Effect of a ctrA promoter mutation, causing a reduction in CtrA abundance, on the cell cycle and development of Caulobacter crescentus
© Curtis et al.; licensee BioMed Central Ltd. 2013
Received: 18 October 2012
Accepted: 12 July 2013
Published: 18 July 2013
Polar development during the alphaproteobacterium Caulobacter crescentus cell cycle is integrated to the point that individual mutations can have pleiotropic effects on the synthesis of polar organelles. Disruption of the genes encoding the histidine kinase PleC, or its localization factor PodJ, disrupts synthesis or functionality of pili, flagella and adhesive holdfast. However, the mechanism by which these mutations affect polar development is not well understood. The aim of this study was to identify new regulators that control multiple aspects of polar organelle development.
To identify mutants with pleiotropic polar organelle synthesis defects, transposon mutagenesis was performed and mutants were selected based resistance to the pili-tropic bacteriophage ΦCbK. Mutants were then screened for defects in motility and holdfast production. Only a single podJ/pleC-independent mutant was isolated which had defects in all three phenotypes. Directed phage assays confirmed the phage resistance phenotype, while the strain demonstrated a similar dispersal radius as a podJ mutant in swarm agar, and treatment with a fluorescent lectin that labels the holdfast showed no staining for this mutant. The transposon had inserted into the promoter region of ctrA, a gene encoding a master transcriptional regulator of the cell cycle, disrupting native transcription but still allowing reduced transcriptional activity and protein production of this essential protein. Transcriptional fusions showed that essential genes controlled by CtrA exhibited minor to moderate changes in expression in the ctrA promoter mutant, while the pilA gene, encoding the subunit of the pilus filament, had a drastic decrease in gene expression. Introduction of a plasmid-born copy of ctrA under its native promoter complemented the phage resistance and holdfast defects, as well as a moderate cell morphology defect, but not the swarming defect.
A mutation was identified that caused pleiotropic defects in polar organelle synthesis, and revealed the surprising result that some CtrA-dependent promoters are more sensitive to changes in CtrA concentration than others. However, the fact that no pleiotropic mutations were found in new regulators suggests that downstream signaling of PleC/PodJ is either essential, redundant, or branching such that all three phenotypes were not simultaneously affected.
KeywordsCtrA P2 promoter PodJ Pleiotrophic
The best-studied asymmetrically dividing prokaryote is the alphaproteobacterium Caulobacter crescentus. At each cell division, predivisional cells of C. crescentus localize different structures at the cell poles: a single flagellum occupies the pole that will be inherited by the swarmer cell and pili are synthesized at this pole after division, whereas a narrow extension of the cell envelope (the stalk) tipped by an adhesive structure (the holdfast) occupies the opposite pole that will give rise to the stalked cell. The stalked cell is able to restart the cell cycle immediately after division, whereas the swarmer cell is unable to initiate DNA replication until it differentiates into a stalked cell.
The C. crescentus cell cycle and developmental program are controlled by three master regulators: CtrA, GcrA, and DnaA (for review, see ). These proteins are regulated such that each one reaches maximal abundance during a different stage of the cell cycle. DnaA reaches peak abundance at initiation of DNA replication occurring in stalked cells, GcrA peaks after DNA replication in early predivisional cells, and CtrA peaks in late predivisional and swarmer stages . All three proteins are required for regulating transcription of different suites of genes. DnaA activates genes involved in chromosome partitioning, nucleotide biosynthesis, and DNA replication, recombination and repair , and initiates replication of the chromosome. DnaA is also required for transcription of gcrA. GcrA activates transcription of genes involved in DNA replication, recombination and repair different from DnaA targets [3–5]. GcrA also activates genes required for polar development (including pleC and podJ, both of which are also activated by DnaA [3, 4]). CtrA, in turn, regulates at least 95 genes in 55 operons: some are repressed (for example gcrA and podJ[4, 6]) whereas others are activated (such as the pilin subunit gene pilA, flagellum synthesis cascade initiation, and the holdfast anchor operon ). Additionally, CtrA binds to the chromosome at the origin of replication where it represses the initiation of DNA replication . Furthermore, CtrA both activates and represses its own promoters.
The ctrA gene has two promoters: P1 and P2 . The weaker upstream P1 promoter is activated first. P1 activation requires that the promoter be in the hemi-methylated state, meaning that DNA replication has initiated and the replication fork has passed the P1 promoter. The P1 promoter is also directly activated by GcrA [4, 9, 10]. The low level of expression from the GcrA-activated ctrA P1 promoter allows some CtrA protein to accumulate. Once sufficient CtrA has accumulated, it represses the P1 promoter (as well as gcrA expression) and activates the strong downstream P2 promoter , leading to a burst of CtrA production and activity.
The sequential activation of the master regulators forms the timeline by which developmental processes are regulated and coordinated. In particular, GcrA contributes to the expression of the key developmental regulators, the histidine kinase PleC and the polar localization factor PodJ. Loss of either protein causes pleiotropic defects in development. A pleC mutant does not synthesize a stalk, holdfast or pili, and though the flagellum is made, flagellar rotation is not activated and the flagellum is not shed during the swarmer cell differentiation [11–13]. A podJ mutant, like pleC, does not synthesize holdfast or pili or shed its flagellum, but it does synthesize a stalk and activates its flagellum, however its motility is impaired in low-percentage agar as compared to wild type [6, 14, 15].
To further elucidate the pathways that lead to these pleiotropic phenotypes a genetic approach was used. We conducted a transposon mutagenesis screen, selecting for resistance to phage ΦCbK, which requires pili for infection, and screening for defects in motility and adhesion, which require the flagellum and holdfast respectively. In this work we report the identification of a transposon insertion in the promoter region of ctrA that causes a drastic reduction of CtrA accumulation, resulting in pleiotropic phenotypes bearing similarities to the pleC and podJ phenotypes.
Results and discussion
A transposon mutation causes a pleiotropic phenotype
Classes of ΦCbK-resistant mutants isolated
# of mutants
Class E (podJ)
Class F (pleC)
Class G (YB3558)
YB3558 transposon insertion is in the ctrA regulatory region
To verify this hypothesis, we generated a fusion of the ctrA mutant promoter from YB3558 to lacZ and compared expression from this promoter to the wild-type ctrA promoter in both CB15 and YB3558 during exponential growth (Figure 6B). Expression from the mutant promoter was only 20% of wild-type ctrA promoter expression in YB3558 and 29% wild-type ctrA promoter expression in the wild-type strain indicating that even when CtrA is present and its activity is normal (as it is in CB15), the mutant promoter is not efficiently transcribed.
Since the mutant ctrA promoter (containing the transposon insertion) from YB3558 demonstrated reduced activity in wild-type, suggesting ctrA transcription is reduced in YB3558, Western blot analysis was performed to measure CtrA abundance. Results showed that CtrA is expressed at a much lower level in YB3558 than in CB15 (Figure 6C). Subsequent quantification of band intensities from six Western blots showed that CtrA is present at approximately 22 +/− 5% of the wild-type CB15 level, demonstrating that the reduced transcription resulting from the transposon insertion leads to drastically lower CtrA protein levels.
Polar development defects are linked to altered CtrA abundance/activity
In order to determine if the lower CtrA levels are involved in the polar development defects found in YB3558, similar assays that were performed on YB3558 were also performed on ctrA401, a temperature sensitive CtrA allele . At the restrictive temperature the allele is lethal, but at the permissive temperature ctrA-dependent promoters demonstrate altered transcription patterns that indicate that CtrA401 has impaired function. Phenotypic analysis demonstrates that a ctrA401 mutant has a reduced swarming phenotype (Figure 1), as well as morphological defects (Figure 2), both of which mirror those of YB3558.
Plasmid pSAL14 was introduced into YB3558, creating strain YB3559. pSAL14 is a low copy plasmid carrying a copy of the ctrA gene with its native promoter . Introduction of the plasmid restored CtrA production to slightly above wild-type levels (Figure 6C). Phenotypic analysis of YB3559 demonstrated that ctrA complementation restores cell morphology (Figure 2) and holdfast synthesis (Figure 3) to wild-type phenotypes, and growth rate to near wild-type levels (Figure 5). Phage sensitivity was increased over that of the parent YB3558 (Figure 4), but not complemented to full wild-type levels (it should be noted pinprick-sized colonies are likely spontaneous suppressors). Interestingly, ctrA complementation appears to have no effect on the swarming defect of YB3558 (Figure 1). The causal relationship between reduced CtrA abundance and the reduced swarming phenotype in this mutant is unknown.
Effect of the ctrA promoter mutation on transcription of developmentally regulated genes
Though expression of the mutant ctrA promoter was reduced regardless of the strain harboring it (Figure 6B), the wild-type ctrA promoter displayed similar expression levels when placed in YB3558, indicating its activity is resistant to the severe reduction in CtrA protein levels in that strain. Given that CtrA is a global regulatory protein for both essential (e.g. cell division) and non-essential (e.g. polar development) genes, and that the drastic CtrA reduction in YB3558 leads to polar developmental defects but the strain is still viable, we hypothesized that transcription of CtrA-regulated genes essential for cell survival will be less affected by CtrA reduction in YB3558 than those that are essential for less important cellular functions. Thus we investigated the transcription level of several CtrA-regulated genes in CB15 and YB3558.
ftsZ and ftsQA promoters had a moderate reduction in activity, and the ccrM promoter had a slight reduction in activity. These genes are essential for viability. The moderate reduction in transcription for these genes agrees with the hypothesis that genes involved in essential cell cycle processes would not be severely affected by the reduction in CtrA in YB3558. In contrast, the pilA promoter exhibited a drastic decrease in activity, as would be expected given the selection by which this mutant was obtained. However, activity from the fliQ promoter (fliQ is a flagellar biosynthesis gene and not essential) was largely unaffected. It is not clear why this promoter is unaffected while the pilA promoter shows such a difference in activity. It could be that the pilA promoter is much more sensitive to CtrA levels.
Regulation of pilA is controlled not only by CtrA, but by SciP. SciP interacts with CtrA to prevent transcription of genes positively regulated by CtrA, such as pilA, in swarmer and late predivisional cells . It is possible that the dramatic decrease in pilA promoter activity in YB3558 is not from CtrA abundance itself, but an indirect effect of reduced CtrA abundance leading to increased SciP activity. However, CtrA positively regulates transcription of sciP and the strong reduction of CtrA activity in the YB3558 mutant should lead to a decrease in SciP levels, not an increase. In agreement with this hypothesis it has been shown that a site-directed mutation that abolishes transcription from the ctrA P1 promoter caused a strong reduction of CtrA abundance , similar to that of the YB3558 mutation in this study, and this lead to significantly reduced expression of SciP, down to 19% of wild-type level . The ctrA P1 mutant also had morphological and growth defects similar to those found here, and several assays demonstrated that CcrM transcription and translation was largely unaffected, agreeing with our results. Therefore it is unlikely that the effects observed on gene expression are the result of increased SciP activity.
CtrA binding sites for CtrA-regulated genes
CtrA binding site
xxxxTTAA xxxxxxxTTAA xxx
CCATTAA CCAGTCTTAA ATTAA CTC
CAGTTAA CCGCCGATTAA CGA
CCGTTAT GACGACATTAA CGA
TGGTTAA CGGCCCGCTAA CCA
CCCCTAA CGCCCTGTTAA CCA
CTGTTTA CTGGCCATTAA GTG
TGGTTAA GAACAAATAA CGGTAAA TACAAATAAA CCA
TGGTCAA CAAAAGACTAA AAT
Though the genes used for analysis in this study mostly have single CtrA-binding sites close to the consensus, the pilA gene, which displays drastically reduced transcription in YB3558 compared to wild-type, appears different compared to the other genes presented in regards to CtrA regulation. CtrA was shown to the bind to three distinct regions in the pilA promoter area. Region 1 has a TTTA-N7-TTAA binding site straddling the −35 site. Region 2, 19 bp upstream of Region 1, has two potential CtrA binding sites, TTAA-N6-ATAA and TAAA-N6-TAAA, separated by 3 bp. Region 3, 71 bp upstream of Region 2, has a single TCAA-N7-CTAA binding site. Though the Region 1 binding site is relatively close to the consensus sequence, all the other binding sites diverge greatly from the consensus in sequence and/or half-site spacing. Clearly CtrA regulation of pilA is more complex than that of the other genes presented. Perhaps the divergent binding sites have low affinity for CtrA and the multiple weak binding sites create cooperative CtrA binding necessary to achieve maximal pilA expression. It would be plausible that this scenario (multiple weak sites working together) would be quite sensitive to changes in CtrA protein levels, leading to the drastic reduction in transcription seen in YB35587. Further analysis of CtrA regulation of pilA will prove informative.
Is it possible that promoters more susceptible to changes in CtrA concentration/activity account for all the pleiotropic defects observed in podJ and pleC strains? Current understanding of PleC’s role (and thus PodJ’s) in developmental signaling is to regulate phosphorylation levels of another signaling protein DivK, which in turn regulates the activity of the CckA phosphorelay that controls CtrA activation [28, 29]. A pleC mutant should have reduced CtrA levels, similar to the CtrA phenotype found in this study. Though CtrA protein levels in pleC are similar to wild-type, there is a significant decrease in CtrA phosphorylation . Also in agreement with this hypothesis, reduced CtrA levels have been implicated as contributing to the null-pili phenotype of podJ mutants . However, the other polar development phenotypes are not as well explained by CtrA-promoter effects. The known link between CtrA and flagellar motility in C. crescentus is that CtrA initiates the flagellum synthesis cascade . The fliQ-lacZ reporter demonstrates that the synthesis cascade is unaffected, which agrees with the fact that both pleC and podJ mutants produce flagella. CtrA must affect motility in a way other than synthesis of the flagellum, possibly two ways since the flagellum is paralyzed in a pleC mutant but capable of rotation in a podJ mutant. The effect of CtrA on motility appears to be independent of CtrA abundance as complementation of CtrA abundance by pSAL14 failed to restore wild-type motility to YB3558 (Figure 1). If the effect is not dependent on CtrA abundance, it may be dependent on timing of CtrA activity. Expression from the mutant promoter in YB3558 is likely constitutive, and may lead to early induction of whatever CtrA-dependent pathway is involved in motility other than flagellum synthesis. However, the CckA/ChpT pathway that controls CtrA activity should not be perturbed in this mutant, so even though CtrA could be produced constitutively, its activity should still be properly regulated. The full link between CtrA and motility is still a mystery.
The connection between CtrA and holdfast synthesis is also not clear. While it is known that at least some of the holdfast synthesis genes display changes in transcription activity during the cell cycle , and microarray experiments have shown that holdfast genes have altered transcription in a ctrA mutant [7, 33], it has also been shown that holdfast synthesis can be stimulated in swarmer cells when they contact a surface , and that developmental holdfast synthesis is also likely regulated by cyclic-di-GMP levels . We have recently shown that the holdfast synthesis and anchoring machineries are synthesized and polarly localized in predivisional cells in preparation for holdfast synthesis in the next cell cycle [36, 37]. Therefore, it is likely that CtrA regulates the synthesis of the holdfast synthesis-anchoring machinery in predivisional cells, but that the activation of this machinery is regulated by surface contact and developmental signals. The additional possibility that CtrA abundance effects post-transcriptional regulation of holdfast synthesis cannot be ruled out. However, both effects on motility and post-transcriptional effects on holdfast synthesis could be downstream effects of CtrA-dependent decrease in promoter activity of one or more other regulators.
In this study we performed a detailed mutagenesis selection/screen to identify new regulators that control multiple aspects of polar development similar to known developmental regulators PleC and PodJ. Our results suggest that potential regulators downstream of those already known may be essential, redundant or branched. In the process we found evidence that suggests at least some of the pleiotrophic phenotypes are the result different affinities between CtrA and CtrA-dependent promoters.
Media and growth conditions
All C. crescentus strains were grown at 30°C in peptone yeast extract (PYE) media . When appropriate, kanamycin (5 μg/ml liquid, 20 μg/ml solid), chloramphenicol (0.5 μg/ml liquid or 1 μg/ml solid), tetracycline (1 μg/ml liquid or 2 μg/ml solid) and nalidixic acid (20 μg/ml) were used. Escherichia coli strains were grown at 37°C in Luria-Bertani (LB) medium  with kanamycin (50 μg/ml), chloramphenicol (20 μg/ml liquid or 30 μg/ml solid), ampicillin (50 μg/ml liquid or 100 μg/ml solid), or tetracycline (12 μg/ml liquid or 12 μg/ml solid).
Transposon mutagenesis and selection of ΦCbKR mutants
The plasmid pFD1 , carrying the mariner transposon and the transposase gene, was introduced into C. crescentus strain CB15 (wild-type) by conjugation with E. coli strain YB2028 (SM10λpir (pFD1)). Cells from five independent conjugations were pooled and frozen at -80°C. Aliquots of cells were thawed, mixed with undiluted Caulobacter phage ΦCbK stock (~1010 pfu/ml), plated on PYE supplemented with kanamycin and nalidixic acid and incubated at 30°C for several days until KanR ΦCbKR colonies appeared.
Overnight cultures of all ΦCbKR mutants were observed with a 100× objective on a Nikon Optiphot-2 microscope. Strains were qualitatively scored on three phenotypes: presence of rosettes, presence of stalks, and presence of motile swarmer cells.
Strains were grown overnight, normalized to equal OD600 and diluted to 100, 10-4 and 10-5. Cell dilutions were mixed in equal volumes with ΦCbK (~1010 pfu/ml) or plain PYE. The mixture was incubated at room temp for 10 minutes, then 5 μl spots were placed onto PYE plates. The plates were incubated at 30°C for 3–5 days. Relative resistance was determined by the number and size of colonies that appeared.
Confirmation of transposon mutant phenotypes and identification of genes
The kanamycin marker in strains of interest were transduced into C. crescentus strain CB15 with the phage ΦCr30, using a standard transduction protocol . KanR colonies were isolated and overnight liquid cultures were shown to have the same phenotype as the parent strain.
Genomic DNA was isolated using a phenol/chloroform extraction method. Briefly, cells were grown overnight at 30°C in 3 ml PYE + kanamycin. The entire culture was pelleted by centrifugation, and resuspended in cold TE pH 7.5 to a final volume of 500 μl. Lysozyme (Sigma) and RNAse (Amresco) were added to final concentrations of 1 mg/ml and 0.1 mg/ml respectively, and the mixture was incubated at 37°C for 30 min before adding 0.1 volumes of 10% w/v SDS. Proteinase K (Amresco) was added to a final concentration of 1 mg/ml. The solution was mixed gently and incubated at 50°C for 2 hours with occasional mixing. 50 μl of sodium chloride (5 M) was added, and the DNA was extracted 3–4 times with one volume of phenol/chloroform/isoamyl alcohol (25:24:1), followed by two extractions with one volume of chloroform. After extraction, DNA was precipitated with 0.6 volumes of isopropanol, washed twice with 70% v/v ethanol, allowed to dry, and resuspended in 50 μl dH2O.
Southern blot analysis
Primers used in this study
For strains of interest that did not have insertions in podJ or pleC, genomic DNA (~3 μg) was digested with Pst I and separated on an agarose gel. DNA was excised from the gel area found to include the band seen by Southern analysis using a probe for the kanamycin resistance gene. The DNA was isolated from the gel using the Qiaquick Gel Extraction kit (Qiagen) and ligated to Pst I-digested pKSII+ (Stratagene) overnight at 16°C. The ligation was electroporated into E. coli strain DH5α (F’, ϕ80dlacZ ΔM15, Δ(lacZYA-argF)U169, endA1, recA1, hsdR17 (rk-, mk+), deoR, thi-1, supE44, λ-, gyrA96, relA1). AmpR KanR colonies were isolated, and plasmid DNA was purified.
Plasmids were sequenced with primer MarRseq (Table 3) using Big Dye version 3.1 (Applied Biosystems), and run on an ABI3730 DNA Analyzer at the Indiana Molecular Biology Institute (Indiana University). The transposon insertion site was identified in the sequence, and the gene was identified by a Basic Local Alignment Search Tool (BLAST) search against the C. crescentus genome (TIGR - http://blast.ncbi.nlm.nih.gov/Blast.cgi?PAGE_TYPE=BlastSearch&PROG_DEF=blastn&BLAST_PROG_DEF=megaBlast&BLAST_SPEC=MicrobialGenomes_155892&DB_GROUP=AllMG).
Characterization of the YB3558 mutant
Cultures of YB3558 were grown overnight in PYE with kanamycin, diluted to an OD600 of approximately 0.15, and allowed to grow to an OD600 of 0.5-0.6, then observed using 100X Plan Apo objective on a Nikon Eclipse E800 microscope. Images were captured using a Princeton Instruments 1317 cooled CCD camera and processed with Metamorph v. 4.5 (Universal Imaging Corporation). Staining of holdfast with fluorescein isothiocyanate-wheat germ agglutinin (FITC-WGA) was performed as described previously . Fluorescence was observed on the Nikon E800 and images were processed using Metamorph.
Strains were grown overnight in PYE supplemented with appropriate antibiotics and diluted to an OD600 of 0.1 in fresh PYE with no antibiotic. They were allowed to grow for two doublings (to OD600 of ~0.4) and diluted again to an OD600 of 0.05 in 10 ml of PYE. 100 μl of the culture was removed and its OD600 recorded every 30 minutes for 5 hours.
Strains were grown overnight in PYE supplemented with appropriate antibiotics, diluted to an OD600 of 0.1, and allowed to grow for two doublings (to OD600 of ~0.4). All strains were diluted to an equal OD600 and 1 μl of the culture was injected into a 0.3% Agar PYE plate. This was incubated at room temperature for 5–7 days in a humid container.
Plasmid pSAL14 , carrying a wild-type copy of the ctrA gene, was transformed into YB3558. The resulting strain, YB3559, was assayed for complementation of the phenotypes seen in YB3558.
To examine levels of CtrA in mixed culture, exponentially growing cells were collected and resuspended to equal OD600 in a final volume of 100 μl in 1X SDS loading buffer (62.5 mM Tris–HCl pH 6.8, 10% v/v glycerol, 2% w/v SDS, 0.05% v/v β-mercaptoethanol, 0.0025% w/v Bromophenol blue). 15 μl of this sample was separated on a 10% SDS-polyacrylamide gel and transferred to a nitrocellulose membrane. The membrane was probed with α-CtrA serum  at 1:10,000 dilution. The membrane was then probed with HRP-conjugated goat anti-rabbit secondary antibody (Biorad) at 1:20,000, developed using Supersignal Pico (Pierce) and imaged on a Kodak imagestation 440CF. For quantification of CtrA levels in wild-type and mutant strains, four replicates of each sample were loaded on one gel and treated as described above. Once exposed, Kodak Molecular Imaging Software version 4.0.3 was used to quantify the intensity of each band and band intensities were averaged for wild-type and mutant.
lacZ fusions of wild-type and mutant ctrA promoters
The ctrAP2::Mn promoter was PCR amplified using the primers M134UP and M134DN (Table 3), incorporating EcoRI and XbaI restriction sites, respectively. The wild-type promoter was amplified using the primers M134DN and CtrAlacUp (Table 3). The digested fragments containing the promoter regions were cloned into the lacZ containing plasmid pLac290 .
Plasmids carrying promoter fusions to lacZ were transferred to YB3558 and CB15 by conjugal mating. The resulting transformants were grown to an OD600 of 0.4 to 0.6 in liquid PYE supplemented with tetracycline. Cells were added to three tubes containing Z-buffer (60 mM sodium phosphate (dibasic), 40 mM sodium phosphate (monobasic), pH 7.0, 10 mM potassium chloride, 1 mM magnesium sulfate, 50 mM β-mercaptoethanol) to a final volume of 800 μl, and 25 μl 0.1% w/v SDS was added. To assay the LacZ activity in the samples, 200 μl of 2-nitrophenyl β-D-galactopyranoside (ONPG) (4 mg/ml in 0.1 M potassium phosphate buffer, pH 7.0) was added to each sample, and reactions were stopped by the addition of 400 μl 1 M sodium carbonate. The time between addition of ONPG and stopping the reaction was recorded. Samples were centrifuged for 5 minutes to pellet cells and debris, then the absorbance at 420 nm (A420) of each sample was measured and recorded. β-galactosidase activity was calculated using the formula (A420 × 1000)/(OD600 × time (min) × volume of cells used (ml)).
This work was supported by grant GM51986 from the National Institutes of Health to YVB. PDC was supported by a postdoctoral National Institutes of Health National Research Service Award F32GM084618 from the National Institute of General Medical Sciences. DK was supported by a National Institutes of Health Predoctoral Fellowship (GM07757).
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