Abstract
Bacterial type IV pili are critical for diverse biological processes including horizontal gene transfer, surface sensing, biofilm formation, adherence, motility, and virulence. These dynamic appendages extend and retract from the cell surface. In type IVa pilus systems, extension occurs through the action of an extension ATPase, PilB, while optimal retraction requires the action of a retraction ATPase, PilT. Many type IVa systems also encode a homolog of PilT called PilU. However, the function of this protein has remained unclear because pilU mutants largely lack obvious phenotypes. Here, we study the type IVa competence pilus of Vibrio cholerae as a model system to define the role of PilU. We show that the ATPase activity of PilU is critical for optimal pilus retraction in PilT ATPase mutants. PilU does not, however, contribute to pilus retraction in ΔpilT strains. Thus, these data suggest that PilU is a bona fide retraction ATPase that supports pilus retraction in a PilT-dependent manner. We also found that a ΔpilU mutant exhibited a reduction in the force of retraction suggesting that PilU is important for generating maximal retraction forces. Additional in vitro and in vivo data show that PilT and PilU act as independent homo-hexamers that likely form a complex to facilitate pilus retraction. Finally, we demonstrate that the role of PilU as a PilT-dependent retraction ATPase is conserved in Acinetobacter baylyi, suggesting that the role of PilU described here may be broadly applicable to diverse type IVa pilus systems.
Author Summary Almost all bacterial species use thin surface appendages called pili to interact with their environments. These structures are critical for the virulence of many pathogens and represent one major way that bacteria share DNA with one another, which contributes to the spread of antibiotic resistance. To carry out their function, pili dynamically extend and retract from the bacterial surface. Here, we show that retraction of pili in some systems is determined by the combined activity of two motor ATPase proteins.
Introduction
Type IV pili are ubiquitous surface appendages in Gram-negative bacteria that promote diverse activities including attachment, virulence, biofilm formation, horizontal gene transfer, and twitching motility [1-5]. These structures can dynamically extend and retract from the cell surface, which is often critical for their function. A detailed mechanistic understanding of pilus dynamic activity, however, remains lacking.
Type IV pili are composed almost exclusively of a single protein called the major pilin, which forms a helical fiber that extends from the cell surface [6]. Extension and retraction of the pilus is hypothesized to occur through the interaction of cytoplasmic hexameric ATPases with the inner membrane platform of the pilus machine, whereby ATP hydrolysis likely facilitates turning of the platform in order to incorporate or remove major pilin subunits from the pilus fiber [7-9]. All type IV pilus systems encode a predicted ATPase (PilB) that facilitates pilus extension [7, 10-12]. The type IVa pilus systems also contain a retraction ATPase (PilT) which depolymerizes the pilus fiber and recycles pilin subunits into the inner membrane. Many type IVa pilus systems also encode another putative ATPase called PilU. PilU is a homolog of the retraction ATPase PilT, however, the function of this protein has remained unclear in most pilus systems because pilU mutants largely lack obvious phenotypes. A notable exception is Pseudomonas aeruginosa, where pilU mutants exhibit reduced twitching motility [13].
The facultative bacterial pathogen Vibrio cholerae uses a type IV competence pilus for DNA uptake during natural transformation. We have recently demonstrated that the type IV competence pili of V. cholerae can be fluorescently labeled by using a technique [14-16] in which an amino acid of the major pilin subunit PilA is replaced with a cysteine (PilAS67C) [17], which allows for subsequent labeling with a fluorescently-labeled thiol-reactive maleimide dye (AF488-mal). This labeling approach does not impede dynamic pilus activity and allows for the direct observation and measurement of pilus extension and retraction by time-lapse epifluorescence microscopy. Using this labeling technique, we have recently demonstrated that the retraction of DNA-bound type IV competence pili is required to initiate the process of DNA uptake during natural transformation [17].
Here, we employ our pilus labeling approach and complementary molecular methods to address the role that PilU plays in type IV competence pilus retraction, and we extend our findings to the type IVa competence pilus of Acinetobacter baylyi. Our results indicate that PilT and PilU form independent homo-hexamers and that the ATPase activity of PilU promotes type IV pilus retraction in a PilT-dependent manner.
Results
PilU promotes type IV competence pilus retraction in a PilT-dependent manner
In order to study the function and role of ATPases in the retraction of type IV competence pili in V. cholerae, we utilize two main methods: (1) our labeling approach to directly measure pilus retraction via epifluorescence time-lapse microscopy and (2) assessing natural transformation, which is dependent on competence pilus retraction [17, 18]. Also, as for other pili [19-21], competence pili can mediate cell-to-cell interactions, which causes hyperpiliated strains to aggregate and pellet to the bottom of culture tubes [22]. Consistent with prior work [17, 18, 23-28], mutant strains that lack the retraction ATPase PilT are poorly transformable, exhibit markedly reduced rates of pilus retraction, aggregate, and are hyperpiliated (≥3 pili per cell) compared to the parent strain (Fig 1A-D). Qualitatively, the number of cells that exhibit dynamic pilus activity is reduced in the ΔpilT strain as observed by time-lapse epifluorescence microscopy, which is consistent with the hyperpiliated phenotype of this mutant. The residual retraction in a ΔpilT strain is not due to PilU because a ΔpilT ΔpilU double mutant strain displays the same phenotype as a ΔpilT strain (Fig 1A-D). By contrast, ΔpilU mutants are largely indistinguishable from the parent, other than a ∼1.3-fold reduction in the rate of pilus retraction (Fig 1B). Thus, these data support a model where PilT plays the dominant role in pilus retraction, which is consistent with prior reports [17, 18, 26].
PilT is a predicted ATPase and its ATPase activity has been shown to be important in the function of other type IV pilus systems [12, 29]. Therefore, we hypothesized that the ATPase activity of PilT would be required for retraction of the competence pilus. We found, however, that strains harboring mutations in PilT that ablate the ATPase activity of this protein—due to a mutation in either an invariantly conserved lysine in the Walker A motif required for ATP binding (pilTK136A), a mutation in a conserved glutamic acid in the Walker B motif required for ATP hydrolysis (pilTE204A), or a double mutant containing both mutations (pilTE204A/K136A)— all transform similarly to the parent strain and retract competence pili at rates that are at most ∼4-fold lower than the parent (Fig 1A-B). This is in stark contrast to a ΔpilT strain where the transformation frequency is reduced ∼4-logs and retraction is >50-fold slower compared to the parent strain (Fig 1A-B). These results were quite surprising and suggested that either (1) the ATPase activity of the retraction machinery is not critical for promoting optimal pilus retraction or (2) that another retraction ATPase may promote retraction in the absence of PilT ATPase activity.
We tested the latter model and specifically assessed whether PilU could promote retraction in PilT ATPase-deficient mutants. Indeed, we found that pilTK136AΔpilU, pilTE204AΔpilU and pilTE204A/K136AΔpilU strains all display reduced transformation, are hyperpiliated, aggregate, and exhibit markedly reduced retraction rates similar to a ΔpilT ΔpilU strain (Fig 1A-D). This is consistent with PilU promoting retraction in the pilTK136A, pilTE204A and pilTE204A/K136A backgrounds. Importantly, the pilTK136AΔpilU, pilTE204AΔpilU and pilTE204A/K136AΔpilU strains could be complemented via ectopic expression of pilT or pilU (S1 Fig). Since loss of pilT alone (i.e. ΔpilT) results in a phenotype comparable to a ΔpilT ΔpilU mutant, this result indicates that PilU is not sufficient to drive pilus retraction in the absence of PilT (Fig 1A-D). Consistent with this, ectopic expression of pilU does not rescue the transformation deficit of a ΔpilT ΔpilU mutant (S1 Fig). Together, these data suggest that PilU supports pilus retraction in a PilT-dependent manner.
Interestingly, the pilTE204A strain is hyperpiliated (Fig 1D). Qualitatively, the frequency of cells that exhibit pilus retraction in this mutant is much lower than the parent and comparable to the ΔpilT ΔpilU strain (as assessed by epifluorescence time-lapse microscopy), which may explain this hyperpiliated phenotype; however, when pilus retraction is observed, the rate of retraction is closer to that of the parent than the ΔpilT ΔpilU mutant (Fig 1B). Considering the canonical functions of the Walker A and B motifs in ATP-binding and hydrolysis, respectively, the Walker B pilTE204A mutant should be able to bind to ATP but not hydrolyze it. Because the ATP-binding deficient pilTK136A Walker A mutant exhibited parental levels of piliation (Fig 1D), we hypothesized that the hyperpiliation observed in pilTE204A may be due to the ATP-binding state of this mutant. Consistent with this, we found that the Walker A and B double mutant (pilTE204A/K136A) exhibits parent levels of piliation in contrast to the pilTE204A single mutant (Fig 1D). These results indicate that the phenotype of the ATP-binding deficient Walker A mutant (pilTK136A) is dominant over the ATP-hydrolysis deficient pilTE204A Walker B mutation. Both of these mutations should result in loss of ATPase activity. Therefore, to avoid phenotypic consequences of variable states of ATP binding associated with the pilTE204A mutant, we utilized the pilTK136A mutant as an ATPase-defective allele of PilT moving forward.
PilU is required for forceful retraction
The data thus far suggest that PilU is sufficient but not necessary to mediate pilus retraction and natural transformation. However, PilU is present in many type IVa pilus systems, so the question remains: Why is PilU so widely conserved if it is not necessary for pilus retraction? Recent work in Pseudomonas aeruginosa indicates that PilU may be required for forceful retraction [30]. Our data thus far indicate that pilus retraction in the pilU mutant is largely indistinguishable from the parent (Fig 1B), however, these measurements are based on free pili that are not retracting under load. We hypothesized that PilU might be required for forceful retraction, such as when pili are attached to a surface. To test this, we assessed pilus retraction using a micropillar assay in which binding of pili to micropillars and subsequent retraction results in measurable bending and displacement of the micropillars. These data can then be used to calculate the rates and force of pilus retraction [31]. In this assay, the ΔpilU mutant exhibited a significant reduction in retraction force (∼2-fold) and speed (∼3-fold) compared to the parent strain, despite the fact that this strain still has functional PilT-dependent retraction (Fig 1B,E-F). This reduction in the force of retraction in the ΔpilU mutant is comparable to the reduction in force of retraction observed in the ΔpilT mutant (Fig 1E). Thus, these data indicate that PilU is necessary for generating maximal retraction forces in the V. cholerae competence pilus.
The ATPase activities of PilT and PilU define the rate of type IV competence pilus retraction
Just like PilT, PilU is a AAA family ATPase with a conserved lysine in its Walker A motif. Because we observed that PilU supports pilus retraction in the pilTK136A mutant (Fig 1B), we hypothesized that the ATPase activity of PilU may be important for supporting pilus retraction. To test the relative roles of each of these ATPases in pilus retraction, we mutated the conserved lysine in the Walker A motifs of PilT and/or PilU. The pilUK134A strain retracts ∼1.5-fold slower than the parent strain, similar to the ΔpilU mutant (Fig 2A). There may be a limited effect of the pilUK134A mutation on pilus retraction because this strain still has a functional PilT ATPase. However, we hypothesized that PilU ATPase activity may be critical for retraction in a background where PilT ATPase activity is also inactivated. Consistent with this, we found that a pilTK136A pilUK134A strain is indistinguishable from a ΔpilT strain for pilus retraction, piliation, aggregation, and transformation (Fig 2A-C). Together, these data suggest that PilU ATPase activity promotes type IV competence pilus retraction.
The data above suggest that the ATPase activities of PilT and PilU are critical for pilus retraction. These ATPases are predicted to be motor proteins that facilitate the depolymerization of pilus fibers. Thus, if we alter the ATPase rates of these motors, we hypothesized that this should alter the rates of pilus retraction. To test this hypothesis, we mutated a leucine in the Walker B domain of PilT and PilU to a cysteine, which was previously shown to reduce the rate of ATP hydrolysis of Neisseria gonorrhoeae PilT ∼2-fold [26]. When these Walker B mutations were tested in isolation they did not alter the rate of pilus retraction (Comparing Parent strain to pilTL201C and pilUL199C) (Fig 2A). This may be due to the fact that these cells still have one fully functional retraction ATPase (i.e. pilTL201C still has a fully functional PilU and pilUL199C still has a fully functional PilT). We hypothesized that these Walker B “slow” ATPase mutations, however, may have an impact on pilus retraction if they represented the sole functional retraction ATPase in the cell. To that end, we generated both a pilTL201C ΔpilU strain and a pilTK136A pilUL199C strain. In both cases, these strains exhibited reduced retraction rates relative to the appropriate parent strain: a ∼1.3-fold reduction for PilTL201C (comparing pilTL201CΔpilU to ΔpilU) and an ∼1.5-fold reduction for PilUL199C (comparing pilTK136ApilUL199C to pilTK136A) (Fig 2A). This provides genetic evidence that the ATPase activities of PilT and PilU are linked to the retraction rate of the type IV competence pilus. Furthermore, it provides additional supporting evidence that PilU promotes pilus retraction through its ATPase activity.
PilT and PilU directly interact with one another
All of the data presented above indicate that PilT and PilU can both support pilus retraction. Therefore, we hypothesized that PilT and PilU may interact to form a complex that supports pilus retraction. To address this, we utilized a Bacterial Adenylate Cyclase Two-Hybrid (BACTH) assay [32, 33]. As expected for predicted hexameric proteins like PilT and PilU, the BACTH analysis showed PilT-PilT and PilU-PilU self-interactions (Fig 3 and S2 Fig). Additionally, the BACTH assay showed that PilT and PilU interact with one another and that the Walker A mutations of either protein (PilTK136A and PilUK134A) did not disrupt this interaction (Fig 3 and S2 Fig). Thus, our BACTH results suggest that PilT and PilU may form a complex to mediate pilus retraction. These results do not, however, indicate whether PilU and PilT form heteromeric complexes or homomeric complexes to facilitate pilus retraction. We test this further below.
PilT and PilU form independent homohexameric rings in vitro and in vivo
PilT in other systems has been shown to form a hexameric complex [34], and by homology we hypothesized that PilU also forms a hexamer. Based on transformation frequency assays, N-terminally tagged 6XHis-PilT and 6XHis-PilU were fully functional (S3A Fig). Therefore, we purified 6XHis-PilT and 6XHis-PilU to characterize these proteins in vitro. Negative stain transmission electron microscopy revealed that both 6XHis-PilT and 6XHis-PilU formed hexameric ring complexes (Fig 4A). These results indicate that these proteins are capable of forming independent homohexameric complexes in vitro. PilT and PilU have both sequence (∼65% similarity) and structural similarity to one another (Fig 4A), thus it is also possible that these proteins can be incorporated into one ring and form heterohexameric complexes that facilitate pilus retraction in vivo.
In order to test if PilT and PilU form heterohexamers or independent homohexamers to facilitate pilus retraction in vivo, we hypothesized that overexpression of ATPase-inactive mutants (i.e. PilTK136A and PilUK134A) in backgrounds that have native PilT and PilU would allow us to distinguish between these two models. For example, if these proteins form heteromeric complexes, then we would predict that overexpression of either ATPase-inactive protein (PilTK136A or PilUK134A) should result in complexes that are mostly composed of ATPase-inactive proteins and result in inhibition of pilus retraction (yielding a hyperpiliated phenotype) and reduce rates of natural transformation (Fig 4B). If these proteins solely form homomeric complexes, however, then we would predict that overexpression of only one ATPase-inactive protein (PilTK136A or PilUK134A) should only inhibit the activity of one motor complex (PilT or PilU, respectively) and should not have a dramatic effect on pilus retraction or natural transformation because the other retraction motor would still be intact (Fig 4B).
In order for us to successfully test this question, we needed to: (1) be able to ectopically overexpress ATPase deficient alleles of PilT or PilU, (2) demonstrate that ATPase deficient alleles can interact with their ATPase sufficient counterparts and form proper hexamers, and (3) show that ectopic expression of ATPase deficient alleles of PilT or PilU can inhibit the activity of their respective natively expressed ATPase. We test each of these in turn below. First, we confirmed that we could ectopically overexpress PilT and PilU. Using functional 3xFLAG-tagged versions of PilT and PilU (S3A-B Fig), we show that our chromosomal Ptac regulated constructs resulted in dramatic overexpression of these proteins (S3C Fig). Importantly, overexpression of WT PilT and PilU did not affect pilus activity or natural transformation (S1 and S3B,D Figs). Next, we purified 6XHis-PilTK136A and 6XHis-PilUK134A and demonstrated that they formed ring structures in vitro (S3E Fig), indicating that these proteins should form complexes with ATPase sufficient alleles. This is also supported by our BACTH data (Fig 3). Finally, we wanted to confirm that overexpressing the ATPase-inactive PilTK136A or PilUK134A could effectively impede the function of the natively expressed ATPase-active PilT or PilU. To test this, we genetically isolated either PilT or PilU as the sole functional retraction ATPase (i.e. PilT activity is isolated in a ΔpilU background, while PilU activity is isolated in a pilTK136A background) and then overexpressed the cognate ATPase-inactive allele (Ptac-pilTK136A or Ptac-pilUK134A, respectively). We assessed piliation, aggregation, and natural transformation as indicators of optimal pilus retraction. We found that, as expected, overexpression of ATPase-inactive alleles was sufficient to disrupt the activity of the natively expressed ATPase (Fig 4C-D - compare ΔpilU to ΔpilU Ptac- pilTK136A and pilTK136A to pilTK136A Ptac-pilUK134A). Also, as expected, overexpression of both ATPase-deficient alleles together (Ptac-pilTK136A and Ptac-pilUK134A) in a background that has natively expressed PilT and PilU, results in hyperpiliation, aggregation and reduced natural transformation (Fig 4C-D), which further demonstrates that overexpression of these ATPase-deficient proteins inhibits the activity of their natively expressed ATPase–sufficient versions.
The experiments above indicated that it would be feasible to test whether PilT and PilU formed homomeric or heteromeric complexes in vivo as described in our model (Fig 4B). When we overexpressed only one ATPase-deficient protein (Ptac-pilTK136A or Ptac-pilUK134A) in a background that has natively expressed PilT and PilU, we found that piliation and transformation were indistinguishable from the parent (Fig. 4C-D). Thus, these data are consistent with PilT and PilU forming independent homohexamers to facilitate pilus retraction.
Using our purified PilT and PilU, we also attempted to directly measure the ATPase activity of these proteins. Unfortunately, the ATPase activity of these purified proteins was indistinguishable from contaminating ATPases and/or below our detection limit (∼1 nmol pi/min/mg protein). Poor ATPase activity in purified retraction ATPases is not unique to our system [12, 29, 35]. This may be due to the fact that the ATPase activities of these proteins are stimulated in vivo via interaction with the proteins that compose the pilus machine, as previously suggested [12]. Regardless, our previous in vivo data (Fig 1A-D) are consistent with the pilTK136A mutant being ATPase deficient.
The presence of PilT is not required for PilU to interact with the pilus platform PilC
Our data thus far indicate that PilU requires the presence of PilT (whether it is a functional ATPase or not) to support optimal pilus retraction. The extension and retraction ATPases are hypothesized to interact with the platform protein PilC to mechanically facilitate polymerization and depolymerization of the pilus fiber [7, 36, 37]. Because PilU is dependent on the presence of PilT to mediate optimal retraction, we hypothesized that PilT can interact with PilC, but that PilU cannot. To test this, we utilized the BACTH assay to test the interactions of PilT and PilU with PilC. This assay showed that PilT and PilU could both interact with PilC, and that the Walker A mutations of these ATPases (PilTK136A and PilUK134A) did not affect these interactions (Fig 3). The interaction between PilC and PilT supports the canonical model that retraction ATPases interact with PilC to facilitate depolymerization of the pilus fiber. Interestingly, PilC and PilU interact independently of PilT. Thus, the simple model of PilU requiring PilT for interaction with the platform PilC appears to be incorrect.
The function of PilU as a PilT-dependent retraction ATPase is conserved
As mentioned earlier, many type IVa pilus systems encode PilU in addition to the canonical retraction ATPase PilT. Therefore, we sought to assess whether PilU could facilitate PilT-dependent retraction in other type IV pilus systems. To test this, we studied the type IVa competence pili of Acinetobacter baylyi.
The competence pili of A. baylyi likely facilitate DNA uptake in a retraction-dependent manner, similar to the competence pili of V. cholerae. Thus, we employed natural transformation assays to dissect the role that PilT and PilU play in A. baylyi competence pilus retraction. These assays showed that the PilT and PilU of A. baylyi behave largely like the PilT and PilU of V. cholerae. The pilU mutant of A. baylyi displays a significant (∼13-fold) reduction in natural transformation (Fig 5), unlike what is observed in V. cholerae (Fig 1A). This may be because DNA uptake in A. baylyi requires more forceful retraction or because PilU plays a more important role in pilus retraction in this species. Most importantly, however, we found that PilU facilitates natural transformation (and therefore pilus retraction) in backgrounds where the ATPase activity of PilT is inactivated, but not in backgrounds where pilT is absent (Fig 5 – compare pilTK136A to pilTK136A ΔpilU and ΔpilT). Also, a strain where the ATPase activity of both PilT and PilU were inactivated (pilTK136A pilUK137A) resulted in the loss of transformation equivalent to a retraction deficient background (e.g. ΔpilTU) or a strain lacking competence pili (ΔcomP). Importantly, strains lacking pilT (i.e. ΔpilT and ΔpilTU) could only be complemented via ectopic expression of pilT but not pilU, while strains that contained an ATPase defective allele of pilT (i.e. pilTK136A ΔpilU or pilTK136A pilUK137A) could be complemented by ectopic expression of pilT or pilU (S4 Fig) These data are consistent with what was observed for V. cholerae competence pili (Fig 1A and S1 Fig) and suggest that PilU ATPase activity facilitates pilus retraction in a PilT-dependent manner in diverse type IVa pilus systems.
Discussion
This study sheds light on the function of PilU as a retraction ATPase of type IVa competence pili in V. cholerae. Specifically, our data indicate that PilU can mediate pilus retraction in a PilT-dependent manner likely through its ATPase activity. Our results also show that PilT and PilU form independent homohexamers that likely interact to form a complex that supports pilus retraction. Furthermore, we demonstrate that the role of PilU in supporting pilus retraction is conserved in at least one other type IVa pilus system.
Our results indicate that PilU can only facilitate pilus retraction in the presence of PilT, regardless of whether that PilT is a functional ATPase. These results are also supported by a recent preprint from the Blokesch lab [22]. We initially tested the hypothesis that this was due to a requirement for PilT to bridge PilU with the platform protein PilC. Our BACTH results, however, indicated that PilU may directly interact with PilC in the absence of PilT. While they may interact, it is possible that PilU does not bind and engage the PilC platform in a conformation that would facilitate retraction. Alternatively, it is possible that PilU does bind PilC in a conformation that can facilitate retraction but additional interactions with PilT are required to stimulate PilU ATPase activity. Another possibility is that PilT interacts with other components of the pilus machine that are required for stimulating retraction, and that PilU lacks these interactions. Future efforts focused on the interaction network of the retraction ATPases with the pilus machine should shed light on this question.
Previous reports have shown that PilU is necessary for twitching motility in P. aeruginosa [13], an activity that likely requires forceful retraction [30]. Consistent with this, our results indicate that PilU is necessary for maximal retraction force for the competence pilus in V. cholerae. Interestingly, because the ΔpilU mutant transforms similarly to the parent strain (Fig 1B and 2A), the forceful retraction mediated by PilU does not seem to be necessary for the function of the V. cholerae competence pilus in DNA uptake under the conditions tested. However, forceful PilU-dependent retraction may be necessary for DNA uptake when tDNA is absorbed onto surfaces and/or in complex environments like biofilms.
Beyond altering the force of retraction, many mutants in PilT and PilU also reduced the speed of pilus retraction (i.e. ΔpilU = ∼1.3-fold, pilTK136A = ∼4.3-fold, pilTK136A pilUL199C = ∼6.5-fold, pilUK134A = ∼1.5-fold, pilTL201C ΔpilU = ∼1.7-fold). This revealed that altering the rates of these motor ATPases results in a corresponding reduction in the rate of pilus retraction. Additionally, despite this reduction in the speed of pilus retraction, these mutants still transformed at levels that were comparable to the parent strain. Thus, these results suggest that the absolute speed of competence pilus retraction is not critical for DNA uptake during natural transformation. Further experiments will focus on defining the parameters that are essential for DNA uptake.
Mutants that completely inactivated the retraction machinery (e.g. ΔpilT, ΔpilT ΔpilU, pilTK136A ΔpilU, pilTK136A pilUK134A) resulted in a hyperpiliated phenotype, lower transformation frequency and a marked reduction in the frequency of pilus retraction (as observed qualitatively by epifluorescence time-lapse microscopy). These strains, however, did display rare pilus retraction, which occurred at rates that were markedly slower than the parent (>40-fold), which is independent of PilT and PilU. Recent work in Neisseria indicates that PilTU-independent retraction is not unique to the V. cholerae competence pilus [38]. The mechanism underlying PilTU-independent retraction is still unclear [5] and will be the focus of future work.
Materials and Methods
Bacterial strains and culture conditions
Vibrio cholerae and A. baylyi strains were routinely grown in LB Miller broth and on LB Miller agar supplemented with erythromycin (10 μg/mL), kanamycin (50 µg/mL), and/or carbenicillin (50 µg/mL) when appropriate.
Construction of mutant strains
All V. cholerae strains used throughout this study are derivatives of the El Tor isolate E7946. All A. baylyi strains used were derivatives of strain ADP1 [39]. All mutant strains were constructed via MuGENT and natural transformation using transforming DNA products made by splicing-by-overlap extension (SOE) PCR exactly as previously described [40-43]. For a detailed list of all mutant strains used throughout this study see Table S1. For a detailed list of all primers used to construct mutant strains see Table S2.
Natural transformation assays
In order to induce competence in V. cholerae strains, the master competence regulator TfoX was overexpressed using an IPTG-inducible or constitutive Ptac promoter and the cells were genetically locked in a state of high cell density via deletion of luxO [41, 44-48]. Chitin-independent transformation assays were performed exactly as previously described [42]. Briefly, strains were grown overnight rolling at 30 °C with 100 µM IPTG then ∼108 colony forming units (CFU) were subcultured into 3mL of LB + 100 µM IPTG + 20 mM MgCl2 + 10 mM CaCl2and grown to late log. Next, ∼108 CFU of this culture was diluted into instant ocean medium (7 g/L; Aquarium Systems) supplemented with 100 µM IPTG and 500ng of transforming DNA was added to each reaction and incubated statically at 30 °C overnight. The tDNA targeted VC1807, a frame-shifted transposase, for deletion as previously described [40]. Negative control reactions where no tDNA was added were also performed for each strain. After incubation with tDNA, reactions were outgrown by adding 1mL of LB to each reaction and shaking (250 rpm) at 37 °C for ∼2 hours. Reactions were then plated for quantitative culture onto medium selecting for transformants (LB + 10 μg/mL Erythromycin) or onto plain LB for total viable counts. The transformation frequency is defined as the number of transformants divided by the total viable counts. For reactions where no transformants were obtained, a limit of detection was calculated and plotted.
In order to sensitize natural transformation assays to differences in the efficiency of DNA uptake, reactions were incubated with transforming DNA for shorter periods of time. These assays were prepared exactly as described above, except that 10 units of DNase I (NEB) was added to reactions 7 minutes after incubation with 500ng of transforming DNA. Reactions were then incubated statically at 30°C overnight, outgrown, and plated exactly as described above.
For A. baylyi, transformations were performed exactly as previously described [49]. Briefly, strains were grown overnight in LB medium. Then, ∼108 cells were diluted into fresh LB medium, and tDNA was added (∼100 ng). Reactions were incubated at 30°C with agitation for 5 h and then plated for quantitative culture as described above to determine the transformation frequency.
Pilin labelling, imaging and quantification
In order to label pili for observation with epifluorescence microscopy, strains were grown to the late-log phase exactly as described above for natural transformation assays. Then, ∼108 CFU were spun down at 18,000 x g for 1 min and resuspended in instant ocean medium. Cells were then incubated with 25 μg/mL AF488-mal for 15 minutes statically at room temperature in the dark. Cells were washed twice using 100μL of instant ocean medium and resuspended to a final concentration of ∼107 CFU. Next, 2μL of these labeled cells were placed under an 0.2% gelzan pad (made in instant ocean medium) on a coverslip and imaged on an inverted Nikon Ti-2 microscope with a Plan Apo ×60 objective, a green fluorescent protein filter cube, a Hamamatsu ORCAFlash 4.0 camera and Nikon NIS Elements imaging software. In order to measure retraction events, labelled cells were imaged by time-lapse microscopy in which a phase-contrast (to image cell bodies) and fluorescent (to image labeled pili) image were taken every second for 1-2 minutes or every 10 seconds for 10-15 minutes. Rates were calculated manually using measurement tools in the NIS Elements analysis software. Only pili that were longer than 0.3 μm and completed retracting within the time-lapse window were analyzed.
In order to calculate the number of pili per cell, static images of cell bodies and labeled pili were captured using phase-contrast and epifluorescence microscopy, respectively. Images from 3 independent biological replicates were sectioned into areas containing ∼100-200 cells and the total number of cells and the number of pili displayed on each cell was manually determined. Representative images of the piliation state of strains were gathered and the lookup tables for each phase or fluorescent image were adjusted to the same range.
To assess aggregation and pelleting of cultures, strains were grown to late log under the conditions described above. Cultures were then allowed to sit statically at room temperature for 30-60 mins. Images of cultures were then taken against a white background.
Protein expression and purification
His tagged (6xHis) versions of PilT, PilTK136A, PilU and PilUK134A were cloned into a pHis expression vector and verified by DNA sequencing (Eurofins). Vectors were transformed into E. coli BL21 DE3 for expression and purification.
For purification of His-PilU and His-PilUK134A, cells were grown in a 1L flask at 37°C with aeration to an OD600 = 0.6. The culture was then induced by adding IPTG (1 mM) and grown to a final OD600 = 4.0 at 30°C with aeration. Cells were harvested by centrifugation at 8,000 x g for 15 min at room temperature and then resuspended in Buffer A [100 mM Tris pH 8.5, 300 mM KCl, 10% glycerol, 20 mM Imidazole] and stored at −80°C. Pellets were thawed on ice and then lysozyme (1 mg/mL) and DNase I (2 mg/L) were added before lysing via sonication with a probe tip sonicator. The soluble fraction was clarified by centrifugation at 12,000 x g for 1 hour at 4°C. The supernatant was loaded onto a nickel-charged HisTrap HP column (1mL; GE) using an ÄKTA FPLC at room temperature. The column was then washed with Buffer A and the protein was eluted with a gradient of Buffer B [100 mM Tris pH 8.5, 300 mM KCl, 10% glycerol, 500 mM imidazole] and fractions were collected on ice. Elution fractions were separated on 15% SDS-PAGE gels and stained with Coomassie Brilliant Blue R-250 (BioRad). Peak fractions were pooled and dialyzed (Fisher, MWCO 12 kDa) into 500 mL Buffer C [100 mM Tris pH 8.5, 300 mM KCl, 10% glycerol] overnight, on a stir plate at 4°C. Then a second round of dialysis was performed using 500 mL Buffer C for 4-6 hours, on a stir plate at 4°C. Following dialysis, the protein concentration was determined using a Bradford Assay and aliquots were stored at −80C.
For purification of His-PilT and His-PilTK136A, cells were grown in a 1L flask at 37°C with aeration to an OD600 = 0.6. The culture was then induced by adding IPTG (1 mM) and grown overnight at 22°C with aeration. Cells were harvested by centrifugation at 8,000 x g for 15min at room temperature and then resuspended in Buffer A [25 mM Tris pH 8.5, 300 mM KCl, 10% glycerol, 1 mM EDTA, 1 mM MgCl2, 5 mM β-mercaptoethanol, 20 mM Imidazole] and stored at −80°C. Pellets were thawed on ice and lysozyme (1 mg/mL), DNase I (2 mg/L) and PMSF (1 mM) were added before lysing via sonication with a probe tip sonicator. The soluble fraction was clarified by centrifugation at 12,000 x g for 1 hour at 4°C. The supernatant was loaded onto a nickel-charged HisTrap HP column (1 mL; GE) using an ÄKTA FPLC at room temperature. The column was then washed with Buffer A and the protein was eluted with a gradient of Buffer B [25 mM Tris pH 8.5, 300 mM KCl, 10% glycerol, 1 mM EDTA, 1 mM MgCl2, 5 mM β-mercaptoethanol, 500 mM Imidazole] and fractions were collected on ice. Elution fractions were separated on 15% SDS-PAGE gels and stained with Coomassie Brilliant Blue R-250 (BioRad). Peak fractions were pooled and dialyzed (Fisher, MWCO 12 kDa) into 500 mL Buffer C [25 mM Tris pH 8.5, 50 mM KCl, 10% glycerol, 1 mM EDTA, 1 mM MgCl2, 0.5mM TCEP] overnight, on a stir plate at 4°C. The next day, the eluate was concentrated with an Amicon conical filter (MWCO 30 kDa) at 4,000 x g for 15 min at 4°C and resuspended to the original volume in Buffer C twice. The protein concentration was determined using a Bradford Assay and then aliquots of protein were snap-frozen with liquid nitrogen and stored at −80C.
Bacterial Adenylate Cyclase Two-hybrid (BACTH) Assays
The Bacterial Adenylate Cyclase Two-Hybrid (BACTH) system was employed to study protein interactions. Gene inserts were amplified by PCR and cloned into pUT18C (CarbR) and pKT25 (KanR) vectors to generate N-terminal fusions to the T18 and T25 fragments of adenylate cyclase, respectively. Miniprepped vectors (Qiagen plasmid miniprep kit) were then cotransformed into E. coli BTH101 and outgrown with 500 µL LB for 1 hour at 37°C with shaking. Transformations were isolation streak plated on LB plates containing kanamycin 50 µg/mL and carbenicillin 100 µg/mL to select for transformants that had received both plasmids. Strains were then grown statically at 30°C from frozen stocks overnight in LB supplemented with kanamycin 50 µg/mL and carbenicillin 100 µg/mL and 3.5µL of each strain was then spotted onto LB plates containing kanamycin 50 µg/mL, carbenicillin 100µg/mL, 0.5 mM IPTG and 40 µg/mL X-gal and incubated at 30°C for 48 hours.
BACTH Miller Assays
For BACTH Miller assays strains were prepared by growing the BTH101 strains with the indicated pUT18C and pKT25 vectors statically overnight at 30°C in LB supplemented with kanamycin 50 µg/mL and carbenicillin 100 µg/mL. Then 5 µL of this overnight culture was subcultured into 3mL of LB supplemented with kanamycin 50 µg/mL, carbenicillin 100µg/mL and 0.5 mM IPTG and grown rolling at 30°C overnight. Cultures were then pelleted and resuspended in 1 mL of Z buffer [60 mM Na2HPO4-7H2O, 40 mM NaH2PO4-H2O, 10 mM KCl, 2 mM MgSO4-7H2O] to an OD600 = 0.5. 50µL of 1% SDS was added to each reaction and vortexed. 100µL of chloroform was added to each reaction, vortexed again and then centrifuged for 1 min at 18,000 x g. The aqueous fraction was then mixed with ONPG (2.5 mg/mL) in a 96-well plate and the OD420 and OD550 were measured every 5 minutes for 10 hours. Miller units were then calculated at the time (in mins) when the A420 reached 0.3 or at 10 hours using the formula: [1000*(A420-(1.75*A550))/ (mins*0.2 mL*A600)].
Western Blotting
Strains were grown to mid-log phase in LB with 100 µM IPTG if applicable and then resuspended to an OD600 = 100 in TBS containing 1 mM PMSF. Cells were then lysed using a FastPrep-24 (MP Bio). Lysates were centrifuged at 16,000 x g for 10 min at 4°C, and the supernatant was then mixed 1:1 with 2X SDS sample buffer [220 mM Tris pH 6.8, 25% glycerol, 1.8% SDS, 0.02% Bromophenol Blue, 5% β-mercaptoethanol] and then 2uL of each sample was separated on a 15% SDS PAGE gel. Proteins were transferred to a PVDF membrane and incubated with one of the following primary antibodies: α-FLAG polyclonal rabbit (Sigma) or α-RpoA monoclonal mouse (Biolegend). Then, blots were washed and incubated with a α-rabbit or α-mouse secondary antibodies conjugated to IRdye 800CW. Blots were then imaged using an Odyssey classic LI-COR system.
Negative-stain electron microscopy
Negative stain specimens were prepared by applying 4 µL of protein solution (0.1 mg/mL + 1 mM AMP-PnP) onto a glow-discharged 300-mesh copper grid coated with continuous carbon film (EMS). After 30 s the protein solution was blotted with a piece of filter paper. The grid was washed with 4 µL of milli-Q water and stained with 4 µL of stain solution (1% (w/v) uranyl acetate, 0.5% (w/v) trehalose). The excess stain solution was removed by filter paper and the grid was left to air dry. Due to the availability of the instrument, we used two different transmission electron microscopes for data collection. For PilU, the data were collected on a 300-kV JEM-3200FS TEM equipped with FEG source and 8k x 8k Direct Electron DE-64 camera. A total of 32 micrograph images were taken at a nominal magnification of 60,000x, which is equivalent to 1.56 Å per pixel. For PilT, the data were collected on a 120-kV JEM-1400plus TEM equipped with LaB6 source and a Gatan Oneview camera. A total of 59 images were taken at a nominal magnification of 60,000x and the pixel size is 1.94 Å. To avoid potential beam induced damage, minimum dose system was employed during data collection. Note that PilU data were collected on a direct electron detector; therefore, each image contains 50 frames and the frame alignment was performed using unblur program with exposure filter applied. Particle picking, defocus estimation and phase flipped particles were all done using EMAN2 software package [50]. These particles were then imported into RELION (v2.1) for 2D classification, Initial model building, 3D refinement, and 3D classification [51]. From 2D classification results, we observed clearly 6-fold symmetry; thus, the symmetry operation was applied to subsequent 3D refinement and 3D classification. The final 3D reconstructions were computed from 45204 particles to 13.1 Å for PilU and 17472 particles to 14.7 Å for PilT. The resolution estimation was calculated using gold-standard method at Fourier shell correlation of 0.143 as implemented in RELION. However, due to the nature of the negative stain, the effective resolution could be equal to or worse than 15 Å. The 3D reconstructions were rendered using UCSF Chimera where the handedness of PilU was chosen to be consistent with PilT [52].
Micropillar retraction assay
To ensure that all measurements recorded were due to type IV competence pili, all the experiments were performed with V. cholerae strains lacking all external appendages other than type IV competence pili as well as exopolysaccharide production (i.e. mutants lacking flagella, MSHA pili, TCP pili and Vibrio polysaccharide (VPS) production). Strains were grown overnight in LB either directly from frozen stocks or from a single colony from an LB agar plate. Of the overnight liquid culture, 50 μl was subcultured into 3 ml LB + 100 μM IPTG + 20 mM MgCl2 + 10 mM CaCl2 and allowed to grow at 30°C for 4h. Then, 100 μl of this culture was centrifuged for 5 min at 20,000 x g and the bacteria were resuspended in instant ocean medium. At different times, 23 μl of the resuspension were added to micropillars in an observation chamber as previously reported [15]. Briefly, silica molds were inverted on activated coverslips with polyacrylamide gels in between. The result is an array of flexible micropillars in a hexagonal array of 3 μm × 3 μm.Once the bacteria were in contact with the micropillars, 10-Hz movies of the top of the pillars were recorded. The motion of the tips of the pillars was tracked using a cross-correlation algorithm in ImageJ. The amplitude and speeds of the pillars’ motions were then analyzed using MATLAB. Finally, we calibrated the pillars’ stiffness constant using optical tweezers as previously described [31]. The pillars used in this study have a stiffness constant of 17±4pNμm−1.
Statistics
Statistical differences were assessed by two-tailed Student’s t-tests using GraphPad Prism software.
Supporting Information for
Acknowledgements
We would like to thank CK Ellison for helpful discussions. We would like to acknowledge JD Newman and CE Dann III for their advice during protein purification. We thank JC van Kessel for providing BACTH strains and plasmids, and Lakshay Khosla and Vijay Deopersaud for technical assistance with micropillar experiments. This work was supported by grants R35GM128674 and AI118863 from the National Institutes of Health to ABD, and by grant AI116566 from the National Institutes of Health to NB.