CdiA Effectors from Uropathogenic Escherichia coli Use Heterotrimeric Osmoporins as Receptors to Recognize Target Bacteria

Abstract

Many Gram-negative bacterial pathogens express contact-dependent growth inhibition (CDI) systems that promote cell-cell interaction. CDI⁺ bacteria express surface CdiA effector proteins, which transfer their C-terminal toxin domains into susceptible target cells upon binding to specific receptors. CDI⁺ cells also produce immunity proteins that neutralize the toxin domains delivered from neighboring siblings. Here, we show that CdiA^EC536 from uropathogenic Escherichia coli 536 (EC536) uses OmpC and OmpF as receptors to recognize target bacteria. E. coli mutants lacking either ompF or ompC are resistant to CDI^EC536-mediated growth inhibition, and both porins are required for target-cell adhesion to inhibitors that express CdiA^EC536. Experiments with single-chain OmpF fusions indicate that the CdiA^EC536 receptor is heterotrimeric OmpC-OmpF. Because the OmpC and OmpF porins are under selective pressure from bacteriophages and host immune systems, their surface-exposed loops vary between E. coli isolates. OmpC polymorphism has a significant impact on CDI^EC536 mediated competition, with many E. coli isolates expressing alleles that are not recognized by CdiA^EC536. Analyses of recombinant OmpC chimeras suggest that extracellular loops L4 and L5 are important recognition epitopes for CdiA^EC536. Loops L4 and L5 also account for much of the sequence variability between E. coli OmpC proteins, raising the possibility that CDI contributes to the selective pressure driving OmpC diversification. We find that the most efficient CdiA^EC536 receptors are encoded by isolates that carry the same cdi gene cluster as E. coli 536. Thus, it appears that CdiA effectors often bind preferentially to "self" receptors, thereby promoting interactions between sibling cells. As a consequence, these effector proteins cannot recognize nor suppress the growth of many potential competitors. These findings suggest that self-recognition and kin selection are important functions of CDI.

Author Summary

Bacterial pathogens often live in crowded communities where cells reside in close contact with one another. Many of these bacteria possess contact-dependent growth inhibition (CDI) systems, which allow cells to touch and inhibit each other using toxic CdiA proteins. CDI⁺ bacteria also produce immunity proteins that specifically protect the cell from the CdiA toxins of neighboring sibling cells. The CDI system from Escherichia coli EC93 was the first to be characterized and its CdiA toxin recognizes a receptor (BamA) that is identical in virtually all E. coli isolates. Here, we describe a different CDI system from uropathogenic E. coli 536, which causes urinary tract infections. In contrast to E. coli EC93, CdiA from E. coli 536 binds to receptor proteins (OmpC/OmpF) that vary widely between different E. coli isolates. Thus, uropathogenic E. coli preferentially bind and deliver toxins into sibling cells and other closely related E. coli strains. These results suggest that CDI systems distinguish between "self" and "non-self" cells. Moreover, because sibling cells are immune to CdiA-mediated growth inhibition, these findings raise the possibility that toxin exchange may be used for communication and cooperative behavior between genetically identical bacteria.

Introduction

Contact-dependent growth inhibition (CDI) systems mediate the transfer of protein toxins between Gram-negative bacteria. CDI was first discovered and characterized in Escherichia coli EC93, which uses CdiB^EC93/CdiA^EC93 two-partner secretion proteins to inhibit the growth of other E. coli isolates [1]. CdiB^EC93 is an Omp85 family β-barrel protein that exports and presents CdiA^EC93 on the cell surface. CdiA^EC93 carries toxic effector activity and is predicted to form a β-helical filament that projects several hundred angstroms from the inhibitor cell [2, 3]. CdiA^EC93 binds to BamA on the surface of neighboring E. coli cells, then delivers its C-terminal toxin domain (CdiA-CT^EC93) into the target cell to inhibit growth [4]. The proton gradient is rapidly dissipated in intoxicated cells [5, 6], suggesting that the CdiA-CT^EC93 toxin forms pores in the cytoplasmic membranes of target bacteria. Because E. coli EC93 cells also express BamA receptors, they receive toxin domains from neighboring siblings. However, inter-sibling toxin exchange does not induce growth arrest because the cdi locus encodes the CdiI^EC93 immunity protein, which neutralizes CdiA-CT^EC93 toxicity [1, 7]. Thus, E. coli EC93 deploys CDI to inhibit competing bacteria, but sibling cells are immune to growth inhibition. Since their discovery in E. coli EC93, cdi genes have been identified and characterized in several other proteobacteria [8–13]. CDI systems are typically encoded on plasmids and genomic islands and therefore are not necessarily found in all isolates of given species. This is exemplified by E. coli, for which ~25% of sequenced isolates carry cdi gene clusters. By contrast, every sequenced strain of Neisseria meningitidis and Burkholderia pseudomallei encodes at least one CDI system. CDI is also characterized by toxin diversity [3, 14]. CdiA-CT sequences vary widely between bacteria, with toxins exhibiting several distinct activities [3, 15]. CdiI immunity proteins are also diverse and specifically protect against cognate CdiA-CT toxins. Thus, cdi genes collectively encode a network of toxin/immunity protein pairs that appear to be rapidly diversifying. Toxin diversity and the specificity of immunity protection suggest that CDI systems are deployed to compete for environmental resources and growth niches.

The molecular mechanisms of CDI toxin delivery have been explored largely using the E. coli EC93 system as a model. Early work showed that CDI^EC93 expression in E. coli K-12 strains promotes auto-adhesion [1] and that CdiA-CT^EC93 delivery is receptor-dependent [4]. The CdiA^EC93 receptor, BamA, was identified from selections for CDI-resistant (CDI^R) target cells. E. coli bamA101 mutants carry a transposon insertion that reduces BamA expression five-fold and confers partial resistance to CDI^EC93-mediated growth inhibition [4]. BamA is an essential outer-membrane protein that forms the core of the β-barrel assembly machine (BAM) complex in Gram-negative bacteria [16–18]. The BAM complex inserts β-barrel proteins into the outer membrane, and BamA orthologues are found in all Gram-negative bacteria and eukaryotic plastids. Though BamA is conserved, its extracellular loops vary considerably between closely related enterobacterial species [19]. For example, E. coli BamA^Eco shares ~94% overall sequence identity with BamA^ECL from Enterobacter cloacae ATCC 13047, but the surface-exposed portions of extracellular loops L4, L6 and L7 are unrelated in sequence [20]. Consequently, E. coli EC93 does not recognize E. cloacae cells as targets and cannot inhibit the growth of this species. Similarly, replacement of bamA ^Eco with bamA genes from other enterobacterial species confers CDI-resistance to E. coli cells [20]. Thus, receptor polymorphism restricts the CDI^EC93 target-cell range to E. coli and Shigella isolates, which share identical BamA proteins. Recognition of "self" receptors appears to be a general feature of CDI, because CdiA effectors from Dickeya dadantii 3937, Burkholderia thailandensis E264, Neisseria meningitidis B16B6 and Pseudomonas aeruginosa PAO1 all deliver toxins to sibling cells [8–13]. This exchange of toxins provides no apparent competitive advantage because siblings are immune. However, the accompanying cell-cell adhesion likely imparts significant benefits by promoting auto-aggregation and biofilm formation [7, 11, 13, 21–25]. Thus, in addition to suppressing the growth of competing bacteria, CDI also contributes to bacterial fitness by mediating cooperative behaviors between sibling cells.

Bacterial surfaces interact directly with the environment and are therefore under strong selective pressure. Bacteriophages and adaptive immune systems exploit surface epitopes to identify bacteria for infection and elimination, respectively. In response, bacteria have evolved mechanisms to rapidly alter their repertoire of surface molecules such that different isolates of the same species commonly express unique extracellular antigens [26–29]. Therefore, CdiA effectors from different bacterial species must presumably recognize distinct receptors. Here, we begin to explore CDI receptor diversity using the CdiA^EC536 effector from uropathogenic E. coli 536 (EC536). We find that E. coli ompC and ompF mutants are resistant to CDI^EC536-mediated growth inhibition, suggesting that the OmpC and OmpF osmoporins function as receptors for CdiA^EC536. CDI^EC536 inhibitor cells only bind to target bacteria that express OmpC and OmpF, indicating that both porins are required for receptor function. Using single-chain OmpF dimers, we provide evidence that heterotrimeric OmpC-OmpF constitutes the receptor for CdiA^EC536. Because OmpF and OmpC exhibit considerable sequence variability in their extracellular loops [30, 31], we examined the effects of these polymorphisms on receptor function. We find that several OmpC proteins from uropathogenic E. coli isolates are not recognized by CdiA^EC536, suggesting that many potential competitors are resistant to CDI^EC536. By contrast, OmpC proteins from E. coli UTI89 and F11 are effective receptors for CdiA^EC536. Notably, E. coli UTI89 and F11 also encode CdiA proteins that are >98% identical to CdiA^EC536, indicating that their effectors also recognize OmpC-OmpF receptors and mediate self-recognition. In these instances, CdiA effector proteins preferentially recognize "self" receptors, suggesting that auto-adhesion and kin selection are important functions of CDI.

Results

Genetic analysis of the CDI^EC536 system

We previously reported that E. coli bamA101 mutants are resistant to CDI^EC536 [32], though the level of protection is less than that observed against CDI^EC93 inhibitors [4]. Because the bamA101 mutation confers incomplete resistance, we revisited these experiments using E. coli target strains that express E. cloacae bamA (bamA ^ECL), which provides full resistance to CDI^EC93 [20]. We first confirmed that E. coli bamA ^ECL target cells are resistant to CDI^EC93 using E. coli EPI100 inhibitor cells that express the cdiBAI ^EC93 gene cluster from a cosmid vector. As expected, E. coli bamA ^ECL targets grew to the same level in co-culture with either CDI^EC93 inhibitors or CDI⁻cells that carry an empty cosmid vector (Fig 1). By contrast, bamA ^Eco target-cell viability decreased ~600-fold during co-culture with CDI^EC93 inhibitors (Fig 1). We then tested bamA ^Eco and bamA ^ECL cells in competitions against E. coli EPI100 that express the E. coli 536 cdiBAI ^EC536 genes from a cosmid and found that both target strains were inhibited to the same extent (Fig 1). These results indicate that BamA^ECL does not protect against CDI^EC536, suggesting that the originally observed bamA101 resistance may be due to decreased expression of an unidentified outer-membrane receptor that requires BamA for assembly.

Fig 1. Heterologous BamA does not protect against CDI^EC536.

Target bacteria that express bamA ^Eco (CH10226) or bamA ^ECL (CH10227) were cultured at a 1:10 ratio with E. coli EPI100 inhibitor cells that carry cosmids pDAL660Δ1–39 (CDI^EC93), pDAL866 (CDI^EC536) or pWEB::TNC (CDI^–). Viable target bacteria are reported as colony-forming units per milliliter (cfu/mL) ± SEM for two independent experiments.

We used a genetic approach to identify the CdiA^EC536 receptor, reasoning that its disruption should confer CDI resistance (CDI^R). We subjected CDI-sensitive E. coli target cells to transposon mutagenesis, then selected CDI^R mutants from co-cultures with E. coli 536 cells that express the cdi locus under control of an arabinose-inducible promoter. Additionally, because CysK (O-acetylserine sulfhydrylase A) is required to activate the CdiA-CT^EC536 toxin domain [33, 34], we provided target cells with plasmid-borne cysK to avoid isolating CDI^R cysK null mutants. CDI^R populations were enriched after three rounds of selection, and randomly isolated clones exhibited complete resistance to growth inhibition. Transposon insertions from CDI^R clones were transduced into CDI-sensitive cells and genetic linkage between insertions and CDI^R phenotypes was established for each transductant. The disrupted genes were then identified from ten randomly selected CDI^R mutants, revealing five independent insertions each in ompC and ompF (Fig 2A). The ompC and ompF genes encode closely related osmoporins, which are trimeric β-barrel proteins that facilitate the diffusion of small hydrophilic molecules across the outer membranes of Gram-negative bacteria [35]. To confirm the roles of OmpC and OmpF in CDI^EC536, we generated an E. coli ΔompC ΔompF double mutant and performed complementation analysis. As expected, the growth of ΔompC ΔompF cells was not inhibited in co-culture with E. coli 536 (Fig 2B). By contrast, wild-type ompC ⁺ ompF ⁺ target cells showed a 50-fold loss in viability under the same co-culture conditions (Fig 2B). This latter growth inhibition is due to CDI^EC536, because the growth of wild-type E. coli was not affected by E. coli 536 mock inhibitors that carry a deletion of the cdiA-CT coding region (Fig 2C). Complementation of the ΔompC ΔompF mutant with a plasmid expressing either ompC or ompF had no effect on the resistance phenotype, but the strain became CDI-sensitive when it harbored plasmids expressing both osmoporins (Fig 2B). Together, these findings demonstrate that OmpC and OmpF expression is required to render bacteria sensitive to the CDI^EC536 system.

Fig 2. E. coli ΔompC and ΔompF mutants are resistant to CDI^EC536.

(A) Positions of mariner transposon insertion sites from CDI^R mutants. (B) Competition co-cultures with CDI⁺ E. coli 536 inhibitors. E. coli CH10013 (ompC ⁺ ompF ⁺) and CH10136 (ΔompC ΔompF) target cells were co-cultured at a 1:10 ratio with CDI-inducible E. coli 536 cells. The plus (+) symbols indicate complementation of ΔompC ΔompF target cells with plasmid-borne ompC ^K-12 (pCH10138) and ompF ^K-12 (pCH10202). Viable target bacteria are reported as colony-forming units per milliliter (cfu/mL) ± SEM for three independent experiments. (C) Competition co-cultures with CDI⁻ E. coli 536. Co-cultures were performed as in panel B, except E. coli 536 cells deleted for the cdiA-CT region were used as mock inhibitors.

OmpC and OmpF are required for CdiA^EC536-mediated cell-cell adhesion

CdiA-mediated adhesion between inhibitor and target cells is a hallmark of CDI. To determine whether OmpC and OmpF are required for CDI-dependent adhesion, we monitored cell-cell binding using a flow-cytometry approach. GFP-labeled inhibitor cells were mixed with DsRed-labeled target bacteria at a 5:1 ratio, and the suspensions analyzed by flow cytometry. Flow events registering both green and red fluorescence correspond to aggregates containing inhibitor and target bacteria. The flow cytometric assay for CDI-mediated adhesion generally requires CdiA over-expression [4, 36], therefore we used inhibitor cells that express cdi genes from cosmids for these experiments. Under these conditions, CDI^EC93 inhibitor cells bind to 60–80% of target bacteria (Fig 3A). This cell-cell adhesion is dependent on CDI, with only about 5% of target cells associated with CDI⁻mock inhibitors (Fig 3A). We then tested whether CDI^EC536 expression also promotes stable adhesion to target cells. As expected, we did not detect CDI^EC536-dependent adhesion to ΔompC target bacteria; but surprisingly there was little increase in adhesion upon complementation with ompC ^K-12 from E. coli K-12 (Fig 3A). Despite the lack of detectable cell-cell adhesion, ompC ^K-12-complemented cells were inhibited during co-culture with E. coli 536 cells (Fig 3B) and immunoblotting confirmed that they produced OmpC^K-12 (Fig 3C). OmpC porins vary significantly between E. coli isolates [30], and the E. coli K-12 and 536 proteins carry different surface-exposed loops (Fig 3D). Because extracellular loop polymorphism could affect receptor function, we tested whether ompC ^EC536 better supports CDI^EC536-dependent cell adhesion. Strikingly, over 40% of ompC ^EC536 complemented bacteria adhered to CDI^EC536 inhibitors, and this adhesion was specific because these target cells did not associate with CDI⁻mock inhibitors (Fig 3A). These results suggest that CdiA^EC536 binds to OmpC^EC536 with higher avidity than OmpC^K-12. We also found that ompC ^EC536 target cells were inhibited to a greater extent than ompC ^K-12 targets in co-cultures with E. coli 536 (Fig 3B). We next tested OmpC proteins from Salmonella Typhimurium LT2 and E. cloacae ATCC 13047 (Fig 3D) to determine whether other species are potential targets for the CDI^EC536 system. E. coli ΔompC cells complemented with S. Typhimurium ompC ^LT2 did not adhere to CDI^EC536 inhibitors (Fig 3A) and were not inhibited in competition co-culture (Fig 3B), though they produced substantial OmpC^LT2 protein (Fig 3C). By contrast, CDI^EC536 inhibitors bound to ompC ^ECL complemented cells and inhibited their growth (Fig 3A and 3B). Together, these results support a role for OmpC as the CdiA^EC536 receptor and indicate that OmpC polymorphism determines whether bacteria are susceptible to CDI^EC536-mediated growth inhibition.

Fig 3. OmpC is required for CDI^EC536 dependent cell-cell adhesion.

(A) Analysis of cell-cell adhesion by flow cytometry. E. coli DL4905 (GFP⁺) inhibitors carrying cosmids pDAL660Δ1–39 (CDI^EC93), pDAL866 (CDI^EC536) or pWEB::TNC (CDI^–) were mixed at a 5:1 ratio with DsRed-labeled CH10099 (ΔompC) target cells. Where indicated, the target cells were complemented with plasmid-borne ompC alleles from E. coli K-12, E. coli 536 (EC536), S. Typhimurium LT2 or E. cloacae (ECL). The suspensions were analyzed by flow cytometry using FL1 (533/30nm, GFP) and FL2 (585/40nm, DsRed) fluorophore filters. The percentage of bound target cells was calculated as the number of dual green/red fluorescent events divided by the total number of red fluorescent events. The average ± SEM is presented for two independent experiments. (B) Competition co-cultures with CDI⁺ E. coli 536. E. coli CH10099 (ΔompC) target cells were co-cultured at a 1:10 ratio with CDI-inducible E. coli 536 cells. Where indicated, the target cells were complemented with plasmid-borne ompC alleles from E. coli K-12, E. coli 536 (EC536), S. Typhimurium LT2 or E. cloacae (ECL). Viable target bacteria are reported as colony-forming units per milliliter (cfu/mL) ± SEM for two independent experiments. (C) Protein from the target bacteria in panel A was subjected to immunoblot analysis using polyclonal antisera against OmpC/F from E. coli K-12. (D) Sequence alignment of OmpC proteins from E. coli K-12, E. coli 536, S. Typhimurium LT2 and E. cloacae ATCC 13047 (ECL). Extracellular loop sequences are indicated above the alignment.

We next examined the contribution of OmpF to CDI^EC536-dependent adhesion by testing whether porins from different bacteria influence receptor activity (Fig 4A). Because OmpC^K-12 does not support detectable adhesion to CDI^EC536 inhibitor cells, we replaced the native ompC ^K-12 locus with the ompC ^EC536 allele to generate a target-cell strain for cell-cell binding studies. The resulting ΔompF ompC ^EC536 cells did not form aggregates with CDI^EC536 inhibitors, but adhered normally to CDI^EC93 inhibitor cells (Fig 4B). We then expressed plasmid-borne ompF from E. coli K-12, E. coli 536, S. Typhimurium and E. cloacae and found that each allele supported adhesion to CDI^EC536 inhibitors (Fig 4B and 4C). These results demonstrate that both OmpC and OmpF are required for CDI^EC536-dependent cell adhesion, indicating that each porin contributes to receptor function.

Fig 4. OmpF is required for CDI^EC536 dependent cell-cell adhesion.

(A) Sequence alignment of OmpF proteins from E. coli K-12, E. coli 536, S. Typhimurium LT2 and E. cloacae ATCC 13047 (ECL). Extracellular loop sequences are indicated above the alignment. (B) Analysis of cell-cell adhesion by flow cytometry. E. coli DL4905 (GFP⁺) inhibitors carrying cosmids pDAL660Δ1–39 (CDI^EC93), pDAL866 (CDI^EC536) or pWEB::TNC (CDI^–) were mixed at a 5:1 ratio with DsRed-labeled CH12134 (ΔompF ompC ^EC536) target cells. Where indicated, target cells were complemented with plasmid-borne ompF from E. coli K-12, E. coli 536 (EC536), S. Typhimurium LT2 or E. cloacae (ECL). Cells suspensions were analyzed by flow cytometry using FL1 (533/30nm, GFP) and FL2 (585/40nm, DsRed) fluorophore filters. The percentage of bound target cells was calculated as the number of dual green/red fluorescent events divided by the total number of red fluorescent events. The average ± SEM is presented for two independent experiments. (C) Protein from the target bacteria in panel B was subjected to immunoblot analysis using polyclonal antisera against OmpC/F from E. coli K-12.

CdiA^EC536 recognizes OmpC-OmpF heterotrimers

At least two models can explain the roles of OmpC and OmpF in CDI. CdiA^EC536 could interact simultaneously with adjacent OmpC and OmpF homotrimers on the target-cell surface. Alternatively, the effector could recognize OmpC and OmpF displayed as heterotrimers, which have been reported previously [37]. To differentiate between these models, we generated and tested the function of covalently constrained OmpC-OmpF heterotrimers. Because osmoporins are obligate trimers, we reasoned that porin dimers produced from a gene fusion should require an additional monomer to assemble properly in the outer membrane. We tested this prediction using dimeric OmpF proteins tethered by 8- and 16-residue linker peptides. OmpF assembly and function was assessed by monitoring sensitivity to colicin E5, which requires OmpF to translocate its C-terminal toxin domain into E. coli cells [38–41]. E. coli ΔompF ΔompC mutants are completely resistant to colicin E5, but become susceptible when complemented with plasmid-borne ompF ^EC536 (Fig 5A). By contrast, neither of the dimeric ompF constructs restored colicin sensitivity (Fig 5A), even though immunoblot analysis confirmed the presence of OmpF dimers in these cells (Fig 5B). We then expressed ompF-ompF fusions in the ΔompF ompC ^EC536 background to test whether monomeric OmpC supports heterotrimer formation. E. coli ΔompF ompC ^EC536 cells are somewhat sensitive to colicin E5 (Fig 5C), consistent with a prior report showing that OmpC can support colicin entry in the absence of OmpF [42]. However, colicin E5 had a greater inhibitory effect on ΔompF ompC ^EC536 cells that were complemented with plasmids that produce either monomeric or dimeric OmpF (Fig 5C). Together, these results indicate that OmpF₂-OmpC heterotrimers assemble in the outer membrane and are active in colicin translocation. Having established a strategy to generate heterotrimeric porin complexes, we tested whether OmpF₂-OmpC is a receptor for CdiA^EC536. Flow cytometry showed that each OmpF dimer supported CDI^EC536-dependent cell-cell adhesion to the same extent as monomeric OmpF (Fig 6A). Moreover, we also found that ΔompF ompC ^EC536 cells expressing dimeric OmpF were inhibited 10- to 100-fold by E. coli 536 cells in competition co-cultures (Fig 6B). Taken together, these results indicate that heterotrimeric OmpF-OmpC is the receptor for CdiA^EC536.

Fig 5. Characterization of functional OmpF-OmpC heterotrimers.

(A) E. coli CH11346 (ΔompF ΔompC) cells were complemented with ompF (pCH11850), ompF-ompF with 8-residue linker (ompF ₂, pCH10796), or ompF-ompF with 16-residue linker (ompF ₂ *, pCH12054) and the resulting strains cultured in LB media without or with colicin E5 (added at 30 min as indicated by the arrow). Cell growth was monitored by measuring the optical density at 600 nm (OD₆₀₀) of the cultures. Data are from two independent experiments. (B) Protein from the strains in panel A was subjected to immunoblot analysis using polyclonal antisera against OmpC/F and BamA (as a loading control) from E. coli K-12. (C) E. coli CH12180 (ΔompF ompC ^EC536) cells were complemented with plasmid-borne ompF, ompF ₂ or ompF ₂* and cultured as described in panel A. (D) Protein from the strains in panel C was subjected to immunoblot analysis using polyclonal antisera against OmpC/F from E. coli K-12.

Fig 6. Heterotrimeric OmpF-OmpC supports CDI^EC536-mediated growth inhibition.

(A) E. coli DL4905 (GFP⁺) inhibitors carrying pDAL866 (CDI^EC536) or pWEB::TNC (CDI^–) were mixed at a 5:1 ratio with DsRed-labeled E. coli CH12180 (ΔompF ompC ^EC536) target cells complemented with ompF (pCH11850), ompF-ompF with 8-residue linker (ompF ₂, pCH10796), or ompF-ompF with 16-residue linker (ompF ₂ *, pCH12054). Cell suspensions were analyzed by flow cytometry using FL1 (533/30nm, GFP) and FL2 (585/40nm, DsRed) fluorophore filters. The fraction of bound target cells was calculated as the number of dual green/red fluorescent events divided by the total number of red fluorescent events. The average ± SEM is presented for two independent experiments. (B) E. coli CH12180 (ΔompF ompC ^EC536) target cells were complemented were complemented with plasmid-borne ompF, ompF ₂ or ompF ₂* and the resulting strains co-cultured at a 1:10 ratio with CDI⁺ (filled symbols) or CDI⁻(open symbols) E. coli 536 cells. Viable target bacteria are reported as colony-forming units per milliliter (cfu/mL) ± SEM for three independent experiments.

OmpC loops L4 and L5 are recognition determinants for CdiA^EC536

We next used natural variations between E. coli OmpC porins to identify potential CdiA^EC536 recognition epitopes. We surveyed predicted OmpC proteins from E. coli isolates and identified over 220 distinct sequences (S1 Table). Though some E. coli OmpC variants differ by single amino-acid residue substitutions, much of the sequence variation is localized to extracellular loops L4 (33 sequences), L5 (30 sequences) and L7 (13 sequences) (S2 Table). Moreover, these loops are found in many different combinations (S1 Table). To examine the influence of OmpC polymorphism on CDI^EC536, we tested a subset of E. coli OmpC proteins with unique extracellular loop combinations (Figs 7A and S1). E. coli ΔompC target cells were provided with plasmid-borne ompC alleles and co-cultured with E. coli 536 inhibitor cells. The ompC alleles from E. coli UTI89, F11 and A33H rendered ΔompC cells sensitive to growth inhibition to roughly the same extent as ompC ^EC536 (Fig 7B). Targets that express ompC ^EC93 from E. coli EC93 were also inhibited in co-culture with E. coli 536, but ompC alleles from uropathogenic isolates CFT073, 6104H, A54H, A42H and A35H were all associated with resistance (Fig 7B and 7C). Immunoblot analysis confirmed that OmpC was produced in all of the complemented strains (Fig 7D), indicating that resistance is not due to a lack of ompC expression. Thus, E. coli ompC allele polymorphism appears to dictate cell susceptibility to CDI^EC536-mediated intoxication.

Fig 7. E. coli OmpC polymorphism restricts the target-cell range of CDI^EC536.

(A) Extracellular loop L4, L5 and L7 sequences from selected E. coli isolates. S1 Fig shows the complete sequence alignment, and the alphanumeric classification scheme for L4, L5 and L7 sequences is presented in S2 Table. (B) E. coli CH10099 (ΔompC) target cells were co-cultured at a 1:10 ratio with CDI⁺ E. coli 536 cells. Target cells were complemented with plasmid-borne ompC alleles from the indicated E. coli isolates. Viable target bacteria are reported as colony-forming units per milliliter (cfu/mL) ± SEM for three independent experiments. (C) The same target-cell strains from panel B were co-cultured at a 1:10 ratio with CDI⁻ E. coli 536 and viable cells quantified in the same manner. (D) Protein from ompC complemented strains was subjected to immunoblot analysis using polyclonal antisera against OmpC/F from E. coli K-12.

To determine which OmpC sequences are critical for recognition by CdiA^EC536, we generated chimeric porins using OmpC^CFT073 as a framework upon which to graft heterologous OmpC loops. OmpC^CFT073 and OmpC^EC536 share loop L4, but differ in their L5, L7 and L8 sequences (Figs 7A and S1). OmpC^CFT073 chimeras carrying L7 or L8 from OmpC^EC536 still provided resistance to CDI^EC536 (Fig 8A), indicating that these loops probably have no role in binding CdiA^EC536. However, replacement of loop L5 converted OmpC^CFT073 into a functional receptor that supports the same level of inhibition as OmpC^EC536 (Fig 8A). Similarly, OmpC^CFT073 chimeras carrying OmpC^EC536 L7 or L8 were functional receptors, provided that they also carried loop L5 from OmpC^EC536 (Fig 8A). We tested the role of loop L4 by grafting the sequence from OmpC^F11 (which is a receptor for CdiA^EC536) onto OmpC^CFT073. Targets that express the resulting chimera were inhibited by E. coli 536 to about the same extent as targets that express OmpC^F11 (Fig 8A). This latter observation shows that OmpC^CFT073 loop L5 is not an anti-determinant, because it supports CdiA^EC536 recognition in other contexts. Collectively, these results indicate that OmpC loops L4 and L5 contribute to target-cell selection, suggesting that the loops may be recognized directly by CdiA^EC536.

Fig 8. OmpC extracellular loops L4 and L5 are important recognition determinants for CdiA^EC536.

(A) E. coli CH10099 (ΔompC) target cells were co-cultured at a 1:10 ratio with CDI⁺ E. coli 536 cells. Target cells were complemented with the indicated chimeric ompC alleles. Viable target bacteria are reported as colony-forming units per milliliter (cfu/mL) ± SEM for three independent experiments. (B) Protein from the strains in panel A was subjected to immunoblot analysis using polyclonal antisera against OmpC/F from E. coli K-12.

Discussion

The results presented here establish a critical role for OmpC and OmpF in the CDI^EC536 pathway. CDI^EC536 inhibitor cells only adhere to target bacteria that express OmpC and OmpF, indicating that both porins contribute to receptor function. Though osmoporins are generally assumed to form homotrimers, we show that heterotrimeric OmpC-OmpF constitutes the CdiA^EC536 receptor. Heterotrimeric porins were first reported by Gehring and Nikaido, who found that OmpC, OmpF and PhoE can form mixed trimers in E. coli K-12 [37]. OmpC^K-12 and OmpF^K-12 share 58.5% sequence identity (S2 Fig), and their β-barrels superimpose with root-square mean deviation of 0.78 Å [43]. The trimer interaction surface is dominated by hydrophobic contacts between β-barrel strands β1, β2, β3, β4, β5 and β1´, and these regions are highly conserved between OmpC^K-12 and OmpF^K-12 (S2 and S3 Figs, S3 Table). The porins also share an inter-protomer ion-pair between a conserved Glu residue (Glu66 in OmpC^K-12 and Glu71 in OmpF^K-12) at the tip of loop L2 and an Arg residue within L3 of the adjacent β-barrel (S3 Fig and S3 Table). Thus, the structures of OmpF and OmpC are compatible with heterotrimer formation. The targeting of heterotrimeric OmpF-OmpC receptors has a number of implications. Firstly, residues from both porins should be directly recognized by CdiA^EC536. The results presented here suggest that OmpC loops L4 and L5 are important CdiA^EC536-binding determinants, but the contribution of OmpF is less clear. OmpF porins from E. cloacae and S. Typhimurium are effective co-receptors in conjunction with OmpC^EC536, so the CdiA-binding epitope is presumably shared by all of the tested OmpF variants. Additionally, because OmpC and OmpF are regulated reciprocally, it is possible that receptor availability fluctuates in response to environmental conditions. OmpF is expressed preferentially in response to low osmolarity, whereas OmpC dominates under hyperosmotic conditions [35]. We find that target cells are effectively inhibited when they over-express ompC alleles, suggesting that OmpF-OmpC₂ heterotrimers are recognized by CdiA^EC536. We have also shown that covalently constrained OmpF₂-OmpC heterotrimers are functional receptors. Because both heterotrimeric forms support CDI^EC536, we conclude that CdiA^EC536 effectors probably make contact with one OmpC and one OmpF protomer in the receptor complex.

The mechanism(s) by which CDI toxins are translocated across the target-cell outer membrane is unknown, but in principle, receptors could mediate this transport. Group A colicins appear to exploit the central pore of OmpF to cross the outer membrane, and this translocation is powered by the proton-motive force via the Tol protein complex [40, 41, 44, 45]. However, CDI toxin translocation is not Tol-dependent, nor does it require the Ton-Exb system, which energizes the import of group B colicins [6, 44]. Further, CDI toxin domains are modular and can be exchanged between CdiA proteins for delivery through different receptor pathways [8, 10, 46]. Therefore, if CdiA receptors mediate outer-membrane transport, then BamA must also be capable of CDI toxin translocation. Like OmpF and OmpC, BamA is a 16-strand β-barrel protein, but its lumen is not exposed to the extracellular milieu. Instead, extracellular loops L3, L4, L6, L7 and L8 form a dome that shields the BamA lumen from the environment [47, 48]. It is possible that the BamA barrel opens upon binding to CdiA^EC93, consistent with work showing that extracellular loop L6 is dynamic during the OMP assembly duty cycle [49, 50]. Buchanan and colleagues have also suggested that the BamA β-barrel opens a lateral gate during OMP biogenesis [51]. These conformational changes could be exploited to translocate toxin across the outer membrane, but we note that inactive BamA lacking the POTRA-3 (polypeptide-transport associated) domain is able to complement the bamA101 mutation and restore sensitivity to CDI^EC93 [4]. Therefore, OMP biogenesis activity is not required for CDI, and thus it remains unclear whether BamA dynamics contribute to toxin translocation.

Putative CdiA-binding epitopes have been localized on BamA and OmpC, but the corresponding receptor-binding regions within CdiA^EC93 and CdiA^EC536 have not yet been identified. Given that BamA and OmpF/OmpC have distinct structures, the two CdiA effectors should contain unique receptor-binding domains. Alignment of CdiA^EC93 and CdiA^EC536 shows several regions of sequence divergence. Prominent among these are the C-terminal toxin region and the sequence corresponding to residues Ser1379 –Tyr1636 of CdiA^EC93 (S4 Fig). Additionally, the CdiA^EC536 effector contains a 69-residue insertion (Ser1988 –Ser2056) relative to CdiA^EC93 (S4 Fig). We recently reported that truncated CdiA^EC93 proteins lacking up to ~1,000 C-terminal residues retain BamA-binding function [7]. Furthermore, CdiA^EC93 lacking internal residues Ala722 –Gly1624 is still secreted and assembled onto the cell surface, but does not bind to BamA [7]. Together, these observations suggest that the receptor-binding domain lies between residues Ser1300 –Asn1600 of CdiA^EC93. There are four distinct classes of E. coli CdiA proteins based on sequence comparisons of the 1300–1600 region, suggesting that there may be additional CDI receptors in E. coli. Alternatively, the other uncharacterized E. coli CdiA effectors could bind to BamA or OmpC/OmpF using epitopes that are distinct from those recognized by CdiA^EC93 and CdiA^EC536 (respectively). Intriguingly, the putative receptor-binding domain is located in the middle of the CdiA primary sequence, between the FHA-1 and FHA-2 peptide repeat regions (S4 Fig). This position is unexpected based on current models of CdiA structure and cell-surface presentation. According to the β-helix structural model [2], the receptor-binding region would be positioned near the center of the CdiA filament rather than the distal tip. It is unclear how a central location is compatible with receptor binding, because the distal end of filament would presumably have to perforate the target-cell outer membrane. Penetration of target bacteria could facilitate toxin delivery, but this model raises the question of why specific receptors should be required for CDI.

Bacterial surface antigens are under intense pressure to diversify, yet E. coli BamA and OmpC proteins show remarkably different responses to this selective pressure. BamA is invariant in over 2,500 E. coli isolates for which sequence information is available. By contrast, E. coli strains encode at least 220 distinct OmpC proteins, many of which are distinguished by unique combinations of extracellular loops L4, L5 and L7. Crystal structures are available for the OmpC^K-12, OmpC^CFT073 and OmpC^6104H porins studied here, and each exhibits a distinct surface topology and electrostatic potential (S5 Fig) [43, 52]. The large number of extracellular loop combinations seen in E. coli OmpC proteins suggests that these elements are modular and can be rearranged independently to generate novel porins. However, OmpC loops L4, L5, L6, L7 and L8 engage in a series of intra-protomer interactions (S4 Table), and therefore some loop combinations may be incompatible with a properly folded porin structure. Most intra-protomer loop interactions (e.g. L5-L7 and L7-L8) are mediated by invariant residues, though the contacts between L4 and L5 in OmpC^CFT073 and OmpC^6104H involve non-conserved residues (S4 Table). We note that chimeric OmpC^CFT073 carrying L4 from OmpC^F11 represents a L4-L5 combination that has yet to be found in any E. coli isolate. Because this OmpC chimera is functional as a CdiA^EC536 receptor, it must be assembled properly in the outer membrane. Therefore, in this instance, loop L4 is clearly modular. This combinatorial complexity has implications for CDI, and almost certainly governs bacterial susceptibility to bacteriophages and immune recognition [53, 54].

CDI receptor polymorphism restricts the number of susceptible target bacteria, yet CdiA^EC536 recognizes multiple OmpC variants. OmpC proteins from E. coli EC93, 536, F11, UTI89 and A33H are all recognized efficiently by CdiA^EC536 though each receptor carries a distinct combination of extracellular loops. E. coli F11 and UTI89 encode CdiA proteins that share >98% identity with CdiA^EC536. Taken together, these observations indicate that the CdiA^F11 and CdiA^UTI89 effectors also bind to OmpF-OmpC and preferentially recognize self-receptors. These findings are consistent with numerous reports that CDI⁺ bacteria tend to auto-aggregate [1, 7, 11, 13, 21–23]. Intriguingly, E. coli CFT073 also encodes a CdiA effector that shares 89.7% identity with CdiA^EC536 over 3,037 N-terminal residues, yet OmpC^CFT073 does not function as a CdiA^EC536 receptor. Inspection of the cdiB gene from E. coli CFT073 reveals that it contains an in-frame stop codon [55], and therefore this strain presumably cannot secrete its CdiA effector. It is tempting to speculate that the loss of CdiA^CFT073 surface expression has allowed ompC ^CFT073 to diverge in response to other selective pressures. However, there are other E. coli isolates with intact cdi loci that also share ompC and ompF alleles with CFT073. Based on the data presented here, it appears that the CdiA proteins from these latter isolates would not preferentially recognize self and therefore may function primarily in competition. Though auto-adhesion is beneficial for bacteria, these phenomena can also be viewed from the perspective of cdi genes as selfish elements. CDI systems are encoded on mobile genetic elements and their toxin/immunity activities act as "gene-addiction" modules [56]. Cells that lose CDI-encoding plasmids or genomic islands are suddenly susceptible to inhibition and risk elimination by neighboring CDI⁺ siblings. Thus, self-recognition would ensure continual selective pressure to retain cdi genes in a population. We speculate that CdiA effectors may have first evolved as adhesins used exclusively for cooperation. Toxin-delivery capabilities could then be grafted onto the ancestral CdiA protein, providing a strong positive selection for cdi retention. Thus, the antagonistic activity of CDI systems may have arisen from evolutionary pressure to retain cdi genes in fluid bacterial genomes.

Materials and Methods

Bacterial strains

Bacterial strains are listed in Table 1. Bacteria were grown in lysogeny broth (LB) or on LB agar unless otherwise noted. Where indicated, media were supplemented with antibiotics at the following concentrations: ampicillin, 150 μg mL⁻¹; kanamycin, 50 μg mL⁻¹; chloramphenicol, 66 μg mL⁻¹; spectinomycin, 50 μg mL⁻¹; rifampicin, 200 μg mL⁻¹; streptomycin, 100 μg mL⁻¹. The bamA::cat allele was introduced into MC4100 pZS21-bamA ⁺ cells by bacteriophage P1-mediated transduction [20]. The ΔompC::kan, ΔompF::kan, Δwzb::kan and ΔrecA::kan disruptions were obtained from the Keio collection [57] and transduced into E. coli strain CH10013. Kanamycin-resistance cassettes were subsequently removed with FLP recombinase expressed from plasmid pCP20 [58]. The ompC ^K-12 gene of E. coli CH10013 was replaced with ompC ^EC536 using Red-mediated recombination. ompC ^EC536 was amplified by PCR with oligonucleotides 3322 and 3323, and a fragment upstream of ompC ^K-12 was amplified using oligonucleotide pair 3320/3321 (all oligonucleotide primers are listed in S5 Table). The two DNA fragments were combined into one fragment by overlap-extension PCR (OE-PCR) using primers 3320/3323 and ligated to pKAN [59] using SacI/BamHI sites to generate plasmid pCH11341. A fragment downstream of ompC ^K-12 was then amplified with primers 3324/3325 and ligated to pKAN using EcoRI/KpnI sites to generate plasmid pCH11342. Fragments from pCH11341 and pCH11342 were amplified with primer pairs 3320/3505 and 3504/3325 (respectively) and the resulting products combined into one fragment by OE-PCR using primers 3320/3325. The final product was introduced into E. coli CH10013 that express phage λ Red proteins from plasmid pSIM6 [60] and recombinant bacteria selected on LB agar supplemented with kanamycin.

Table 1. Bacterial strains.

Bacterial strain	Description a	Reference
EC536	uropathogenic isolate of E. coli	[66]
CFT073	uropathogenic isolate of E. coli	[67]
F11	uropathogenic isolate of E. coli	[68]
UTI89	uropathogenic isolate of E. coli	[69]
6104H	uropathogenic isolate of E. coli	[70]
A54H	uropathogenic isolate of E. coli	[70]
A42H	uropathogenic isolate of E. coli	[70]
A35H	uropathogenic isolate of E. coli	[70]
A33H	uropathogenic isolate of E. coli	[70]
EC93	E. coli isolate from rat feces	[1]
EPI100	F− mcrA Δ(mrr-hsdRMS-mcrBC) φ80dlacZΔM15 ΔlacXcZΔM15 ΔlacX recA1 endA1 araD139 Δ(ara, leu)7697 galU galK λ−rpsL nupG	Epicentre
MFDpir	RP4-2 Tc::[ΔMu1::aac(3)IV-ΔaphA-Δnic35-ΔMu2::zeo] ΔdapA::(erm-pir) ΔrecA	[62]
DL4905	MC4100 λ640–13 PpapBA-gfp-mut3, KanR	[71]
DL6536	E. coli 536 Δkps15::cat ΔaraBAD spc-araC-Para::cdiBAI, CmR StrR SpcR	[8]
DL6381	E. coli 536 Δkps15::cat ΔaraBAD spec-araC-Para::cdiBA(ΔCT)I, CmR StrR SpcR	[8]
CH2016	X90 (DE3) Δrna ΔslyD::kan, KanR	[72]
CH10013	spontaneous rifampicin-resistant isolate of JCM158, RifR	[73]
CH10099	CH10013 ΔompC	This study
CH10100	CH10013 ΔompF	This study
CH10136	CH10013 ΔompC ΔompF	This study
CH10226	MC4100 Δbam::cat pZS21amp-bamA Eco, AmpR	This study
CH10227	MC4100 Δbam::cat pZS21amp-bamA ECL, AmpR	This study
CH11346	CH10013 Δwzb ΔompF ΔompC ΔrecA, RifR	This study
CH12133	CH10013 ompC EC536 -kan, KanR	This study
CH12134	CH10013 ΔompF ompC EC536 -kan, KanR	This study
CH12180	CH10013 Δwzb ΔompF ompC EC536 ΔrecA::kan, RifR KanR	This study
CH12345	JCM158 Δwzb ΔompF ΔompC ΔrecA::kan, KanR	This study

^aAbbreviations: Amp^R, ampicillin resistant; Cm^R, chloramphenicol resistant; Kan^R, kanamycin resistant; Rif^R, rifampicin resistant; Spc^R, spectinomycin resistant; Str^R, streptomycin resistant

Plasmid constructions

Plasmids constructs used in this study are listed in Table 2. The E. coli cysK gene was amplified with primer pair 2111/2110 and ligated to pBR322 using EcoRV and BamHI restriction sites to generate plasmid pCH9763. The ompF ^K-12 allele was amplified from E. coli JCM158 with primer pair 2385/2386, and the fragment ligated to pZS21 using BamHI and XbaI restriction sites to generate plasmid pCH10202. Additionally, ompF ^K-12 and ompF ^EC536 were amplified with primers 2729/2734, and the fragments ligated to pTrcKX using KpnI and XhoI restriction sites to generate plasmids pCH11850 and pCH10780. The ompF genes from S. Typhimurium LT2 and E. cloacae ATCC 13047 were amplified with primer pairs 3468/3469 and 2729/3470 (respectively) and ligated to pTrcKX as described for the E. coli genes. The ompC genes from S. Typhimurium LT2 and E. cloacae were amplified with primers 3326/2388, digested with NcoI/XhoI and ligated to plasmid pTrc99a to generate pCH11852 and pCH11853, respectively. All E. coli ompC alleles were amplified with primer pair 2387/2388, followed by ligation to plasmid pTrcKX using KpnI and XhoI restriction sites. The coding sequences for OmpC^CFT073 loops L4, L5, L7 and L8 of OmpC^CFT073 were replaced using megaprimer PCR [61]. Plasmid pCH10778 template DNA was amplified with reverse primer 2388 in conjunction with primers 2935 (L5^EC536), 2936 (L7^EC536) and 2937 (L8^EC536). The resulting products were used as megaprimers to amplify ompC ^CFT073 in conjunction with forward primer 2387. The chimeric products were digested with KpnI/XhoI and ligated to pTrcKX to generate plasmids pCH11334, pCH11765 and pCH11335. The same procedure and primers were used to generate ompC ^CFT073 constructs encoding two loops from ompC ^EC536. The loop L4 exchange megaprimer was generated with primers 2387/2807 and used with 2388 to generate the final product. All PCR products were digested with KpnI and XhoI and ligated to plasmid pTrcKX.

Table 2. Plasmid constructs.

Plasmids	Description a	Reference
pTrc99a	IPTG-inducible expression vector, AmpR	Amersham
pTrcKX	Derivative of pTrc99a with additional KpnI and XhoI restriction sites, AmpR	[74]
pCP20	temperature-inducible expression of FLP recombinase, CmR AmpR	[58]
pSC189kan	Mobilizable plasmid with R6Kγ replication origin. Carries the mariner transposon containing a kanamycin-resistance cassette, AmpR KanR	[63]
pWEB::TNC	pWEB::TNC (CDI–), AmpR, CmR	Epicentre
pDAL660Δ1–39	Constitutively expresses the E. coli EC93 cdiBAI gene cluster, AmpR	[1]
pDAL866	Arabinose-inducible expression of the E. coli 536 cdiBAI gene cluster, AmpR CmR	[32]
pZS21	pSC101-derived plasmid vector, KanR	[75]
pZS21amp	pZS21 derivative with ampicillin-resistance cassette, AmpR	[4]
pZS21amp-bamA Eco	Constitutive expression of E. coli bamA, AmpR	[4]
pZS21amp-bamA ECL	Constitutive expression of E. cloacae bamA, AmpR	[20]
pET21::colE5-immE5	Over-produces colicin E5 and ImmE5-His6, AmpR	[74]
pCH450	pACY184 derivative that carries E. coli araC and the L-arabinose-inducible PBAD promoter. TetR	[59]
pCH9181	pTrcKX::cdiA-CT/cdiI Dd3937-his 6, AmpR	[76]
pCH9763	pBR322::cysK, AmpR	This study
pCH10138	pTrcKX::ompC K-12, AmpR	This study
pCH10201	pTrcKX::ompC F11, AmpR	This study
pCH10202	pZS21-ompF K-12, KanR	This study
pCH11338	pTrcKX::ompC EC536, AmpR	This study
pCH10458	pTrcKX::ompC EC869, AmpR	This study
pCH10460	pTrcKX::ompC EC93, AmpR	This study
pCH10778	pTrcKX::ompC CFT073, AmpR	This study
pCH10779	pTrcKX::ompC UTI89, AmpR	This study
pCH10780	pTrcKX::ompF EC536, AmpR	This study
pCH10796	pTrcKX::link8-ompF, AmpR	This study
pCH10797	pTrc99a::ompF 2, AmpR	This study
pCH10959	pTrcKX::ompC(L4F11)CFT073, AmpR	This study
pCH11330	pTrcKX::ompC 6104H, AmpR	This study
pCH11331	pTrcKX::ompC A54H, AmpR	This study
pCH11332	pTrcKX::ompC A42H, AmpR	This study
pCH11333	pTrcKX::ompC A35H, AmpR	This study
pCH11341	pKAN-micF-ompC EC536, AmpR KanR	This study
pCH11342	pKAN-abpE´, AmpR KanR	This study
pCH11496	pCH450::DsRed; arabinose-inducible expression of DsRed, TetR	This study
pCH11627	pTrcKX::ompC A33H, AmpR	This study
pCH11334	pTrcKX::ompC(L5536)CFT073, AmpR	This study
pCH11335	pTrcKX::ompC(L8536)CFT073, AmpR	This study
pCH11336	pTrcKX::ompC(L5/L7536)CFT073, AmpR	This study
pCH11337	pTrcKX::ompC(L7/L8536)CFT073, AmpR	This study
pCH11765	pTrcKX::ompC(L7536)CFT073, AmpR	This study
pCH11767	pTrcKX::ompC(L5/L8536)CFT073, AmpR	This study
pCH11850	pTrcKX::ompF K-12, AmpR	This study
pCH11851	pTrcKX::ompF ECL, AmpR	This study
pCH11852	pTrcKX::ompF LT2, AmpR	This study
pCH11853	pTrcKX::ompC ECL, AmpR	This study
pCH11854	pTrcKX::ompC LT2, AmpR	This study
pCH12054	pTrc99a::ompF 2*, AmpR	This study

^aAbbreviations: Amp^R, ampicillin resistant; Cm^R, chloramphenicol resistant; Kan^R, kanamycin resistant; Tet^R, tetracycline resistant

Dimeric OmpF expression plasmids were constructed through sequential ligation of ompF coding sequences. The ompF ^EC536 allele was amplified with primers 2733/2734 and ligated to pCH9181 using SpeI/XhoI restriction sites to generate intermediate construct pCH10796. A second ompF fragment was amplified with primers 2732/2729 and ligated to pCH10796 using KpnI/SpeI restriction sites. The resulting pCH10797 (ompF ₂) construct produces OmpF^EC536 dimers linked by a Ser-Gly-Thr-Thr-Ser-Thr-Gly-Gly peptide. To introduce a longer linker sequence, ompF was amplified with primers 2729/2765 and ligated to pCH9181 using KpnI/SpeI sites to generate plasmid pCH11893. The second ompF module was amplified with primers 3008/2734 and ligated to pCH11893 using BamHI/XhoI restriction sites. The resulting pCH12054 (ompF ₂*) construct produces OmpF^EC536 dimers linked by a Ser-Gly-Thr-Gly-Ser-Asp-Thr-Ser-Gly-Gly-Thr-Asp-Gly-Thr-Gly-Gly peptide.

Competition co-cultures

Inhibitor and target bacteria were grown to mid-log phase in LB media supplemented with the appropriate antibiotics, then mixed at a 10:1 ratio in fresh LB medium without antibiotics and incubated for 4–5 h at 37°C with vigorous shaking in baffled flasks. Viable target-cell counts were enumerated on selective LB-agar as colony forming units (cfu) per mL. E. coli EPI100 carrying pDAL660Δ1–39 [1] or pDAL866 [33] were used as inhibitor cells for the comparison of CDI^EC93 and CDI^EC536 systems. The mock CDI⁻inhibitors were E. coli EPI100 carrying the pWEB::TNC vector. All other competitions used the E. coli 536 derivative DL6536, which expresses cdiBAI ^EC536 under the control of an arabinose-inducible promoter [8]. The mock inhibitor strain DL6381 is a derivative of DL6536 that carries a deletion of the cdiA-CT coding sequence [8]. DL6536 and DL6381 were cultured in 0.2% arabinose to induce cdiBAI expression and competition co-cultures were supplemented with 0.2% arabinose.

Transposon library construction and selection for CDI^R mutants

Plasmid pSC189 (encoding the mariner transposon) was introduced into E. coli CH10013 cells carrying pCH9763 (pBR322::cysK) through conjugation with E. coli MFDpir donors [62, 63]. Donor and recipient cells were grown to mid-log phase in LB media supplemented with ampicillin and 30 μM diaminopimelic acid (for donors). Donors (~6.0 ×10⁸ colony forming units, cfu) and recipients (~3 ×10⁸ cfu) were mixed and collected by centrifugation for 2 min at 9,000 rpm in a microcentrifuge. The supernatant was removed by aspiration and the cell pellet resuspended in 100 μL of 1× M9 salts. Cell mixtures were spotted onto 0.45 μm nitrocellulose membranes and incubated on LB-agar without inversion for 4 h at 37°C. Cells were then harvested into 2 mL of 1× M9 salts and transposon-insertion mutants selected on LB-agar supplemented with rifampicin and kanamycin. Over 50,000 colonies from each mating were harvested into 1× M9 salts, and inoculated into 50 mL of LB medium in a 250 mL baffled flask. Inhibitor cells (E. coli 536 derivative DL6536) were grown in parallel in LB medium supplemented with L-arabinose until mid-log phase. Inhibitors and targets were mixed at a 10:1 ratio in fresh LB medium supplemented with L-arabinose and cultured for 4 h with shaking at 37°C. Viable target cells were enumerated as cfu/mL on LB agar supplemented with rifampicin and kanamycin. Survivors from the first round of CDI^EC536 selection were harvested into 1× M9 salts and inoculated into 50 mL of LB medium supplemented with L-arabinose for a second round of selection. After the third round of selection, target-cell populations were completely resistant to the CDI inhibitor cells. Individual colonies were screened for CysK activity in MOPS minimal media containing 3 mM 1,2,4-triazole, which inhibits the growth of cysK ⁺ bacteria [64]. CDI^R phenotypes for all cysK ⁺ clones were confirmed in competition co-cultures with E. coli 536 inhibitors. Transposon mutations were transduced into CDI-sensitive cells, and the resulting transductants tested in competition co-cultures for CDI^R phenotypes.

Flow cytometry analysis of cell-cell adhesion

Overnight cultures of GFP-labeled E. coli DL4905 cells carrying pDAL660Δ1–39 (CDI^EC93), pDAL866 (CDI^EC536) or pWEB::TNC (CDI⁻) were diluted into fresh tryptone broth (TB) and grown to mid-log phase at 30°C. Inhibitor cells were then mixed at a 5:1 ratio with DsRed-labeled E. coli CH10099 (ΔompC) or CH10100 (ΔompF) cells complemented with the indicated ompC and ompF expressing plasmids. Cell suspensions were incubated with aeration for 15 min at 30°C, diluted 1:50 into filtered 1× phosphate-buffered saline (PBS), and then analyzed on an Accuri C6 flow cytometer using FL1 (533/30 nm, GFP) and FL2 (585/40 nm, DS-Red) fluorophore filters (Becton Dickinson) as described [36].

Immunoblot analysis

Bacteria were grown to mid-log phase in LB media supplemented with ampicillin. Cells were collected by centrifugation at 6,000 ×g. Cell were frozen at –80°C, then broken by freeze-thaw cycles in 8 M urea, 150 mM NaCl, 50 mM Tris-HCl (pH 8.0). Extracts were quantified by Bradford assay and equal amounts of protein were resolved on 6 M urea/10% SDS polyacrylamide gels for 4 h at 100 V. Gels were electroblotted onto nitrocellulose membranes and OMPs detected using polyclonal antisera raised against E. coli OmpC/OmpF and BamA. Immunoblots were visualized using IRDye 680 (LI-COR) labeled anti-rabbit secondary antibodies and an Odyssey infrared imager.

Colicin E5 purification and activity assays

Colicin E5 was purified in complex with its His₆-tagged immunity protein as described previously [6, 65]. E. coli strains CH12180 and CH12345 carrying pCH11850 (ompF), pCH10796 (ompF-ompF, 8-residue linker), or pCH12054 (ompF-ompF, 16-residue linker) were grown to mid-log phase in LB media supplemented with ampicillin, then collected by centrifugation and re-suspended in fresh pre-warmed media at OD₆₀₀ = 0.05. After 30 min of incubation with shaking, purified colicin E5 was added to a final concentration of 1 μM and cell growth monitored by measuring the OD₆₀₀ every 60 min.

Supporting Information

S1 Fig. Alignment of OmpC sequences from selected E. coli isolates.

The sequences of mature OmpC proteins were aligned using Clustal-Omega and identical residues indicated with asterisks (*). Extracellular loop sequences are shown in red and β-strands in boldface.

(PDF)

S2 Fig. Inter-protomer interactions for OmpC^K-12 and OmpF^K-12.

The sequences of mature OmpC^K-12 and OmpF^K-12 were aligned using Clustal-Omega and identical residues indicated with asterisks (*). Extracellular loops are shown in red font and β-strands are underlined. Buried interfacial residues are shown in blue, and residues involved in direct inter-protomer H-bonds and salt-bridges are in orange. Contacts were determined with PDBePISA using PDB:2J1N (OmpC^K-12) and PDB:3POX (OmpF^K-12).

(PDF)

S3 Fig. Inter-protomer contacts are conserved between OmpC^K-12 and OmpF^K-12.

Individual protomers of (A) OmpC^K-12 (PDB:2J1N) and (B) OmpF^K-12 (PDB:3POX) are viewed from the inter-protomer interface (left) and from the extracellular milieu (right). Extracellular loops L2 and L4 from adjacent protomers are shown as yellow sticks and β-strands involved inter-subunit contacts are labeled in the left panels. Buried interfacial residues are shown in blue, and residues involved in direct inter-protomer H-bonds and salt-bridges are in orange. Contacts were determined with PDBePISA. The conserved inter-subunit ion-pairs are shown in the right panels with residues labeled.

(TIF)

S4 Fig. Alignment of CdiA^EC93 and CdiA^EC536.

CdiA^EC93 and CdiA^EC536 sequences were aligned using Clustal-Omega and identical residues are highlighted in blue. Predicted domains and peptide motifs are as described by the InterPro web server. CdiA^EC93 annotations are available at http://www.ebi.ac.uk/interpro/protein/Q3YL96, and CdiA^EC536 at http://www.ebi.ac.uk/interpro/protein/Q0T963. The signal sequence and TPS transport domain are required for CdiA export. FHA-1 (Pfam: PF05594) and FHA-2 (PF13332) peptide repeats are predicted to form a β-helix. The pretoxin-VENN domain (PT-VENN; PF04829) demarcates the variable C-terminal toxin region.

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S5 Fig. Comparison of E. coli OmpC structures.

Individual OmpC protomers from E. coli K-12 (PDB:2J1N), CFT073 (PDB:2XE1) and 6104H (PDB:2XE2) are shown in surface representation as viewed from the extracellular milieu. Extracellular loops L4 (maroon), L5 (green) and L7 (orange) are labeled in the top row of images. The bottom row shows the OmpC protomers in the same orientation, but with the surface rendered as an electrostatic potential map. Red areas are electronegative, blue are electropositive and white are neutral.

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S1 Table. Predicted OmpC proteins from E. coli isolates.

OmpC from E. coli str. K-12 substr. MG1655 was used to query all E. coli isolates using BLASTP 2.2.32+ at https://blast.ncbi.nlm.nih.gov/Blast.cgi. Proteins with identical amino acid sequences are grouped with colored backgrounds. Extracellular loop L4, L5 and L7 variations are indicated according the scheme outlined in S2 Table. Asterisks (*) indicate OmpC variants tested in this study.

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S2 Table. E. coli OmpC loop L4, L5 and L5 sequences.

Loop sequences were determined for 2707 predicted E. coli OmpC proteins. Sequences were grouped and numbered according to sequence identity. Red residues are shared with OmpC from E. coli K-12.

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S3 Table. OmpC and OmpF interprotomer H-bonds and salt-bridges.

Predicted hydrogen-bonds (H-bonds) and salt-bridges between individual protomers of E. coli K-12 OmpC (PDB: 2J1N) and OmpF (PDB: 3POX) were determined using PDBePISA (http://www.ebi.ac.uk/pdbe/pisa/). Solvent accessible surface areas (ASA) and buried surface areas (BSA) were determined using PDBePISA (http://www.ebi.ac.uk/pdbe/pisa/). Residues highlighted in orange are buried at interprotomer interfaces. Predicted hydrogen bonds are indicated by H and salt-bridges by SH.

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S4 Table. OmpC intraprotomer loop-loop interactions.

Extracellular loop residues involved in intraprotomer loop-loop interactions are presented for three E. coli OmpC proteins. Loop L4, L5 and L7 sequences are labeled according to the scheme outlined in S2 Table.

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S5 Table. Oligonucleotides used in this study.

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Data Availability

All relevant data are within the paper and its Supporting Information files.

Funding Statement

This work was supported by grant R01 GM117930 (CSH) from the National Institutes of Health. CMB was supported in part by Dean's and Chang Fellowships from the University of California, Santa Barbara, and JLEW was supported by National Science Foundation Graduate Research Fellowship DGE-1144085. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.