for EM

Four microliters of the OM core complex diluted to 0.01 mg/ml in the gel filtration buffer (see above) was applied on glow-discharged carbon-coated copper grids (400 mesh grid copper, Agar Scientific). After incubation for 2 min, the sample was washed twice with 10 μl water and then stained for 1 min with 10 μl 2 % uranyl acetate. Then the grids were blotted to remove excess stain, dried, and kept for the microscope sessions. The data were collected on a F20 microscope (FEI) operating at 200 kV at a magnification of 45,500xg using a low dose mode (~ 30 e Ų) and a defocus range of −0.7 to −2.0 μm. Images were recorded on a Gatan UltraScan 4000 CCD camera (Gatan) with a calibrated pixel size of 3.3 Å. 60 micrographs were collected. Quality was assessed visually and through CTF estimation. Images with distorted Thon rings and drift were not considered for the further processing.

Preprocessing

The contrast transfer function (CTF) of the microscope was estimated using CTFFIND3 (Mindell & Grigorieff, 2003) and the CTF correction of entire images was done by phase flipping using Bsoft (Heymann & Belnap, 2007). Particle images were picked from the CTF corrected micrographs. A total of 1746 particles were manually selected and extracted with a box size of 240×240 pixels using EMAN/BOXER (Ludtke, 2010). The following processing was done using IMAGIC software (van Heel et al., 1996). Images of particles were normalized, band pass filtered, centered, subjected to reference-free multi-statistical analysis (MSA), and then the alignment and classification were refined using iteratively multi-reference-alignment (MRA) and MSA where the best classes were used as new references. Quality of classes was assessed by variations between images that constitute classes: the lower variations -> the better class (van Heel et al., 2000). This refinement procedure was considered complete when changes in image shifts were no longer observed. A subset of the best 96 representative class averages (from 300) were assigned Euler angles by angular reconstitution (Van Heel, 1987) using the structure of the pKM101 core complex (EMDB – 5031) as a starting model. The three-dimensional structure was refined using reconstructions of the OM core complex generated using the best classes with the lowest errors at determination of angular orientations. The process was iterated (reconstruction – refinements of angles) until errors between class averaged images and reprojections of 3D maps stabilized and showed no further improvement. The final resolution of the map was 21 Å. The resolution was evaluated using the Fourier Shell Correlation and the threshold of 0.5 (van Heel et al., 2000). Surface rendering of EM maps was done using UCSF Chimera (Pettersen et al., 2004)