Surface stiffness stimulates pilus production through transcriptional and post-transcriptional mechanisms

The increase in the number of pili upon association with solid surfaces suggested that growth conditions might stimulate the production of pilus components. To investigate the effects of surface association on the transcriptional regulation of pilus genes, we used real-time PCR to examine the mRNA levels of pilA (a gene encoding the major pilin subunit (Strom & Lory, 1986)) and pilT (a gene encoding the ATPase responsible for retraction of the pilus (Whitchurch et al., 1990)) in liquid and surface grown cells. Examination of pilin subunit levels provides an indirect measure of pilus production while characterization of pilT levels complements the protein localization studies described below. When compared to liquid grown cells in stationary phase, pilA transcription increased ~15 fold when cells were grown on the surface of agar plates (Fig. 2A), while pilT levels were unaffected by changes in growth condition (Fig. 2B). To control for differences due to growth phase or cell density, we examined pilA and pilT mRNA levels in liquid grown cells that were in either log or stationary phase and found no significant differences (Fig. 2). As an additional comparison, we found that the gene encoding the flagellum subunit (fliC) showed no change in expression under the same conditions (data not shown). To gain a further understanding of the environmental regulation of pilus factors, we monitored pilus protein levels during the different growth conditions using Western blot analysis. In accordance with the transcriptional regulation of pilA, PilA protein levels were also regulated in response to growth conditions. Surface growth resulted in a ~15-fold increase in PilA levels (Fig. 2A). Although pilT mRNA levels were the same in liquid grown cells and surface grown cells, PilT protein levels were ~3 fold greater upon surface association (Fig. 2B), suggesting a post-transcriptional mechanism for PilT regulation.

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Figure 2

Surface association regulates both pilus gene expression and pilus protein levels

(AB) Relative levels of pilA (A) and pilT (B) gene (white bars) and PilA (A) and PilT (B) protein (black bars) expression in cells grown to logarithmic (log) or stationary phase in LB broth and in cells taken from the leading edge of motility on the surface of plates containing increasing concentrations of agar. Error bars represent the standard deviation (n=3) and asterisks denote significant differences from stationary phase cells (P < 0.01 for mRNA levels and P < 0.05 for protein levels).

To further explore the role of surface association in regulating pilus expression, we monitored pilus mRNA and protein levels in cells grown on plates with increasing concentrations of agar. Increased levels of both pilA mRNA and PilA protein directly correlated with increased agar concentration in the plates (Fig. 2A). Similarly, PilT protein levels increased with increased levels of surface stiffness (Fig. 2B), demonstrating that substrate stiffness may be one factor that impacts pilus expression. These results show that the effects of surface association are not limited to regulation of pilin expression and represent both transcriptional and post-transcriptional modes of regulation.

The polar localization of GFP-PilT is regulated by growth on surfaces

Since pili are predominantly found at the cell poles, another potential mechanism for regulating pilus production is to modulate the subcellular localization of pilus proteins. To monitor the localization of one type IV pilus protein, we created an arabinose-inducible N-terminal GFP fusion to PilT. While PilT is not required for pilus assembly, PilT does function at the cytoplasmic base of the pilus such that PilT localization was predicted to serve as a reporter for pilus localization. The GFP-PilT fusion protein was confirmed to be functional by complementation of the twitching motility defect of a pilT mutant and had no adverse effects on wild-type twitching motility (Fig. S2). As expected, the GFP-PilT fusion protein primarily localized to the cell pole (Fig. 3). The majority of cells had GFP-PilT located at one or both poles (50.7% and 14.9%, respectively) while a smaller proportion of the population (29.8%) exhibited diffuse fluorescence (Fig. 3). In addition, a small number of cells (4.6%) had GFP-PilT spots at both polar and nonpolar sites in the cell (Fig. 3). The overall distribution of GFP-PilT (unipolar, bipolar, nonpolar, or diffuse) correlated well with our direct observations of pili using TEM (Fig. 1G), indicating that GFP-PilT may reflect the location of the assembled pilus structure.

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Figure 3

GFP-PilT is more polarly localized during growth on a solid surface

(A) GFP-PilT localization in logarithmic phase cells grown in suspension is unipolar (P), nonpolar (NP), bipolar (BP), or diffuse (Dif). The arrow indicates a nonpolar GFP-PilT focus. (B) Colocalization of GFP-PilT (top) and PilQ-mCherry (bottom) in cells grown on agar plates. (C–D) GFP-PilT localization in stationary phase cells taken from the leading edge of motility on the surface of agar plates (C) or grown in suspension (D). Scale bars represent 2 μm. (E) The localization patterns of GFP-PilT in cells collected from the surface of agar plates. Error bars represent the standard deviation (n=3 replicates of 100 cells each). (F) GFP-PilT fluorescence intensity across the cell length for representative liquid grown (white squares) and surface grown (black squares) cells. The cells used to create these plots are indicated by arrowheads in A and C.

As an additional assay for the accuracy of GFP-PilT as a reporter for pilus location, we found that GFP-PilT colocalized with a mCherry fusion to PilQ (the multimeric outer membrane secretin required for pilus assembly) in greater than 80% of cells (Fig. 3B, n = 4 replicates of ~250 cells each). Colocalization was defined as an identical localization pattern for each fusion (i.e. if GFP-PilT was unipolar, then PilQ-mCherry was considered colocalized only if found exclusively at the same pole). We also confirmed the functionality of PilQ-mCherry by showing that the fusion complemented the twitching motility defect of a pilQ mutant and had no adverse effects on wild-type twitching motility (Fig. S2). Previous work using a YFP-PilT fusion showed that PilT localizes in a bipolar fashion in more than 90% of cells (Chiang et al., 2005). It remains unclear as to the reason for the difference in our results, but this discrepancy could result from differences in growth conditions or simply from strain differences (PAO1 versus PAK). Together, the agreement of pilus and GFP-PilT localization distributions and the colocalization of PilT and PilQ indicate that GFP-PilT localizes with other pilus components and can therefore serve as a reporter for pilus protein localization.

To determine if surface association regulates PilT localization, we examined localization of GFP-PilT in liquid and surface grown cells. As with the pili themselves, GFP-PilT was more abundant (brighter) at the poles when cells were grown on a surface compared to fluorescence from liquid grown cells (Fig. 3A, C). Quantitative fluorescence intensity plots are shown in Figure 3F for representative liquid and surface grown cells. The difference between liquid and surface grown cells was not the result of fusion protein induction times or growth phase since cells grown to stationary phase in liquid medium with induction for 24 h showed no increase in polar fluorescence of GFP-PilT (Fig. 3D). To further confirm that the differences in PilT localization are not merely a secondary consequence of increased GFP-PilT abundance during surface growth, we found that induced gfp-pilT expression and GFP-PilT protein levels in liquid grown cells were similar to those from surface grown cells (Fig. S3).

To characterize the dynamics of GFP-PilT localization in response to contact with a surface, we induced expression of GFP-PilT in liquid-grown cells and then monitored GFP-PilT localization immediately upon transfer to solid LB agarose pads. GFP-PilT fluorescence became more defined (brighter and more punctate) at the cell poles over time (Fig. 4A). By quantitating the fluorescence intensity of GFP-PilT in cells transitioning from liquid to surface growth, we found that the total cellular fluorescence did not increase over time, indicating that the increased polar fluorescence came at the expense of the GFP-PilT distributed throughout the rest of the cell (Fig. 4B). To control for cell density differences between liquid and surface grown conditions, we examined the dynamics of GFP-PilT localization at varying cell densities and found that GFP-PilT accumulated at the cell pole at the same rate, regardless of cell density on the agarose pad (data not shown). The representative images shown in Figure 4 are from the low cell density experiments, where individual cells were isolated from neighboring cells at the start of the experiment. Similarly, cells grown at high and low densities in liquid culture had no detectable difference in GFP-PilT localization (Fig. 3). These results suggest that in addition to regulating pilus component mRNA and protein levels, surface association also regulates the polar localization of GFP-PilT.

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Figure 4

GFP-PilT accumulates at the cell pole during solid surface growth

(A) Time-lapse images of a representative P. aeruginosa cell carrying the GFP-PilT fusion taken every 10 min during growth on agarose pads. For each time-point, GFP fluorescence (top) and phase contrast (bottom) images are shown. Scale bar represent 2 μm. (B) GFP-PilT fluorescence intensity across the cell length for the topmost cell in (A). Closed squares indicate intensities at 10 min and open squares indicate intensities at 60 min of growth on the agarose pad.

P. aeruginosa MreB forms a cell cycle regulated helix that is sensitive to treatment with A22

Having demonstrated that pilus protein localization is actively controlled, we next sought to investigate the molecular basis of this regulation, focusing on the MreB actin homolog as a candidate localization determinant. In several bacterial species, MreB is essential for maintenance of cell shape, chromosome segregation, and polar protein localization (Dye et al., 2005, Divakaruni et al., 2007, Gitai et al., 2004, Gitai et al., 2005, Kruse et al., 2005). Previous work has demonstrated that P. aeruginosa mreB is essential and is targeted by an antibacterial indole compound CBR-4830 (Robertson et al., 2007). However, the localization and functional properties of MreB have yet to be explored in P. aeruginosa. To determine if MreB is involved in polar pilus protein localization in P. aeruginosa, we first characterized P. aeruginosa MreB itself. We examined the localization of P. aeruginosa MreB by constructing an arabinose-inducible N-terminal mCherry fusion to MreB (mCherry-MreB) and observed that mCherry-MreB forms a helical pattern in P. aeruginosa using deconvolution microscopy (Fig. 5A-B). We also monitored the dynamics of mCherry-MreB localization during the cell division cycle by timelapse-imaging individual cells mounted on agarose pads containing rich medium. P. aeruginosa mCherry-MreB undergoes spatial rearrangement during the cell cycle, transitioning from a spiraled structure stretching the length of the cell (Fig. 5A-B) to a ring structure at the cell division site (Fig. 5C). These dynamics closely resemble those observed in Caulobacter crescentus (Figge et al., 2004, Gitai et al., 2004), but have not been described for many other bacteria. Unlike GFP-PilT, mCherry-MreB localization is unaffected by liquid versus surface growth conditions (data not shown).

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Figure 5

P. aeruginosa MreB forms a helix that is sensitive to treatment with A22

(A–B) Deconvolution images of the mCherry-MreB helix. (C) Medial condensation of mCherry-MreB localization during growth on LB agarose pads (fluorescence (left) and phase contrast (right) images). (D–E) P. aeruginosa cells incubated for 3 h in the absence (D) or presence (E) of A22 (100 μg/ml). (F–G) Fluorescence images of mCherry-MreB localization 30 seconds after exposure to A22 (100 μg/ml) (F) and 2 minutes after removal of A22 (G). Scale bars represent 2 μm.

In C. crescentus, treatment with the chemical inhibitor A22 (S-(3,4-dichlorobenzyl)isothiourea) leads to gradual changes in cell morphology due to perturbation of MreB localization and function (Gitai et al., 2005). Recent work has also determined that A22 causes similar changes in P. aeruginosa cell shape in an efflux-compromised strain (Robertson et al., 2007). We thus sought to use A22 to characterize the function of P. aeruginosa MreB. We found that after several hours in the presence of growth-inhibitory levels of A22 (100 μg/ml; A22¹⁰⁰), wild-type P. aeruginosa transitions from normal rod-shaped cells to lemon-shaped cells (Fig. 5D–E). We monitored P. aeruginosa mCherry-MreB localization patterns in the presence of A22 and observed that, in all cells examined, A22¹⁰⁰ rapidly disrupted mCherry-MreB localization in a reversible manner: MreB became completely diffuse within 30 seconds after exposure to A22¹⁰⁰ (Fig. 5F) and relocalization occurred 2–3 minutes after A22 was removed (Fig. 5G).

The cellular target of A22 was identified as mreB in C. crescentus by selecting for spontaneous resistance to A22 (Gitai et al., 2005). We performed a similar selection for A22-resistant P. aeruginosa and identified 12 strains with missense mutations in mreB (Fig. S4), indicating that A22 also targets MreB in P. aeruginosa. These findings are consistent with a previous study that linked P. aeruginosa mreB point mutants to increased resistance to the unrelated indole compound CBR-4830 and A22 in an efflux-compromised strain (Robertson et al., 2007). Six additional A22-resistant strains were isolated in our selection, but we found no sequence changes in the mreBCD operon or its predicted promoter region. Our results suggest that A22 may have additional targets in P. aeruginosa, but we cannot eliminate the possibility that resistance also may be due to indirect effects on MreB activity.

To confirm that the increased resistance to A22 is due to the point mutations in mreB and not the consequence of a secondary mutation, we moved two representative mutations (I171S and G189C) into a clean genetic background. The original mutant strains containing the MreB I171S and G189C mutations had wild-type growth rates and normal cell shape (data not shown), indicating that the majority of MreB function was maintained. Strains carrying plasmid-born copies of the mutated versions of mreB displayed the same increased resistance to A22 as the original spontaneous mutants (data not shown), demonstrating that these mutations are responsible for the phenotype observed and are dominant over the wild-type mreB. These strains can thus also serve as controls strains to dissect the MreB-dependent effects of A22 treatment. Taken together, these data indicate that as in other species, P. aeruginosa MreB localizes in a helix that is in a position to potentially regulate polar protein localization and that the small molecule inhibitor A22 is a useful reagent for rapidly perturbing MreB function.

Disruption of MreB affects the polar localization of PilT

Having validated MreB as a candidate localization determinant and A22 as a tool for studying MreB function in wild-type P. aeruginosa, we sought to use A22 to begin to characterize the role of MreB in the polar localization of the P. aeruginosa type IV pilus. Initially, we examined the effects of A22 treatment on polar GFP-PilT. Within one minute after treatment with A22¹⁰⁰ (a condition that causes mCherry-MreB to become diffuse in all cells), liquid grown cells displayed a rapid change in GFP-PilT localization patterns. As described above, without A22 treatment, the majority of cells contained concentrated polar foci (Fig. 3, ​,6A).6A). Once exposed to A22, most cells maintained polar localization of GFP-PilT, but a number of those cells also acquired nonpolar GFP-PilT foci (Fig. 6B). Analysis of these images revealed that a significant proportion of the A22-treated cell population had increased numbers of nonpolar GFP-PilT at the expense of the cells with unipolar GFP-PilT (Fig. 6C, P < 0.005). To confirm that GFP-PilT sensitivity to A22 was a result of MreB disruption and not a secondary effect of A22 treatment, we observed GFP-PilT in the A22-resistant mreB-mutant P. aeruginosa strains I171S and G189C. GFP-PilT localization in the A22-resistant mreB mutants was unaffected by A22 treatment (Fig. S4), indicating that MreB contributes to the polar localization of PilT.

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Figure 6

GFP-PilT localizes to the bacterial cell pole in an MreB-dependent manner

(A–B) Maintenance of GFP-PilT localization in liquid grown cells in the absence (A) or presence (B) of A22 (100 μg/ml) treatment after GFP-PilT induction. Arrowheads denote nonpolar GFP-PilT. (C) The distribution of GFP-PilT within a population of cells (diffuse, unipolar, bipolar, or nonpolar) in the absence (white bars) or presence (black bars) of A22 (100 μg/ml). Error bars indicate standard deviation (n=3 replicates of 100 cells each). Asterisks denote significant differences due to treatment with A22 (P < 0.005). (D–E) Maintenance of GFP-PilT localization in surface grown cells in the absence (D) or presence (E) of A22 (100 μg/ml) treatment after GFP-PilT induction. Scale bars represent 2 μm.

MreB control of PilT localization is regulated by surface association

Based on the data described above, we hypothesize that MreB is important for directing the localization of PilT to the cell pole. MreB could be involved in the initial placement of PilT at the pole or may be responsible for maintaining that localization once PilT has reached the pole by other means. By varying the order of GFP-PilT induction and treatment with A22, we were able to address the role of MreB in initiation and maintenance of PilT localization. In addition, since both growth conditions and MreB affect PilT localization, we examined the possibility that association with a surface might influence PilT by regulating MreB’s involvement in PilT localization.

If MreB were required for initial localization of PilT to the cell pole, then treatment with A22 prior to or concurrently with induction of GFP-PilT expression would prevent GFP-PilT from reaching the pole. When GFP-PilT expression was induced at the same time as treatment with A22, either in cells grown in liquid medium containing A22¹⁰ or in cells grown on agar plates containing A22¹⁰, GFP-PilT fluorescence was diffuse in 100% of cells examined (n > 100, Fig. 7D), indicating that functional MreB is indeed needed for initiation of polar PilT in both growth conditions. Meanwhile, if MreB were involved in maintenance of polar PilT localization, then treatment with A22 after induction of GFP-PilT expression would detach localized PilT from the cell pole. In liquid grown cells, A22 treatment after GFP-PilT induction caused an increase in nonpolar GFP-PilT localization (Fig. 6B). In contrast, A22 treatment of surface grown cells after GFP-PilT induction had no effect on GFP-PilT localization (Fig. 6D–E), suggesting that MreB is important for maintenance of PilT localization in liquid grown cells but not in cells grown on a surface. These results indicate that A22 can perturb both the initiation and maintenance of polar PilT localization when cells are grown in suspension, but only affects the initiation of polar PilT localization when cells are grown on a surface (Fig. 8).

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Figure 7

A22 disruption of MreB alters pilus production prior to affecting cell shape

(A) The distribution of pili on the cell surface (no pili, unipolar pili, bipolar pili, or nonpolar pili) was monitored by TEM for P. aeruginosa cells taken from the leading edge of motility on the surface of LB plates with no treatment (white bars), plates with 10 μg/ml A22 (black bars), or treated with 100μg/ml mecillinam for 2 h (gray bars). Error bars represent the standard deviation (n=3 replicates of 100 cells each). Asterisk indicates significant differences from untreated cells (P < 0.005). (B–C) TEM images of P. aeruginosa taken from the surface of plates containing 10 μg/ml A22 (B) or treated with 100 μg/ml mecillinam for 2 h (C). Closed arrowheads denote flagella, open arrowheads indicate representative pili, and the arrow indicates a nonpolar pilus. (D–E) The effects of A22 and mecillinam treatment on initiation of GFP-PilT localization (top) and cell shape (phase contrast, bottom) were performed by treating cells with A22 at the time of GFP-PilT induction. Scale bars represent 500 nm (B–C) and 2 μm (D–E).

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Figure 8

Model for how surface association regulates the role of MreB in PilT localization

In the absence of A22 (far left), MreB contributes, either directly or indirectly, to polar localization of GFP-PilT (green circles). In surface grown cells (bottom), GFP-PilT becomes more concentrated at the pole, potentially due to the presence of a surface-growth-specific anchoring factor (yellow rectangle). When cells are treated with A22 after GFP-PilT induction (middle), the maintenance of polar PilT localization is disrupted in liquid grown cells (top), while the putative anchoring factor retains PilT localization in surface grown cells (bottom). When GFP-PilT induction occurs concurrently with A22 treatment (far right), initiation of polar PilT localization is prevented and GFP-PilT remains diffuse (green shaded area) under both growth conditions.

P. aeruginosa respond to surface association by regulating both the production and localization of pilus components

P. aeruginosa utilizes type IV pili for surface motility and attachment (Mattick, 2002, Kohler et al., 2000), indicating that the bacterium likely has little need for pili during growth in liquids. Consistent with this hypothesis, we and others see an increase in the number of cells with pili during growth on solid surfaces compared to liquid growth. To our knowledge, this study is the first detailed quantification of multiple aspects of pilus production in response to surface contact. The increased assembly of pili stems at least in part from increased expression of pilus factors when cells are grown in contact with a surface. Thus, P. aeruginosa stimulates pilus production when it encounters environmental conditions in which pili can provide a selective advantage. Interestingly, we found that pilus production is controlled at both the transcriptional and post-transcriptional level. The transcription factors described above may be responsible for the effects we observed on pilA levels, but the mechanism underlying post-transcriptional regulation of PilT is unclear. Post-transcriptional regulation of PilT would provide a rapid regulation step that could potentially alter pilus function even if the pili are completely assembled.

Eukaryotic cells can respond to contact with a surface based on the mechanical properties of the surface substrate (Discher et al., 2005). Depending on the cell type, variations in surface stiffness can lead to changes in cell adhesion, cytoskeletal organization, and cell differentiation. Recently, this type of mechanosensing has also been described for bacteria. Independent of membrane composition or cell shape, increasing substrate stiffness leads to increased adherence of both Staphylococcus epidermidis and Escherichia coli (Lichter et al., 2008). Although a full characterization of this phenomenon is outside the scope of this work, our findings suggest that P. aeruginosa can also sense and respond to its mechanical environment by increasing production of pilus components in response to increased surface stiffness (as measured by increasing concentrations of agar in solid media). Thus, just as bacteria use chemical cues from their environment to modulate developmental programs, they may also incorporate information about their mechanical environment in order to regulate physiological responses such as motility, adhesion, biofilm formation, and virulence factor production. In the future, it will be interesting to determine how bacteria sense and respond to these and other mechanical environments.

Although transcriptional and post-transcriptional modes of regulation are well established as bacterial adaptations to the environment, it has only recently been appreciated that changes in protein localization can also occur in response to extracellular signals. Here, we show that PilT accumulates at the pole in response to surface association. The regulation of protein localization in response to contact with a surface has also been seen with the P. aeruginosa chemotaxis protein WspR (Guvener & Harwood, 2007). However, PilT accumulation at the pole is visible within one cell cycle while WspR localization changes take at least 3 cell division cycles, suggesting that these responses are distinct. In Myxococcus xanthus, PilT oscillates between poles when cells reverse direction to provide the retractive force necessary for pilus-mediated motility (Bulyha et al., 2009). The structural properties of pili in M. xanthus and P. aeruginosa are highly similar (Mattick, 2002), suggesting that pilus protein localization may be similarly regulated. However, our PilQ localization results suggest that pilus regulation in these organisms is not fully conserved. In M. xanthus, structural components of the pilus such as the outer membrane secretin PilQ remain static at both poles, regardless of the direction of motion (Bulyha et al., 2009, Nudleman et al., 2006). The oscillation of M. xanthus PilT and PilB (the ATPase required for pilus extension) between poles is thought to instruct the direction of cell movement (Bulyha et al., 2009). The colocalization of P. aeruginosa PilT and PilQ at unipolar, bipolar, nonpolar, or diffuse sites indicate that a different mechanism may regulate pilus activity in Pseudomonas. Clearly, unique strategies have evolved in different bacteria to regulate protein localization, and continued characterization of multiple systems will be required to fully understand the mechanisms of protein localization regulation.

Transmission electron microscopy

For liquid grown cells, overnight bacterial cultures were subcultured 1:100 into fresh LB and grown for 3 h (logarithmic phase) or 24 h (stationary phase) at 37°C with gentle shaking. Cells were then pelleted (3,000 rpm for 1 min) and resuspended gently in PBS. For surface grown cells, overnight cultures were inoculated to 1% agar LB plates and incubated at 37°C for 24 h. Surface cells were collected from the leading edge of surface motility and gently resuspended in PBS. For TEM, bacteria were placed on 200 mesh thin-film carbon with nitrocellulose, glow-discharged grids and stained with 1% uranyl acetate. Bacterial pili were visualized at 80kV on a Zeiss 912AB Transmission Electron Microscope equipped with an Omega Energy Filter (Confocal and Electron Microscopy Core Facility Laboratory, Princeton University) at a magnification of 5000X. Micrographs were captured using a digital camera from Advanced Microscopy Techniques. For the distribution of pilus locations within cell populations, 100 cells from three independent replicates were counted for each treatment. Pili could be clearly distinguished from the single, polar flagellum because they are smaller in diameter, present at higher numbers, and lack the sinusoidal curve associated with the flagellum. Samples were blinded to assure objective scoring and nonpolar pili were counted only when distinct from sheared pili. As an additional control, we counted the number of sheared pili (those not associated with a bacterial cell) for each treatment. To examine the effect of A22 treatment on pilus distribution, 10 μg/ml A22 was added to 1% agar LB plates and cells were collected as described above. To monitor the effect of mecillinam on pilus distribution, cells were collected as described above and incubated with mecillinam (100 μg/ml) for 2 h.

Construction of fluorescent fusions

The Gateway^® Technology (Invitrogen, Carlsbad, CA) was utilized to construct N-terminal fluorescent fusions to mreB and pilT and a C-terminal fusion to pilQ using a site-specific recombinase derived from bacteriophage λ. Briefly, mreB, pilT, and pilQ genes were PCR-amplified using Gateway-compatible primers MreBGWUp, MreBGWDn, PilTGWUp, PilTGWDn, PilQGWUp, and PilQGWDn, respectively, and cloned into pDONR223 with the BP recombination reaction. The resulting entry clones were confirmed by sequencing (Genewiz, South Plainfield, NJ). mreB and pilT were then transferred into the destination vectors pSW(mCherry-Gateway) for mreB and pSW(GFP-Gateway) and pJN(GFP-Gateway) for pilT using the LR recombination reaction. pilQ was transferred into the destination vector pJN(Gateway-mCherry) using the same reaction. pSW(mCherry-Gateway) and pSW(GFP- Gateway) were constructed by PCR-amplifying mCherry or GFP in conjunction with the Gateway^® cassette using the primers RFPNtermSpeIFor and GWNtermSpeIRev and cloning the products into pSW196 (Baynham et al., 2006). Primer RFPNtermSpeIFor contains a ribosome-binding site for efficient expression in Pseudomonas. pSW196 contains an arabinose-inducible promoter upstream of the multiple cloning site and will integrate at the attP site on the P. aeruginosa chromosome. pJN(GFP-Gateway) was constructed by PCR-amplifying GFP-Gateway as described above and cloning the product into pJN105(Newman & Fuqua, 1999). pJN(Gateway-mCherry) was constructed by PCR-amplifying mCherry in conjunction with the Gateway^® cassette using the primers GWrfpFORSpe and GWrfpREVSpe and cloning the products into pJN105 (Newman & Fuqua, 1999). The expression vectors pSW(mCherry-mreB), pSW(gfp-pilT), pJN(gfp-pilT), and pJN(pilQ-mCherry) were conjugated into P. aeruginosa PAO1. Integration of pSW(mCherry-mreB) and pSW(gfp-pilT) at the attP site was confirmed using primers SerUp and SerDown, and maintenance of pJN(gfp-pilT) and pJN(pilQ-mCherry) was confirmed using primers AraCAmp and GWrfpREVSpe. Primer sequences are listed in Table S1.

Microscopy of fluorescent fusions

P. aeruginosa carrying the mCherry-MreB, GFP-PilT, or PilQ-mCherry protein fusions were grown overnight in LB broth, subcultured 1:100 into fresh LB broth containing 0.02% arabinose, and grown for 3 h (logarithmic phase) or 24 h (stationary phase) at 37°C. To collect single images of protein localization, bacterial cells were placed on pads made from 1% agarose in water and visualized using a 100X 1.4NA objective on a Nikon 90i microscope equipped with a Rolera XR camera and NIS Elements software. For timelapse imaging of growing cells, bacteria were placed on pads made from 1% agarose in LB broth and coverslips were sealed with valap (1:1:1 mixture of Vaseline, lanolin, and paraffin). Phase contrast and fluorescence images were collected every 10 min for 3 h. To monitor localization in cells grown on solid surfaces, bacterial cultures were inoculated to 1% agar LB plates containing 0.02% arabinose, grown for 24 h at 37°C, scraped from leading edge of motility on the plate, resuspended in PBS, and visualized on pads as described above. For studies examining colocalization of GFP-PilT and PilQ-mCherry, ~250 cells were counted from 4 independent samples. Colocalization was defined as an identical localization pattern for each fusion (i.e. if GFP-PilT was unipolar, then PilQ-mCherry was considered colocalized only if found exclusively at the same pole).

Images were normalized by subtracting background fluorescence (defined as fluorescence attributed to the agarose pad (not cell-associated)) from each image being compared using the NIS Elements software. The subtracted value was defined by the image with the highest background fluorescence. To analyze the increased accumulation of polar GFP-PilT in Figures 3 and ​and4,4, background fluorescence was subtracted as described above and the fluorescence intensity along the length of the cell was measured using the Plot Profile function in ImageJ software, which gives fluorescence values in arbitrary units.

Functionality of GFP-PilT and PilQ-mCherry fusions

To confirm that the GFP-PilT and PilQ-mCherry fusions were functional, pJN105 (empty vector), pJN(gfp-pilT), and pJN(pilQ-mCherry) were transformed into wild-type PAO1 and the transposon mutants pilT::Tn and pilQ::Tn (Jacobs et al., 2003). To examine pilus-dependent twitching motility in these strains, overnight cultures were stabbed into 1% agar LB plates with 0.02% arabinose and inoculated at 37°C for 48 h. Twitching motility was observed by removing the agar and staining cells attached to the Petri dish with 1% crystal violet.

A22 treatment

To measure the effects of A22 treatment on P. aeruginosa cell morphology, overnight bacterial cultures were subcultured 1:100 into fresh LB broth in the absence or presence of growth inhibitory levels of A22 (100 μg/ml) and incubated at 37°C for 3 h. Phase contrast images of the resulting cells were taken as described above. To examine the effect of A22 on the initiation of protein localization in liquid-grown cells, growth permissive levels of A22 (10 μg/ml) were added to cultures at the time of arabinose induction. To monitor initiation of GFP-PilT localization in cells grown on solid surfaces in the presence of A22, bacterial cultures were inoculated to 1% agar LB plates containing 0.02% arabinose and A22 (10 μg/ml), grown for 24 h at 37°C, scraped from the leading edge of motility on the plate, resuspended in PBS, and visualized on pads as described above. The effect of A22 on the maintenance of protein localization was studied by preparing bacterial cultures as described above in the absence of A22 and adding A22 (100 μg/ml) to cooled molten agarose before pouring the pads. To monitor the rate at which P. aeruginosa mCherry-MreB was delocalized by A22, phase contrast and fluorescence images were collected every 30 sec for 5 min after initial placement on the A22-containing agarose pad. To determine whether this delocalization was reversible, the mCherry-MreB fusion was induced as described above, and cells were incubated in A22 (100 μg/ml) for 10 min to delocalize mCherry-MreB. The A22-treated bacteria were then placed on 1% agarose pads without A22, and images were collected every 30 sec for 5 min.

Isolation and characterization of A22-resistant strains

To determine the target of A22, overnight cultures of P. aeruginosa were plated on LB plates containing 100 μg/ml A22 and incubated at 37°C for 2–3 days. Colonies that arose after that time were considered A22-resistant. Chromosomal DNA was isolated from individual colonies and the mreBCD region was PCR-amplified and sequenced.

To confirm the causality of the point mutations in mreB, mreB was PCR-amplified from wild-type P. aeruginosa and from two representative A22-resistant mutants (I171S and G189C) using primers mreBFOREco and mreBREVXba and cloned into pJN105. pJN105, pJN(mreB), pJN(mreB*I171S), and pJN(mreB*G189C) were conjugated into wild-type P. aeruginosa PAO1 and dilutions were plated on LB plates containing 100 μg/ml A22 for CFU counts.

Real-time PCR

Total RNA from wild-type P. aeruginosa PAO1 was isolated from cells grown in LB broth to an OD₆₀₀ of 0.8 (logarithmic phase) or OD₆₀₀ of 5.0 (stationary phase) and from cells collected at the leading edge of motility from the surface of LB agar plates after 24 h with 3 replicates per growth condition. RNA was isolated using the RNeasy Mini kit (Qiagen, Valencia, CA), treated with DNase I (Ambion, Austin, TX), and used to make cDNA with random decamer primers and MMLV Reverse Transcriptase (RT) (Ambion, Austin, TX). Reactions for real-time PCR were performed in duplicate in 20 ml reactions with SYBR^® Green PCR Master mix (Applied Biosystems, Foster City, CA), cDNA template, and appropriate primers. Reactions were carried out with a two-step cycling protocol on a 7900HT Fast Real-Time PCR System (Applied Biosystems) and results were analyzed with SDS RQ Manager software (Applied Biosystems). Transcript levels of pilT and pilA were measured with primers PilTForRT, PilTRevRT, PilAForRT, and PilARevRT, respectively. As a negative control, water was used in the place of template cDNA. Additionally, DNase-free RNA without RT treatment from each sample was used as template to detect any potential DNA contamination. As expected, no product was detected in any negative control samples. Cycle threshold results for each sample were normalized according to rpoD levels (amplified with RpoDForRT and RpoDRevRT) and then converted to relative values factoring in a two-fold change in product amount per cycle. Primer sequences are listed in Table S1.

Western blot analysis

Whole cell lysates from wild-type P. aeruginosa PAO1 were isolated from cells grown in LB broth to an OD₆₀₀ of 0.8 (logarithmic phase) or OD₆₀₀ of 5.0 (stationary phase) and from cells collected at the leading edge of motility from the surface of LB agar plates with 0.25%, 0.5%, 0.75%, or 1.0% agar for 24 h with 3 replicates per growth condition. Cell samples were washed in PBS, pelleted, solubilized in 2X SDS/PAGE loading buffer to an equivalent of OD₆₀₀ = 10.0, boiled for 10 minutes, and serially diluted two-fold (PilT) or 10-fold (PilA) in 2X SDS/PAGE loading buffer for immunoblot quantification. Samples were separated on 11.25% (for PilT) or 15% (for PilA) SDS/PAGE gels and transferred to nitrocellulose membranes. For immunoblot analysis, membranes were blocked with 5% milk (w/v) in TBS-T, then washed and immunodetected in TBS-T. Primary anti-PilT antiserum (kindly provided by L. Burrows, McMaster University) was used at 1:1,000, primary anti-PilA antiserum (kindly provided by J. Engel, UCSF) was used at 1:100,000, and secondary anti-rabbit-HRP IgG conjugate (GE Healthcare, NA934V) was used at 1:8,000. ECL Western Blotting Reagent (GE Healthcare, RPN2106) and film were used for signal detection; ImageJ software was used to quantify signal intensity (Abramoff et al., 2004). To quantify PilT levels in P. aeruginosa carrying the GFP-PilT fusion, cultures were grown in LB broth or on 1% agar LB plates in the absence or presence of 0.02% arabinose and samples were collected and processed as described above.

Motility assays

To assess the gross effects of A22 treatment on pilus-mediated motility, overnight cultures of wild-type P. aeruginosa were stabbed into 1% agar LB plates containing no A22 or 10 μg/ml A22. Surface motility was defined as movement from the point of inoculation on the surface of the agar plate. Subsurface twitching motility was observed by removing the agar and staining cells attached to the Petri dish with 1% crystal violet. To microscopically observe the motility of individual cells, cells were scraped from the leading edge of surface motility from plates with or without A22 (10 μg/ml), resuspended in PBS, and spotted to 1% agarose pads containing A22. Timelapse images were taken over the course of 1 h and cells that exhibited discontinuous jerking motion typical of pilus-driven motility were classified as motile. Movement due to flagellum-based swimming motility was not considered as pilus-driven motility for these experiments. Standard error was calculated from 3 replicates of at least 250 cells each.

This work was supported in part by a NIH New Innovator Award (Number 1DP2OD004389-01), a Beckman Young Investigator Award, and a Grand Challenges Seed Program from Princeton University awarded to Z.G., and Award Number F32AI074271 from the National Institute of Allergy and Infectious Diseases given to K.N.C. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infectious Diseases or the National Institutes of Health.

We would like to thank M. Bisher for technical assistance with transmission electron microscopy, C. Cowles for technical assistance with Westerns, L. Burrows for the PilT antibody, J. Engel for the PilA antibody, M. Pomianek for synthesis of A22, B. Boles for plasmids and C. Manoil for P. aeruginosa PAO1.