Abstract
Bacteria use extracellular appendages called type IV pili (T4P) for diverse behaviors including DNA uptake, surface sensing, virulence, protein secretion, and twitching motility1. Dynamic extension and retraction of T4P is essential for their function, yet little is known about the mechanisms controlling these dynamics or the extent to which their regulation is conserved across bacterial species. Here, we develop Acinetobacter baylyi as a new model to study T4P by employing a recently developed pilus labeling method2,3. Our findings overturn the current dogma that T4P extension occurs through the action of a single, highly conserved motor, PilB, by showing that T4P synthesis in A. baylyi is dependent on an additional, phylogenetically distinct motor, TfpB. Furthermore, we uncover an inhibitor of T4P extension that specifically binds to and inhibits PilB but not TfpB. These results expand our understanding of T4P regulation and highlight how inhibitors might be exploited to inhibit T4P synthesis.
Introduction
T4P are thin, proteinaceous appendages that are broadly distributed throughout bacteria and archaea4. T4P are composed primarily of major pilin protein subunits that are polymerized or depolymerized through the activity of ATPases to mediate fiber extension and retraction, respectively5. T4P are subdivided into subcategories (generally T4aP, T4bP, and T4cP) based on protein homology and pilus function, with T4aP being the best-characterized. In the T4aP systems found in Gram-negative organisms, polymerization and depolymerization of pilins occurs through interactions of the extension ATPase PilB or retraction ATPase PilT with the integral inner membrane platform protein PilC (Supplemental figure 1a). The growing fiber spans an alignment complex from the inner membrane through the periplasm composed of PilNOP to exit through the PilQ outer membrane secretin pore. Dynamic cycles of T4P extension and retraction are critical for the diverse processes that these structures mediate including twitching motility6, surface sensing2,7, virulence8,9, and DNA uptake10,11.
Acinetobacter species like A. baumannii and A. nosocomialis have emerged to become an urgent medical threat due to their prevalence in hospital-acquired infections and their capacity to acquire antibiotic resistance genes; a process that is achieved in part by natural transformation through T4aP-mediated DNA uptake12,13. Acinetobacter baylyi is the most naturally transformable species reported to date14, with up to 50% of cells undergoing natural transformation in laboratory conditions, making it an ideal candidate to study T4P-mediated DNA uptake and natural transformation.
Results
To study A. baylyi T4P, we applied a recently-developed labeling method2,3 by targeting the major pilin, ComP, for cysteine substitution and subsequent labeling with thiolreactive maleimide dyes. Maleimide-labeling of the functional comPT129C strain (Supplemental figure 1b, 1c) revealed external T4P filaments as seen in other species using this method3. T4P in A. baylyi appear much shorter than those found in other species like V. cholerae, and they localize close together in a line along the long axis of the cell (Supplemental figure 1c). Incubation of cells with fluorescently-labeled DNA resulted in co-localization of DNA with T4P, which is consistent with the essential role of T4P-DNA binding during natural transformation in Vibrio cholerae11 (Figure 1a). We reasoned that natural transformation could be used to screen for other factors that regulate T4P synthesis in A. baylyi. To that end, we performed a high throughput transposon-sequencing screen (Tn-seq)15 to identify genes required for natural transformation. In this screen, loss of known T4P-related genes resulted in negative selection, indicating that they were critical for natural transformation as expected (Figure 1b). To validate our Tn-seq results, we made in-frame deletions of representative genes from each T4P-encoding operon, as well as genes known to be essential for natural transformation that act downstream of T4P. Mutations in the T4P platform protein gene pilC, the pilus regulatory gene pilY1, the outer membrane secretin gene pilQ and the retraction ATPase gene pilT all showed a marked reduction in natural transformation, with most mutants exhibiting transformation rates below our limit of detection (Figure 1c). T4P-labeling of pilC, pilY1, and pilQ mutants revealed no visible T4P fibers (Figure 1d). Mutations in genes that act downstream of T4P, including the periplasmic DNA-binding protein gene comEA, the inner membrane DNA-transporter protein gene comEC, and the DNA-recombination helicase gene comM likewise resulted in a reduction in transformation frequency, however, these strains still produced T4P fibers as expected (Figure 1c, d).
We next sought to determine whether any of the previously uncharacterized genes from our Tn-seq screen reduced natural transformation in A. baylyi by affecting T4P synthesis. Tn-seq revealed that mutations in pilB, the canonical extension ATPase gene that is typically cotranscribed with pilC and the pre-pilin peptidase, pilD, resulted in reduced natural transformation (Figure 1b). We also found an additional pilB homologue that likewise exhibited lower rates of transformation (Figure 1b). We named this new PilB homologue TfpB for “type four pilus PilB-like protein” because although it has homology to PilB, it does not exhibit the same gene synteny as pilB genes that are typically co-transcribed with their cognate inner membrane platform gene pilC. Surprisingly, deletion of the canonical pilB did not ablate transformation as would be expected based on homology to other T4P10 (Figure 2a). Mutation of tfpB reduced transformation rates to a level that was similar to the pilB mutant; however, natural transformation was undetectable in the pilB tfpB double mutant (Figure 2a).
Deletion of pilT prevents T4P retraction16 and may thus reveal more subtle effects on pilus extension. T4P labeling in pilB, tfpB, and pilB tfpB mutants revealed that all three mutants were defective in T4P synthesis while pilT was intact (Figure 2c). In a pilT deletion background, pilB mutant populations had similar numbers of cells producing T4P compared to the ΔpilT parent while, surprisingly, tfpB mutants were highly defective in T4P synthesis (Figure 2b, c). The pilB tfpB pilT triple mutant produced no detectable T4P fibers (Figure 2b, c). Together, these results indicate that both PilB and TfpB are essential for efficient T4P extension in A. baylyi, with TfpB playing the dominant role.
Phylogenetic analysis of extension ATPase homologues found among Gammaproteobacteria4,17, which include members of the PilT/VirB11 family of secretion ATPases, revealed that TfpB clusters with a group of proteins that are phylogenetically distinct from the canonical PilB ATPase (Figure 2d, Supplemental figure 2). PilB and TfpB proteins are as divergent from one another as PilB and the type II secretion system proteins XcpR/GspE, or PilB and the MshE motors that drive mannose-sensitive haemagglutinin (MSHA) pilus synthesis, suggesting that TfpB evolved as a functionally divergent class of proteins that is distinct from canonical PilB extension motors. TfpB is highly conserved in other Acinetobacter species (Figure 2d, Supplemental figure 2), implying that use of multiple extension motors may be prevalent in the Acinetobacter clade. These results suggest that proteins that cluster with TfpB may play similar functions in other bacterial species and that multiple extension ATPases may be a common feature of diverse T4P.
In addition to revealing the importance of TfpB in T4P synthesis, our Tn-seq results also uncovered an uncharacterized transcriptional regulator belonging to the XRE-family of transcriptional repressors that is critical for natural transformation (Figure 1b, Figure 3a). Deletion of this regulator, named cpiR for competence pilus inhibition repressor, resulted in a 100-fold reduction in transformation (Figure 3b). We hypothesized that CpiR repressed a factor that inhibits natural transformation. Transcriptional repressors are often transcribed immediately adjacent to the genes they repress. We thus deleted the upstream gene, named cpiA for competence pilus inhibition actuator (Figure 3a), and found that transformation frequency was restored in the cpiR cpiA double mutant background (Figure 3b). Expression of cpiA under the control of an IPTG-inducible promoter (Ptac) was sufficient to reduce transformation frequency even in the presence of cpiR, further suggesting that CpiR represses cpiA transcription. Furthermore, we found that CpiR was sufficient to repress the cpiA promoter (using a PcpiA-GFP reporter) in the heterologous host Vibrio cholerae, which suggests that CpiR is a direct repressor of PcpiA (Figure 3c). Finally, we found that CpiA protein is only produced in a cpiR mutant (Figure 3d). These results demonstrate that under lab conditions, CpiR represses cpiA transcription to allow for high rates of natural transformation; however, in the absence of CpiR, CpiA is produced and natural transformation is inhibited ~100-fold.
CpiA lacks primary sequence or structural homology to any proteins or domains of known function18,19, so we next sought to determine the mechanism of transformation inhibition by CpiA. Labeling of T4P in a cpiR mutant (i.e. when CpiA is expressed) revealed that cells were deficient in T4P synthesis (Figure 4a). This defect was dependent on cpiA because piliation was restored in the cpiR cpiA double mutant (Figure 4a). Deletion of pilT restored T4P synthesis in the cpiR mutant (i.e. the cpiR pilT double mutant) (Figure 4a, b). This phenotype was reminiscent of the restoration of piliation observed in the pilB pilT double mutant, so we hypothesized that CpiA may regulate natural transformation by inhibiting PilB.
To test if CpiA functions by inhibiting PilB, we made pilB cpiR and tfpB cpiR double mutants. T4P synthesis was unaffected in pilB cpiR pilT (where TfpB is the sole extension ATPase present), while no T4P were detected in tfpB cpiR pilT (where PilB is the sole extension ATPase present) (Figure 4a, b). The defect in T4P synthesis in the tfpB cpiR pilT strain was restored when cpiA was deleted in this background, demonstrating that CpiA acts as a PilB inhibitor to control natural transformation (Figure 4a, b). Transformation frequency assays corroborated these results by demonstrating that deletion of cpiR only inhibited transformation in the tfpB mutant background (where PilB is the sole extension ATPase) and not the pilB mutant background (where TfpB is the sole extension ATPase) (Figure 4c). Immunoblotting revealed that CpiA does not affect PilB production or stability (Figure 3d), and we thus hypothesized that CpiA may interact with PilB to disrupt its function. Coimmunoprecipitation experiments with functional fusion proteins (Supplemental figure 3) revealed that CpiA and PilB interact with each other, and that their interaction is not dependent on the pilus machinery proteins PilMNOPQ (Figure 4d and Supplemental figure 4), ruling out indirect binding of CpiA to PilB through other machinery components required for T4P synthesis. CpiA and TfpB do not interact (Supplemental figure 4), consistent with our genetic evidence that CpiA specifically inhibits PilB and not TfpB (Figure 4a-c). Together, these data suggest that CpiA directly binds to and inhibits PilB.
Discussion
Environmental conditions play an integral role in regulating how bacteria respond to and interact with their surroundings. The mechanisms of T4P synthesis regulation identified here likely reflect how environmental conditions influence bacterial physiology. The acquisition of the additional extension motor TfpB and the PilB-specific inhibitory protein CpiA likely resulted from a need to modulate T4P extension under different environmental conditions. Bacteria generally limit T4P synthesis to environments where they provide a selective advantage. For example, in V. cholerae, the production of competence T4aP requires chitin and quorum sensing20, which are environmental conditions that cells experience when they are likely to encounter other bacterial cells for horizontal gene transfer; while the toxin-coregulated T4bP (TCP) required for intestinal colonization are produced in response to host-specific cues21. In A. baylyi, both PilB and TfpB are required for T4P extension and, consequently, efficient natural transformation. While CpiA is not expressed under laboratory conditions (due to CpiR repression), it is possible that different environmental conditions derepress cpiA to decrease pilus activity.
Secretion ATPases like PilB belong to a broadly distributed class of proteins that play essential roles in diverse microbial behaviors, yet there are few known inhibitors of these proteins. The ones that are characterized are encoded by phages that use T4P to infect cells, and it is speculated that PilB inhibition by these proteins prevents superinfection by other T4P-dependent phages22,23. In Acinetobacter baylyi, cpiA is encoded on the chromosome. Thus, it is tempting to speculate that CpiA was acquired after phage infection of an ancestral strain and co-opted for regulation of T4P activity. A better understanding of secretion ATPase function and the evolution of mechanisms by which they can be inhibited may enable the development of tools to control their activity. Precise control of T4P synthesis may provide a means to manipulate behaviors that require T4P like biofilm formation, virulence, and natural transformation, which are clinically relevant in diverse pathogens.
Methods
Bacterial strains and culture conditions
Acinetobacter baylyi strain ADP1 was used throughout this study. For a list of strains used throughout, see Supplemental Table 1. A. baylyi cultures were grown at 30 °C in Miller lysogeny broth (LB) medium and on agar supplemented with kanamycin (50 μg/mL), spectinomycin (60 μg/mL), gentamycin (30 μg/mL), and/or chloramphenicol (30 μg/mL), rifampicin (30 μg/mL), zeocin (100 μg/mL), and/or streptomycin (10 μg/mL) as appropriate.
Construction of mutant strains
Mutants in A. baylyi were made using natural transformation as described previously16. Briefly, mutant constructs were made by splicing-by-overlap (SOE) PCR to stich 1) ~3 kb of the homologous region upstream of the gene of interest, 2) the mutation where appropriate (for deletion by allelic replacement with an antibiotic resistance cassette, or the fusion protein), and 3) the downstream region of homology. For a list of primers used to generate mutants in this study, see Supplemental Table 2. The upstream region was amplified using F1 + R1 primers, and the downstream region was amplified using F2 + R2 primers. All antibiotic resistance (AbR) cassettes were amplified with ABD123 (ATTCCGGGGATCCGTCGAC) and ABD124 (TGTAGGCTGGAGCTGCTTC). Fusion proteins were amplified using the primers indicated in Supplemental Table 2. In-frame deletions were constructed using F1 + R1 primer pairs to amplify the upstream region and F2 + R2 primer pairs to amplify the downstream region with ~20 bp homology to the remaining region of the downstream region built into the R1 primer and ~20 bp homology to the upstream region built into the F2 primer. SOE PCR reactions were performed using a mixture of the upstream and downstream regions, and middle region where appropriate using F1 + R2 primers. SOE PCR products were added with 50 μl of overnight-grown culture to 450 μl of LB in 2 ml round-bottom microcentrifuge tubes (USA Scientific) and grown at 30 °C rotating on a roller drum for 1-3 hours. For AbR-constructs, transformants were serially diluted and plated on LB and LB + antibiotic. For in-frame deletions and protein fusion constructs, after the three-hour incubation, cells were incubated with 10 μl of DNaseI (New England Biolabs) for 5 min at room temperature before 450 μl of LB was added. Transformations were grown for an additional hour before cells were diluted and 100 μl of 10-6 dilution was plated on LB plates. In-frame deletions were confirmed by PCR using primers ~150 bp up and downstream of the introduced mutation, and fusions were confirmed by sequencing.
Mutants in V. cholerae were made by chitin-induced natural transformation exactly as previously described24,25. SOE products were generated exactly as described above. The chromosomally integrated PBAD ectopic expression construct is described in detail in ref 26.
Natural transformation assays
Assays were performed exactly as previously described27. Briefly, strains were grown overnight in LB broth at 30°C rolling. Then, ~108 cells were subcultured into fresh LB medium and 100 ng of tDNA was added. In this study, a ΔACIAD1551::SpecR PCR product (with 3kb arms of homology) was used as the tDNA. Reactions were incubated with end-over-end rotation at 30°C for 5 hours and then plated for quantitative culture on spectinomycin plates (to quantify transformants) and on plain LB plates (to quantify total viable counts). Data are reported as the transformation frequency, which is defined as the (CFU/mL of transformants) / (CFU/mL of total viable counts).
Tn-seq Analysis
Transposon mutant libraries of A. baylyi were generated by electroporation of cells with pDL1093, a vector that allows for KanR mini-Tn10 mutagenesis28,29, and plating on kanamycin plates. Approximately 100,000 Tn mutants were scraped off of plates and pooled to generate the input transposon mutant library. This mutant library was then subjected to natural transformation in three separate replicates exactly as described above using three different sources of tDNA: a ΔACIAD1551::SpecR PCR product, an RpsL K43R (SmR) PCR product, or an RpoB PCR product from a spontaneous rifampicin resistant mutant. Following incubation with tDNA, reactions were split and outgrown in selective medium overnight (i.e. with the appropriate antibiotic added to select for transformants = “transformant pool”) or grown overnight in nonselective medium (i.e. in plain LB medium = “total pool”). Sequencing libraries of the transposon-genomic junctions were generated for the Illumina platform using HTM-PCR exactly as previously described29,30. Sequencing data were mapped to the V. cholerae N16961 genome31 and the relative abundance of Tn insertions in all samples was determined on the Galaxy platform32. Comparative analyses for Tn-seq were performed treating the “transformant pool” as the output and the “total pool” as the input to determine genes that were over- and under-represented following selection for transformants. Gene fitness was only assessed if a gene contained at least 1 transposon insertion in at least two out of the three replicates of the “total pool” samples. Also, the normalized abundance of insertions within a gene had to be greater than 0.01 in the “total pool” when averaged across all three replicates (where 1 is the expected normalized abundance for a “neutral” gene). These cutoffs allowed us to assess the phenotype of ~83% of the genes (2747/3310) in the A. baylyi ADP1 genome. For visualization, the relative abundance of Tn insertions within each gene is plotted from the “transformant pool” relative to the “total pool”.
Pilin labeling, imaging, and quantification
Pilin labeling was performed as described previously with some changes2,3. Briefly, 100 μl of overnight-grown cultures was added to 900 μl of LB in 1.5 ml microcentrifuge tube, and cells were grown at 30 °C rotating on a roller drum for 70 min. Cells were then centrifuged at 18,000 x g for 1 min and then resuspended in 50 μl of LB before labeling with 25 μg/ml of AlexaFluor488 C5-maleimide (AF488-mal) (ThermoFisher) for 15 min at room temperature. Labeled cells were centrifuged, washed once with 100 μl of LB without disrupting the pellet, and resuspended in 5-20 μl LB. Cell bodies were imaged using phase contrast microscopy while labeled pili were imaged using fluorescence microscopy on a Nikon Ti-2 microscope using a Plan Apo 100X objective, a GFP filter cube for pili, a Hamamatsu ORCAFlash4.0 camera, and Nikon NIS Elements Imaging Software. Cell numbers and the percent of cells making pili were quantified manually using ImageJ33. All imaging was performed under 1% UltraPure agarose (Invitrogen) pads made with phosphate-buffered saline (PBS) solution.
Western blotting
~109 Cells from overnight cultures were concentrated into a pellet by centrifugation, and the culture supernatant was removed. Cell pellets were resuspended in 50 μl PBS and then mixed with an equal volume of 2×SDS-PAGE sample buffer (125 mM Tris, pH 6.8, 20% glycerol, 4% SDS, 0.4% bromophenol blue, and 10% β-mercaptoethanol) and boiled using a heat block set to 100 °C for 10-15 min. Proteins were separated on a 4-20% pre-cast polyacrylamide gel (Biorad) by SDS electrophoresis, electrophoretically transferred to a nitrocellulose membrane, and probed with mouse monoclonal α-FLAG antibodies (Sigma), mouse monoclonal α-GFP and/or mouse monoclonal α-RpoA (BioLegend) primary antibodies. Blots were then incubated with α-mouse IRDye secondary antibodies (Licor) and imaged using a Licor imaging system.
Coimmunoprecipitation (pulldown) experiments
Overnight cultures of cells grown in tubes in LB medium were diluted by 1/10 into fresh LB for a total volume of 30 ml or 50 ml in 125 ml or 250 ml volume flasks respectively. 30-50 ml cultures were grown to exponential growth phase by shaking for 1.5 hr at 30 °C. The total culture volume was then harvested at 10,000 x g for 10 min at room temperature, and the supernatant was removed. Cell pellets were resuspended in 2 ml of Buffer 1 (50 mM Tris-Cl pH 7.4, 150 mM NaCl, 1 mM EDTA) and transferred to 2 ml volume microcentrifuge tubes and centrifuged at 18,000 x g for 1 min. Cells were washed once more with 2 ml of Buffer 1, and washed pellets were resuspended in 1 ml of Buffer 2 (50 mM Tris-Cl pH 7.4, 150 mM NaCl, 1 mM EDTA, 10 mM MgCl2, 0.1% Triton X-100, 2% glycerol). To lyse cells, 4200 units of Ready-Lyse lysozyme (Lucigen), 30 units of DNase I (New England Biolabs), and 10 μl of concentrated protease inhibitor cocktail (Sigma) (one pellet dissolved in 500 μl of Buffer 1) were added to cell suspensions and incubated at room temperature for 45 min. Cell debris was removed by centrifugation at 10,000 x g for 5 min at 4 °C. 50 μl of cell lysates were set aside and used as an “input” sample. 50 μl aliquots of α-FLAG magnetic bead slurry (Sigma) or α-GFP magnetic bead slurry (MBL biotech) in 1.5 ml microcentrifuge tubes were washed three times with 1 ml of Buffer 2 using a magnetic collection stand. 1 ml of cell lysates was added to washed magnetic beads and subjected to end-over-end rotation at 4 °C for 2 hr. Beads were then washed three times with 0.5 ml of Buffer 2, with 10 min incubations in Buffer 2 at 4 °C between each wash step. Beads were briefly washed a 4th time with 0.5 ml Buffer 2. To elute proteins from α-FLAG beads, 100 μl of elution buffer (150 μg/ml 3X-FLAG peptide, Sigma, in Buffer 2) was added and samples were subjected to end-over-end rotation at 4 °C for 30 min. To elute proteins from α-GFP beads, beads were resuspended in 50 μl PBS, 2x SDS-PAGE sample buffer was added to tubes, and samples were boiled as described above. Eluates and input samples were subjected to western blotting as described above.
Measuring GFP fluorescence in reporter strains
V. cholerae reporter strains harboring PcpiA-GFP were grown to late log in LB medium with or without 0.2% arabinose as indicated. Cells were then washed once in instant ocean medium (7g/L; Aquarium Systems) and transferred to a 96-well plate. Fluorescence was then determined on a Biotek H1M plate reader with monochromater set to 500 nm for excitation and 540 nm for emission.
Fluorescent DNA binding/uptake
A ~7 kb PCR product was fluorescently labeled as described previously11 using the Cy3 LabelIT kit (Mirus Biosciences) as per manufacturer recommendations. For the parent strain, 1 μl (100 ng) of Cy3-DNA was added to 900 μl of LB in a 1.5 ml microcentrifuge tube along with 100 μl of overnight-grown cultures, and cells were grown at 30°C rotating on a roller drum for 70 min. Pili were then labeled as described above.
To visualize DNA-binding in the pilT mutant, cells were grown as described above, but 1 μl (100 ng) of Cy3-DNA was added to cells along with AF488-mal and incubated for 25 min before washing. Cells were imaged using the same microscopy setup described above, using a dsRed filter cube to image Cy3-DNA.
Phylogenetic analysis
TfpB homologues were identified by BLAST using 43 manually selected Gammaproteobacteria genomes34 through Integrated Microbial Genomes and Microbiomes online resources (IMG)35 resulting in 209 protein sequences. The FtsK protein from A. baylyi ADP1 was added manually and used as an outgroup. Through the NGPhylogeny.fr server36, sequences were aligned using the default parameters of MAFFT37, and phylogenetic tree construction was performed on the aligned sequences using the default parameters of FastTree software38,39 set to perform 100 bootstraps40. The resulting tree was visualized using the Interactive Tree of Life (iTOL) visualization software41, from which trees were exported for publication.
Author contributions
CKE and ABD designed and coordinated the overall study. CKE, TND, CAK and ABD performed the experiments. All authors analyzed and interpreted data. CKE and ABD wrote the manuscript with help from ZG and JWS.
Competing interests
The authors declare no competing interests.
Data availability statement
The data that support the findings of this study are available from the corresponding authors upon request.
Supplemental Materials for
Acknowledgements
We would like to thank E. Geisinger for A. baylyi ADP1 ATCC33305 wildtype strain, and we would like to thank K. Hummels and B. Bratton for helpful suggestions on phylogenetic analysis. CKE is a Damon Runyon Fellow supported by the Damon Runyon Cancer Research Foundation (DRG-2385-20). This work was supported in part by the National Science Foundation, through the Center for the Physics of Biological Function (PHY-1734030). This work was supported by grant R35GM128674 from the National Institutes of Health awarded to ABD and the National Institutes of Health Pioneer Award 1DP1AI124669-01 awarded to ZG.