Probing flagellar promoter occupancy in wild-type and mutant Caulobacter crescentus by chromatin immunoprecipitation

Authors

  • Nicole J. Davis,

    1. Department of Molecular Biology and Microbiology, Case Western Reserve University, Cleveland, OH, USA
    2. Department of Infectious Diseases, School of Medicine, University of California San Francisco, San Francisco, CA, USA
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  • Patrick H. Viollier

    1. Department of Molecular Biology and Microbiology, Case Western Reserve University, Cleveland, OH, USA
    2. Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland
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  • Editor: Olga Ozoline

Correspondence: Nicole J. Davis, Department of Infectious Diseases, School of Medicine, University of California San Francisco, 513 Parnassus Avenue, Room HSE418, San Francisco, CA 94143, USA. Tel.: +1 925 321 9988; fax: +1 415 476 9364; e-mail: nicole.davis@ucsf.edu

Abstract

In the asymmetric predivisional cell of Caulobacter crescentus, TipF and TipN mark the cellular pole for future flagellar development. TipF is essential for motility and contains a cyclic-di-GMP phosphodiesterase-like (EAL) domain that is necessary for proper function. TipN is localized to the flagellar pole before TipF and is essential for the proper placement of the flagellum in C. crescentus. Using β-galactosidase promoter-probe assays and quantitative chromatin immunoprecipitation, we investigated the influence of the C. crescentus flagellar assembly regulator TipF on flagellar gene transcription. We compared the transcriptional activity of class II-fliF-lacZ, class III-flgE-lacZ, and class IV-fljL-lacZ fusions in a ΔtipF mutant with that of other flagellar mutants and the wild-type strain. We subsequently verified the in vivo occupancy of the fliF, flgE, and fljL flagellar promoters by the flagellar regulators CtrA, FlbD, and FliX in addition to RNA polymerase. We deduce that TipF contributes to proper expression of flagellar genes in C. crescentus by acting both within and outside of the canonical flagellar gene expression hierarchy.

Introduction

The alphaproteobacterium Caulobacter crescentus divides asymmetrically every cell cycle to form two dissimilar progeny: a nonmotile stalked cell and a motile, polarly flagellated swarmer cell. Assembly of the single, polar flagellum in the predivisional cell occurs with the aid of the birth scar markers TipN (Huitema et al., 2006; Lam et al., 2006) and TipF (Huitema et al., 2006). The latter contains an EAL domain homologous to the catalytic domain in bis-(3′-5′)-cyclic dimeric GMP (cyclic-di-GMP) phosphodiesterases (Bobrov et al., 2005; Schmidt et al., 2005; Tamayo et al., 2005; Huitema et al., 2006). Cells that lack TipF are nonmotile and impaired in the translation and secretion of the FljK flagellin, a class IV flagellar gene product and major component of the flagellar filament (Huitema et al., 2006). With the goal of further characterizing the flagellar assembly defect of TipF cells, we studied flagellar gene expression in ΔtipF cells, comparing it with that of wild-type (WT) cells and other flagellar assembly mutants.

Flagellar biogenesis in C. crescentus requires over 50 genes organized into a regulatory hierarchy of four expression classes (Fig. 1) to link the assembly of flagellar gene expression to cell cycle progression (Minnich & Newton, 1987; Ohta et al., 1991; Ramakrishnan et al., 1994). The master cell cycle transcriptional regulator CtrA, encoded at the class I transcriptional level, accumulates and initiates the transcription of class II flagellar genes in S-phase (Quon et al., 1996). As class II gene products are expressed and assembled into the early (MS-ring basal body) substructure, their transcription ceases as a result of the repressive action of the σ54-dependent transcriptional regulator FlbD and its interacting partner FliX (Mohr et al., 1998; Anderson & Gober, 2000; Gober & England, 2000) at the time of cell division. Concurrent with the repression of class II genes, FlbD/FliX and σ54-containing RNA polymerase (Eσ54) activate the transcription of class III/IV flagellar genes that form the hook (FlgE), P-, and L-rings and the flagellar filament (Anderson & Gober, 2000). An additional layer of regulation operates on the expression of class IV (flagellin) genes, whose message stability is modulated by the negative regulator FlbT, an RNA-binding protein (Mangan et al., 1999), and FlaF, a protein with unknown biochemical activity (Llewellyn et al., 2005). Collectively, all levels of regulation ensure the accrual of gene products at the time when they are needed for the ordered expression and assembly into the growing flagellum structure.

Figure 1.

 Illustration of flagellar biogenesis in Caulobacter crescentus divided into four classes. The upper diagram depicts flagellar assembly as it proceeds from the inner cell membrane (IM) to the outer cell membrane (OM). The hook is assembled on the outside of the cell, followed by the filament. The lower section shows a schematic of representative genes (fliF, flgE, and fljL) from each class with the regulatory elements: FliX, FlbD, CtrA, σ70-, and σ54-containing RNAP, integration host factor (IHF), and flagellar transcriptional regulator (ftr) present at the promoters.

The transcription of flagellar genes is controlled by FlbD, a σ54-dependent protein that binds the flagellar transcriptional regulation (ftr) sequence motif located in the −10 to −35 region of the class II promoters, and the −100 region of the class III promoters, to repress and activate transcription, respectively (Benson et al., 1994; Mullin et al., 1994; Wingrove & Gober, 1994; Dutton et al., 2005). Direct evidence of FlbD binding to flagellar promoters in vivo has not been shown. FlbD activity is modulated by the trans-acting factor FliX that links class II flagellar assembly to class III/IV flagellar gene transcription in two ways (Wingrove & Gober, 1994; Muir et al., 2001; Muir & Gober, 2004). First, FliX stimulates the activation of class III genes by FlbD during the assembly of the basal body. Second, when flagellar assembly is blocked, FliX prevents the activation of the class III gene pathway by FlbD (Muir & Gober, 2002, 2004). Genetic and biochemical studies provide evidence for FliX binding directly to FlbD (Muir & Gober, 2002, 2004) to prevent binding to ftr (Dutton et al., 2005); yet, whether FliX associates with FlbD-dependent promoters in vivo remains to be determined.

TipF, a predicted 50-kDa protein with two N-terminal transmembrane domains, a coiled-coil region, and a C-terminal EAL domain, is required for flagellum biogenesis (Huitema et al., 2006). TipN, a membrane-embedded landmark protein, dictates the proper localization of TipF and the flagellar structure (Huitema et al., 2006; Lam et al., 2006). Little is known about how TipF and TipN affect flagellar gene expression. Here, we use β-galactosidase promoter probe assays and quantitative chromatin immunoprecipitation (qChIP) analyses to explore how a ΔtipF mutation affects the activity of flagellar promoters when compared with WT, a flagellar assembly (ΔfliG) mutant, positioning (ΔtipN), and regulatory (fliX∷Tn5 and flbD∷Tn5) mutants. These experiments reveal, for the first time, the direct quantification of the occupancy of flagellar promoters by their cognate transcriptional regulators in vivo.

Materials and methods

Strains and growth conditions

Caulobacter crescentus NA1000, a synchronizable derivative of the CB15 wild-type strain (Evinger & Agabian, 1977), and derivatives were grown at 30 °C in peptone yeast extract (PYE) [2 g peptone, 1 g yeast extract, 0.2 g MgSO4, and 1 mL CaCl2 (0.5 M) per liter] (Poindexter, 1964; Johnson & Ely, 1977).

β-Galactosidase assays

β-Galactosidase activity (Miller, 1972) was measured at 30 °C with log-phase cultures grown in PYE–tetracycline (0.5 μg mL−1). Assays were performed in triplicate, with a minimum of two independent cultures for each promoter construct.

Antibody production

For the generation of anti-FlbD antibodies, FlbD was overexpressed in Escherichia coli Rosetta (DE3)/pLysS using pET28a (Novagen) as an N-terminal His6-tagged variant and purified using Ni-NTA agarose (Qiagen). Purified proteins were cut out from a 12.5% sodium dodecyl sulfate (SDS) polyacrylamide gel and used to immunize rabbits (Josman LLC).

qChIP

Cells (20 mL) were grown to the mid-log phase and cross-linked in 10 mM sodium phosphate (pH 7.6) and 1% formaldehyde for 10 min at room temperature and on ice for 30 min thereafter. Cells were then washed three times in phosphate-buffered saline (pH 7.4), resuspended in 500 μL of TES buffer [10 mM Tris-HCl (pH 7.5), 1 mM EDTA, and 100 mM NaCl] containing 20 μL of 250 U μL−1 of Ready-Lyse (Epicentre), and incubated at room temperature for 15 min. Following incubation, 500 μL of ChIP buffer [1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris-HCl (pH 8.1), and 167 mM NaCl] containing one protease inhibitor tablet (Roche) was added to the lysates and incubated at 37 °C for 10 min. The lysates were then sonicated (Sonicator 3000, Misonix Inc., Farmingdale, NY) on ice using 10 bursts of 20 s at output level 4.5 to shear DNA fragments to an average length of 300–700 basepairs and cleared by centrifugation at 10 956 g for 2 min at 4 °C. The protein content of the lysates was normalized, diluted to 1 mL in ChIP buffer with 0.01% SDS, and precleared with 100 μL of Protein-A agarose (Roche), 100 μg bovine serum albumin (BSA), and 300 μg herring sperm DNA for 1 h at 4 °C. The supernatant (10%) was removed and used as total chromatin input DNA. Antisera: anti-CtrA (2 μL) (Quon et al., 1996); anti-RNA polymerase (RNAP) against RpoC subunit (2 μL) (Neoclone); anti-FlbD (1 μL); or anti-FliX (0.5 μL) (Mohr et al., 1998) was added to the remaining lysate, respectively, and incubated overnight at 4 °C. After overnight incubation, the supernatant was incubated with Protein-A agarose beads (100 μL), previously incubated with BSA and herring sperm, for 2 h at 4 °C. The beads were then washed once with: low-salt buffer [0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl (pH 8.1), 150 mM NaCl]; high-salt buffer [0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl (pH 8.1), 500 mM NaCl], and LiCl buffer [0.25 M LiCl, 1% Triton X-100, 1% sodium deoxycholate, 1 mM EDTA, 10 mM Tris-HCl (pH 8.1)], and twice with TE buffer [10 mM Tris-HCl (pH 8.1) and 1 mM EDTA]. The protein–DNA complexes were eluted from the beads by adding 500 μL of elution buffer (1% SDS, 0.1 M NaHCO3) with 300 mM NaCl to the beads, and incubating them overnight at 65 °C to reverse cross-linking. The samples were then incubated with 2 μg of Proteinase K for 2 h at 45 °C in 40 mM EDTA and 40 mM Tris-HCl (pH 6.5). DNA was extracted using phenol : chloroform : isoamyl alcohol (25 : 24 : 1) and precipitated with 100% ethanol, using glycogen (20 μg) as a carrier, and resuspended in 50 μL of water.

Quantitative real-time PCR

Real-time PCR was performed using a MyIQ single-color real-time PCR detection system (Bio-Rad, Hercules, CA) using 5% of each ChIP sample, 12.5 μL of SYBR green PCR master mix (Bio-Rad or Quanta), 10 pmol of primers, and 5.5 μL of water per reaction. A standard curve generated from the cycle threshold (Ct) value of the serially diluted chromatin input was used to calculate the percentage input value of each sample. Average values are from triplicate measurements performed per culture. The final data were generated from three independent cultures. DNA regions analyzed by real-time PCR were from nucleotides −190 to −8 relative to the start codon of fliF; from −176 to +23 relative to the start codon of flgE; and from −214 to −21 relative to the start codon of fljL.

Results and discussion

To examine the transcription of flagellar genes in WT and the ΔtipF mutant, we first measured β-galactosidase activity of lacZ transcriptional reporters fused to class II-fliF (MS-ring), class III-flgE (hook), and class IV-fljL (flagellin) promoters. The ΔtipF mutant strain was also compared with a hook basal-body mutant ΔfliG (lacking a component of the flagellar switch bound to the MS-ring), the flagellar placement mutant ΔtipN, and the transcriptional regulatory mutants, fliX∷Tn5 and flbD∷Tn5. Relative to WT, the class II-fliF-lacZ fusion was upregulated in ΔtipF (174 ± 5%) and ΔfliG (318 ± 4%) (Fig. 2). Because the promoter activity of class III flagellar genes is impaired in class II flagellar mutants due to an unknown regulatory mechanism imposed by the absence of the basal body, the transcription of class III-flgE-lacZ fusion was less active in ΔfliG (19 ± 1%) and ΔtipF (57 ± 1%) relative to WT (Fig. 2). Unlike the ΔfliG mutant (5 ± 0.5%), the class IV-fljL-lacZ fusion was as active in the ΔtipF mutant as in the WT background (87 ± 1%) (Fig. 2). These indirect in vivo assays suggest that class IV flagellar genes are efficiently transcribed in the ΔtipF mutant despite the absence of an assembled flagellum.

Figure 2.

 β-Galactosidase promoter-probe assays. LacZ activity of class II-fliF (MS-ring), class III-flgE (hook), and class IV-fljL (flagellin) in WT, ΔfliG, tipF, tipN, fliX∷Tn5, and flbD∷Tn5 strains was quantified as a percentage relative to WT.

fliX∷Tn5 and flbD∷Tn5 mutant strains were included as controls, while the ΔtipN mutant allowed for comparison with a strain that can possess multiple flagella that are frequently misplaced (Huitema et al., 2006; Lam et al., 2006). Subsequently, similar to canonical class II flagellar mutants, the class II-fliF-lacZ fusion was upregulated in the fliX∷Tn5 (142 ± 9%) and flbD∷Tn5 (316 ± 7%) mutants, while the class III-flgE-lacZ fusion (22 ± 2% and 19 ± 1%, respectively) and class IV-fljL-lacZ fusion (6 ± 0% and 5 ± 0%, respectively) were less active in the fliX∷Tn5 and flbD∷Tn5 strains when compared with the WT background (Fig. 2). Interestingly, the ΔtipN mutant transcribed class II-fliF-lacZ (146 ± 1%) and class III-flgE-lacZ (169 ± 2%) at higher levels than those observed in the WT background, while class IV-fljL-lacZ (112 ± 1%) was transcribed at levels near WT (Fig. 2). We speculate that the increased levels of flagellar gene transcription seen in the ΔtipN for class II-fliF and class III-flgE are a consequence of the multiple flagella present in the absence of TipN.

To validate the β-galactosidase promoter-probe assays, we relied on qChIP experiments to directly measure the in vivo occupancy of the transcriptional factors CtrA, FlbD, FliX, and RNAP at the fliF, flgE, and fljL promoters using polyclonal antibodies to CtrA, FlbD, and FliX, and a monoclonal antibody to the RpoC subunit of RNAP. The occupancy of flagellar promoters in ΔtipF was compared with WT, ΔfliG, ΔtipN, fliX∷Tn5, and flbD∷Tn5 mutants, with minor modifications (Radhakrishnan et al., 2008). Measurement of RNAP occupancy at the fliF promoter by qChIP corroborated the β-galactosidase results, with comparable trends being observed (i.e. elevated levels in ΔfliG) (Fig. 3). Notably, qChIP experiments revealed that CtrA occupied the fliF promoter at similar levels in ΔfliG and ΔtipF (99 ± 4% and 80 ± 6% relative to WT, respectively) (Fig. 3), indicating that the increase in class II flagellar gene transcription in ΔfliG and ΔtipF mutants is not due to an elevated occupancy of CtrA at the promoter(s). Consistent with fliF upregulation seen in ΔfliG and ΔtipF by the β-galactosidase assay, qChIP revealed that the occupancy of FlbD (repressing class II genes) was decreased at the fliF promoter in the ΔfliG (45 ± 1%) and ΔtipF (51 ± 8%) strains (Fig. 4a). FliX, the regulatory factor that links the status of flagellar assembly to FlbD activity (Muir & Gober, 2005), was present at the class II promoters, at higher levels than WT, in ΔfliG (170 ± 7%) and ΔtipF (144 ± 4%), consistent with the decreased levels of FlbD at the fliF promoter (Fig. 4a). FliX has been shown to interact with FlbD and block its access to enhancer DNA sequences in vitro (Dutton et al., 2005), and this new qChIP-based approach further suggests that FliX occupies the promoters to modulate FlbD activity at the class II-fliF promoter in vivo.

Figure 3.

 qChIP assays. The occupancy of the class II-fliF flagellar promoter by flagellar regulators CtrA and RNAP in vivo is shown for WT, fliG, tipF, tipN, fliX∷Tn5, and flbD∷Tn5 strains. Promoter regions were immunoprecipitated with a polyclonal antibody to CtrA or a monoclonal antibody to the RpoC subunit of RNAP before analyses by quantitative real-time PCR. Results demonstrate the percentage of promoter occupancy by CtrA and RNAP relative to WT.

Figure 4.

 qChIP assays. The occupancy of flagellar promoters by flagellar regulators FlbD and FliX in vivo for WT, fliG, tipF, tipN, fliX∷Tn5, and flbD∷Tn5 strains. (a) Class II-fliF (MS-ring). (b) Class III-flgE (hook). (c) Class IV-fljL (flagellin). Promoter regions were immunoprecipitated with polyclonal antibodies to FlbD or FliX, respectively, before analyses by quantitative real-time PCR. Results demonstrate the percentage of promoter occupancy by FlbD and FliX relative to WT.

Next, we determined the presence of FlbD and FliX at the class III-flgE and class IV-fljL promoters. qChIP showed that FlbD occupancy at the class III-flgE promoter was reduced in ΔfliG (68 ± 5%) and ΔtipF strains (75 ± 10%) (Fig. 4b), while that of FliX was elevated (155 ± 5% in ΔfliG and 227 ± 9% in ΔtipF) (Fig. 4b). These data demonstrate that the ΔtipF strain is similar to the ΔfliG mutant strain with regard to the occurrence of FlbD and FliX at the flgE promoter. It is further consistent with the view that FliX is also present at class III promoters to block FlbD access. The class IV-fljL promoter, however, had an abundance of FlbD similar to WT (123 ± 8%) and decreased levels of FliX (64 ± 7%) in ΔtipF, while the ΔfliG mutant had decreased FlbD (20 ± 2%) and increased FliX (200 ± 9%) (Fig. 4c). These results, also supported by the β-galactosidase promoter-probe assays (Fig. 2), suggest that, unlike FliG, TipF is not necessary to confer the transcription of class IV flagellar genes.

Both flbD∷Tn5 and fliX∷Tn5 mutant strains were included as controls. Accordingly, FlbD was considerably decreased at the fliF (7 ± 1%), flgE (22 ± 3%), and fljL (7 ± 1%) promoters in the flbD∷Tn5 mutant compared with WT (Fig. 4a–c). Similarly, the fliX∷Tn5 mutant had decreased levels of FliX at the fliF (8 ± 2%), flgE (15 ± 1%), and fljL (15 ± 1%) promoters (Fig. 4a–c).

The ΔtipN mutant possessed lowered levels of FlbD at the fliF (69 ± 5%) and flgE (57 ± 3%) promoters, while fljL (103 ± 9%) was near WT levels (Fig. 4a–c). FliX was present at the fliF (109 ± 8%), flgE (166 ± 9%), and fljL (129 ± 25%) promoters in the ΔtipN mutant relative to WT. Because the ΔtipN mutant frequently possesses multiple flagella that are often misplaced (Huitema et al., 2006; Lam et al., 2006), these results suggest that the levels of FlbD present at the class II/III promoters are sufficient to allow for flagellar development in the absence of TipN.

Our data demonstrate the in vivo occupancy of fliF, flgE, and fljL flagellar promoters by the transcriptional regulators CtrA, FlbD, and FliX of the C. crescentus WT and flagellar mutants for the first time, thus providing direct in vivo evidence for the previously proposed hierarchical scheme in the negative and positive transcriptional regulation of flagellar genes. While FlbD and FliX have been shown to interact, an inverse correlation was observed here between FlbD and FliX at the site of flagellar promoters in vivo, consistent with the hypothesis that FliX blocks FlbD access to flagellar promoters to regulate flagellar gene transcription.

The results from the transcriptional activity and promoter occupancy of flagellar regulators in the ΔtipF mutant suggest that tipF does not conform to the canonical flagellar hierarchy, akin to flgBC-fliE (Boyd & Gober, 2001) and fljK (Muir & Gober, 2005). We speculate that the transcriptional data presented point to hitherto unknown coupling mechanisms or interactions of TipF with regulatory components of the flagellar gene expression hierarchy, the cell cycle, and/or organizers of the flagellum assembly.

Acknowledgements

We thank the US Department of Energy, Office of Science (Biological and Environmental Research, grant DE-FG02-05ER64136) and the Mount Sinai Health Care Foundation for funding support and acknowledge William Davis for IT support and graphic design.

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