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J Bacteriol. Nov 2006; 188(22): 7932–7940.
Published online Sep 8, 2006. doi:  10.1128/JB.00964-06
PMCID: PMC1636300

Two Chemosensory Operons of Rhodobacter sphaeroides Are Regulated Independently by Sigma 28 and Sigma 54[down-pointing small open triangle]


Rhodobacter sphaeroides has a complex chemosensory system, with several loci encoding multiple homologues of the components required for chemosensing in Escherichia coli. The operons cheOp2 and cheOp3 each encode complete pathways, and both are essential for chemosensing. The components of cheOp2 are predominantly localized to the cell pole, whereas those encoded by cheOp3 are predominantly targeted to a discrete cluster in the cytoplasm. Here we show that the expression of the two pathways is regulated independently. Overlapping promoters recognized by σ28 and σ70 RNAP holoenzyme transcribe cheOp2, whereas cheOp3 is regulated by one of the four σ54 homologues, RpoN3. The different regulation of these operons may reflect the need for balancing responses to extra- and intracellular signals under different growth conditions.

Many motile bacteria sense and respond to changes in their environment by using a variation of the Escherichia coli chemotaxis paradigm (for a review, see reference 27). This two-component signal transduction pathway responds to changes in the concentration of specific ligands via transmembrane methyl-accepting chemotaxis proteins (MCPs). These proteins, together with the linker protein CheW, regulate the activity of a sensor kinase, CheA, which autophosphorylates on a conserved histidine residue. This phosphoryl group is transferred to an aspartate residue on a response regulator CheY. CheY-P diffuses to the flagellar motor, where it binds to the switch protein FliM, altering the direction of flagellar motor rotation and ultimately causing a change in swimming direction. Receptor adaptation is mediated by a constitutively active methyltransferase, CheR and a second response regulator, the methylesterase CheB, which work antagonistically to modify the methylation state of the MCPs. CheB activity is also regulated by CheA-dependent phosphorylation.

Rhodobacter sphaeroides is an α-subgroup, purple nonsulfur, photosynthetic bacterium. It has multiple homologues of the E. coli chemotaxis proteins that are expressed from three operons and other unlinked loci. Although a role for cheOp1 has yet to be identified under laboratory conditions, cheOp2 and cheOp3 are both essential for chemotaxis (6, 12, 13, 18). cheOp2 contains the genes for the putative phosphatase, cheP2, cheY3, cheA2, cheW2, cheW3, cheR2, and cheB1, and a cytoplasmic chemoreceptor, tlpC. cheOp3 contains cheA4, cheR3, cheB2, cheW4, the partitioning factor ppfA, tlpT, cheY6, and cheA3. Interestingly, whereas the genes encoding membrane-spanning chemoreceptors are scattered around the genome, those encoding the soluble chemoreceptors (Tlps) are within the chemotaxis operons. The chemosensory proteins encoded in cheOp2 are predominantly clustered with the MCPs at the cell poles, whereas the proteins encoded in cheOp3 are targeted with Tlps to a discrete cytoplasmic locus (28). This suggests that the pathway encoded by cheOp2 is responsible for sensing extracellular signals, whereas that encoded by cheOp3 responds to cytoplasmic signals.

R. sphaeroides has a versatile metabolism enabling growth in many environmental conditions. There is evidence to suggest that the bacterium is able to customize the expression of its chemosensory machinery according to the prevailing growth conditions, for example, during aerobic growth cheOp2 is expressed at much higher levels than during photoheterotrophic growth (21). However, there are no data on how either cheOp2 or cheOp3 are regulated and no detailed mapping of the promoter regions. The majority of bacterial gene transcription is affected by core RNA polymerase bound to the constitutive “housekeeping” specificity factor, σ70. However, alternative sigma factors are often used to facilitate promoter recognition and transcription of specific regulons. Although expression of early flagellar genes is regulated by a σ70 master operon, these in turn regulate expression of σ28 ensuring regulated sequential expression of the late flagellar and chemotaxis genes in Bacillus subtilis, E. coli, and many other motile bacterial species (4, 7). σ54 (RpoN) proteins are generally constitutive but require an enhancer binding protein (EBP), generally activated by specific growth conditions, (reviewed in reference 5). Although σ54 was first implicated in the regulation of nitrogen metabolism, σ54 homologues have been found to regulate many other functions, including chemotaxis gene expression in Rhodospirillum centenum (9). The genome sequence of R. sphaeroides has revealed 16 genes for putative alternative sigma factors, including 1 for σ28 (fliA) and 4 encoding σ54 (rpoN). This is the only species thus far identified to encode four, apparently noninterchangeable σ54 proteins (14, 16). This suggests that they each regulate promoters interacting with different EBPs and therefore presumably respond to different cellular signals. It has been shown that, as with other species, the expression of the late flagellum genes is regulated by σ28, whereas the expression of the earlier genes depends on one of the four σ54 proteins (RpoN2) and two EBPs (17).

In the present study we show that cheOp2 is regulated by σ28 in addition to the “housekeeping” σ70, whereas cheOp3 is regulated by RpoN3, distinct from the RpoN2 involved in flagellum gene expression.


Strains and growth conditions.

Bacterial strains and plasmids are shown in Table Table1.1. R. sphaeroides strains were grown microaerobically in succinate medium (22) at 30°C with slow shaking. The expression of cheOp2 and cheOp3 is highest under microaerobic growth conditions (21; unpublished observations). E. coli strains were grown in Luria-Bertani medium at 37°C with shaking. DH5α was used for all molecular cloning procedures and S17-1λpir was used for conjugal transfer into R. sphaeroides. Nalidixic acid, kanamycin, and streptomycin were used at 25 μg ml−1 and ampicillin was used at 100 μg ml−1 when appropriate.

Strains and plasmids

Molecular genetic techniques.

All cloning steps were performed as described by Sambrook and Russell (19). Sequencing quality DNA was extracted by using the Plasmid Midi-Kit (QIAGEN) or the WizardPlus kit (Promega), sequenced by the University of Oxford Biochemistry sequencing service and analyzed with Clone Manager version 7.10 (Sci Ed Central). All primers were from SigmaGenosys. PCRs were performed with Pfu DNA polymerase (Promega) according to the manufacturer's directions. Products were purified from agarose gels with QiaexII (QIAGEN) or GenElute (Sigma).

Construction of mutant strains.

DNA upstream of the gene or region to be deleted was amplified by PCR, using primers that introduced EcoRI and XbaI restriction sites, and cloned into pK18mobsacB (20). Similarly, DNA downstream of the gene or region to be deleted was amplified using primers that introduced XbaI and HindIII restriction sites. These were cloned into the XbaI and HindIII sites of the appropriate pK18mobsacB derivatives containing the upstream sequences. A similar strategy was used for the construction of the orfE omega insertion construct. The first 9 bp of orfE and upstream sequences was amplified by PCR using primers that introduced EcoRI and SmaI restriction sites and the product was cloned into pK18mobsacB. The remainder of orfE and downstream sequences was amplified by using primers that introduced SmaI and XbaI restriction sites. The product was cloned into the SmaI and XbaI sites of the pK18mobsacB derivative containing the upstream sequences. Finally, the omega cartridge was cloned into the engineered SmaI site. All constructs were sequenced to ensure that there had been no PCR misincorporation errors and that, when appropriate, they were in frame. The constructs were introduced into the genome by allelic exchange to create unmarked deletion mutants as described previously (6, 20).

Phenotypic analysis.

Immunoblotting of strains was performed as described elsewhere (11). Swarm plates containing 0.25% agar (BiTek; Difco) and 100 μM propionate were performed as described previously (12) and incubated for 48 h microaerobically before the swarm diameters were recorded. Growth rates were measured as described in Martin et al. (12). For electron microscopy, bacterial cells on Formvar-coated Ni grids were negatively stained with 1.7% phosphotungstic acid and examined in a Philips EM410 transmission electron microscope.

Mapping of the transcriptional start.

Transcriptional start sites were mapped by the primer extension of mRNA isolated from R. sphaeroides WS8N cells grown microaerobically to an optical density at 700 nm of 0.6. Transcription was halted through the addition of RNAlater (Ambion), and total RNA was isolated by using lysozyme and the GramCracker kit (Ambion) according to the manufacturer's instructions. Then, 30 pmol of primer was labeled with 32P using T4 polynucleotide kinase (Ambion) and was extended by using 4 U of Omniscript reverse transcriptase (QIAGEN) for 1 h at 37°C. The resultant cDNA was precipitated, suspended in formamide loading dye (95% formamide, 20 mM EDTA, 0.05% bromophenol blue, 0.05% xylene cyanol), heated to 85°C for 3 min, and quickly cooled on wet ice before being electrophoresed on a 6% sequencing gel alongside a double-stranded DNA sequencing reaction prepared with the same primer and a Sequenase Version 2 DNA sequencing kit (USB). Gels were dried and either exposed to film (Amersham Hyperfilm) or analyzed on a PhosphorImager with ImageQuant version 5.0 software (Molecular Dynamics). The sequence of primer P1NEW used for mapping the start of cheOp2 was 5′-GGAGAATCGTCCACGGCGAG-3′.


Total RNA was isolated from R. sphaeroides WS8N cells as described above. Contaminating DNA was digested with DNase I (Amersham) for 1 h at 37°C and removed with the RNeasy kit (QIAGEN). Reverse transcription-PCR (RT-PCR) analyses were performed with 2 μg of RNA, the one-step RT-PCR kit (QIAGEN) and primers designed to yield a tlpC cDNA product of 259 bp and a tlpT cDNA product of 298 bp. Thermocycling was carried out in a Thermal Mini-Cycler (MJ Research) as follows: step 1, 50°C, 30 min; step 2, 95°C, 15 min; step 3.1, 94°C, 1 min; step 3.2, 65°C, 1 min; step 3.3, 72°C, 1 min (25 cycles); and step 4, 72°C, 10 min. Products were analyzed by electrophoresis on a 2.5% agarose gel and visualized after incubation with SYBR Green (Molecular Probes) on a FLA-3000 analyzer (Fuji). To control for contaminating DNA, PCRs were also performed on the prepared RNA using Pfu DNA polymerase (Promega) and cycling conditions as described above with a 10-min incubation at 98°C instead of steps 1 and 2. The primers used for the amplification of tlpT cDNA were 5′-GCTGCTTCGAGGCCATCG-3′ and 5′-GACGGTCTGCGCGGTCTG-3′ and for the amplification of tlpC cDNA were 5′-GCCACCTGACCTCCGACG-3′ and 5′-CTCGACCGTCTGCGCTCG-3′.

LacZ fusion assay.

The broad-host-range vector pUI523A permits translational fusions with a promoterless lacZ gene (25). The copy number of this tetracycline-resistant plasmid is 4 to 6 in R. sphaeroides and does not vary with the growth condition. For the analysis of cheOp2 expression, a 1.6-kb fragment comprising the 5′ end of cheY3 and upstream regions was cloned in each orientation into pUI523A to generate a CheY3-LacZ fusion product expressed from the promoter(s) within the insert (21). For ease of use, the omega cartridge, which confers streptomycin resistance, was inserted upstream of this fusion, into the opposite orientation control plasmid, and into pUI523A without an insert to form pACM37, pACM38, and pACM27, respectively. Similarly, for the analysis of cheOp3 expression, a 0.5-kb fragment comprising the 5′ end of cheA4 and upstream regions was cloned in each orientation into pUI523A so as to generate a CheA4-LacZ fusion product. After the introduction of the streptomycin resistance (Smr) cartridge upstream of these fusions, the resultant plasmids were designated pACM76 and pACM77 for the forward and reverse orientations, respectively. The plasmids were introduced into R. sphaeroides WS8N by conjugation. Cells were harvested at an optical density at 700 nm of 0.6, lysed by sonication, and β-galactosidase activity was determined by using the Promega assay system according to the manufacturer's instructions.

Purification of σ28 and the electrophoretic mobility shift assay.

The gene encoding σ28 (lacking the ATG start codon) was amplified to include 5′ and 3′ restriction sites and ligated in frame into the expression vector pQE80. This expression vector introduces a six-histidine N-terminal tag to the gene product and allows control of expression through the IPTG (isopropyl-β-d-thiogalactopyranoside)-inducible tac promoter. σ28 protein was purified on a nickel affinity column (QIAGEN) and was reconstituted to purified core RNA polymerase (a gift from T. Donohue [1]) in buffer containing 100 mM NaCl, 10 mM MgCl2, 100 μg of bovine serum albumin/ml, 5% glycerol, 20 mM Tris (pH 8.0), and 1 mM dithiothreitol so that there was a 15-fold molar excess of σ28 over core enzyme.

A DNA fragment encompassing bases −91 to +85 relative to the σ28-dependent transcriptional start of cheOp2 was amplified by PCR so as to include 5′ and 3′ KpnI restriction sites and sequenced. Digestion with KpnI yielded 3′ overhangs, which were labeled with biotin as described by the manufacturer (Pierce). The electrophoretic mobility shift assay was performed by using the LightShift Chemiluminescent EMSA kit (Pierce) as recommended by the manufacturer using 3 ng of labeled DNA, 1.6 μg of protein, and a 100-fold excess of competitor (unlabeled) DNA.

Tn mutagenesis.

The plasmid pBR322 harboring the transposon Isomegokmhah was mobilized into R. sphaeroides WS8N from E. coli S17-1λpir as described previously (6, 20). Chemotaxis mutants were identified by their reduced movement through soft nutrient agar swarm plates (12). The position of the transposon (Tn) was determined by DNA sequencing.


cheOp2: identification of transcriptional start sites.

LacZ translational fusions showed that a region of DNA 1.6 kb upstream of the cheY3 gene of cheOp2 (Fig. (Fig.1A)1A) contains the promoter sequences responsible for its expression (21). This region was scanned for promoter recognition sequences. Two motifs were identified (TCTCGC and TAGCGT separated by 14 bp) 86 and 66 bp upstream of the translational start, with some similarity to the known, poorly conserved, σ70 promoter recognition sequences in R. sphaeroides (Fig. (Fig.1B)1B) (24, 26). In addition, two motifs with 66% identity to the −35 and −10 σ28 consensus recognition sequences of E. coli (2, 8) were identified (TAAAT and GCCGTTCT separated by 14 bp), 99 and 80 bp from the translational start (Fig. (Fig.1B1B).

FIG. 1.
(A) Diagram showing the architecture of cheOp2. Each gene is represented by large gray arrows and is labeled individually. The region contained within the lacZ construct pACM37 is indicated. (B) Diagram illustrating the promoter region of cheOp2. The ...

Primer extension was performed to determine the transcriptional start site(s). Using a primer which bound within cheY3 and mRNA prepared from WS8N, two primer extension products of equal intensity were identified (Fig. (Fig.2).2). One product ran level with a T residue in the ladder and was 8 bp downstream of the −10 element of the putative σ28 promoter and 65 bp upstream of the translational start of CheP2. The other also ran level with a T residue in the ladder and was 9 bp downstream of the −10 element of the putative σ70 promoter and 52 bp upstream of the translational start of CheP2. There were no bands in the tRNA control lane, indicating that the primer did not hybridize nonspecifically. Since the start sites are appropriate distances from each of the proposed promoters and the initiating residue of each is an adenine, it suggests there are two overlapping promoters regulating cheOp2 expression.

FIG. 2.
Mapping of the transcriptional start sites of cheOp2 by primer extension analysis. Lanes containing tRNA (negative control) and WS8N mRNA are marked. A sequencing ladder of a pUC19 derivative containing the putative promoter region (in the order ACGT) ...

cheOp2: mutagenesis of promoter regions.

Unmarked mutants containing deletions of the putative promoter regions upstream of cheOp2 were constructed based on the data from the primer extension experiments and on the homology to known promoter recognition sequences (Fig. (Fig.1B).1B). The mutants in the cheOp2 putative promoter region were JPA455 with an 11-bp deletion, including the putative −35 recognition sequence of σ28; JPA459 with a 49-bp deletion encompassing both the putative −35 and the −10 σ28 recognition sequences (these also overlap with the putative σ70 −35 recognition sequence); and JPA461 with a 62-bp deletion removing all putative promoter recognition sequences. The ability of these strains to respond to a gradient of the chemoattractant propionate on swarm plates was compared to the wild-type strain, to the nonchemotactic strain ΔcheOp3 (JPA1301), and to the nonmotile strain cheY6 D57N (JPA1213) (Fig. (Fig.3a).3a). The swarm diameters of all of the putative promoter mutants were comparable to the nonchemotactic strain JPA1301 and were not caused by changes in the growth rates (data not shown). This suggests that the regions deleted contained the promoter sequences for these operons.

FIG. 3.
(a) Comparison of the swarm diameters of the wild-type strain WS8N, the putative cheOp2 promoter mutant strains JPA455, JPA459, and JPA461, and the nonmotile JPA1213 and nonchemotactic JPA1301 controls in response to 100 μM propionate under microaerobic ...

The expression of CheW2 from cheOp2 was examined in WS8N and JPA455 under microaerobic conditions by immunoblotting with anti-CheW2 antibodies. The expression of CheW2 was completely abolished in strain JPA455 (Fig. (Fig.3b).3b). Again, this suggests that the promoter sequences for cheOp2 expression have been deleted in this strain.

cheOp2: involvement of σ28 in transcription.

Since the promoter regions appear to overlap, deletion of the σ28 recognition site in JPA455 probably also altered the architecture of the σ70 site. The elimination of transcription from the σ70 promoter would account for the total abolition of both CheW2 expression and chemotaxis. To confirm that σ28 is involved in the transcription of cheOp2, the gene encoding σ28, fliA, and flanking sequences was cloned from a cosmid library of R. sphaeroides WS8N chromosomal DNA using a probe based on the published sequence of R. sphaeroides 2.4.1. An in-frame, unmarked deletion was constructed (which did not affect the stop codon of an overlapping gene) and designated JPA467. The strain was nonmotile and (as determined by electron microscopy) nonflagellate (data not shown). This was expected, since flagellin genes require σ28 for transcription. The growth rate of the mutant was normal (data not shown).

The expression of the cheOp2-encoded CheW2 was assessed in the σ28 deletion mutant by immunoblot analysis with anti-CheW2 antibodies. The expression of CheW2 was reduced to approximately a third relative to the wild-type (Fig. (Fig.3c),3c), suggesting a key role for σ28 in the regulation of cheOp2 expression.

To further determine the effect of the fliA deletion on the expression of cheOp2, Smr derivatives of the lacZ translational fusions used previously (21) were constructed. These derivatives of the broad-host-range vector pUI523A permit fusions with a promoterless lacZ gene. The resultant plasmid pACM37 generated a CheY3-LacZ fusion product expressed from the promoter(s) within the insert. The expression of lacZ from this plasmid and a reverse orientation control (pACM38) conjugated into the σ28 deletion mutant and the wild-type WS8N strain was assayed. β-Galactosidase production (in Miller units) in the mutant strain was reduced by one-third compared to the wild type (Fig. (Fig.3d3d).

Similar plasmids were constructed to assess whether the expression of cheOp3 was affected by the deletion of σ28. A 570-bp region of DNA containing the 5′ end of cheA4 and upstream sequences was cloned in each orientation into pUI523A, and the Smr Ω cartridge was subsequently ligated upstream of each insert to form pACM76 and the control plasmid pACM77. The expression of lacZ from these plasmids in the σ28 mutant was similar to that of the wild-type strain (data not shown). Taken together, these data suggest that whereas σ70 is the principal sigma factor, σ28 contributes significantly to the expression of cheOp2 but not of cheOp3.

If the expression of cheOp2 is partly regulated by σ28, the primer extension band relating to the transcript produced from this promoter should be absent from the σ28 deletion strain. Primer extension was repeated using mRNA from the mutant strain (JPA467) and the wild-type WS8N (Fig. (Fig.4).4). For a more rigorous negative control, a primer extension reaction using mRNA from the strain in which 11 bp had been deleted from a region that encompassed the putative σ28 −35 promoter sequence (JPA455) was used. The top band, which represents cDNA from the σ28 regulated transcript, was present in the wild-type reaction but absent from the σ28 deletion mutant lane. There were no bands in the negative control lane, indicating that the bands observed were specific. This experiment provides further evidence that σ28 regulates cheOp2 expression.

FIG. 4.
Primer extension analysis of the promoter region of cheOp2 showing the absence of the σ28 regulated start site in the fliA mutant JPA467 and the σ28 and σ70 start sites in the wild-type strain WS8N. JPA455 is a negative control. ...

An electrophoretic mobility shift assay was performed to measure the interaction between σ28 and the proposed promoter. Purified σ28 was reconstituted with pure R. sphaeroides core RNA polymerase and bound to a biotin-labeled DNA fragment encompassing bases −91 to +85 relative to the transcriptional start. This was electrophoresed on a native polyacrylamide gel alongside free DNA. A second reaction was performed in which unlabeled competitor DNA was preincubated with the protein prior to the addition of labeled DNA. A bandshift was evident in the lane with both DNA and σ28 holoenzyme (data not shown), and the intensity of the shifted band was significantly reduced in the presence of competitor DNA, indicating that the σ28 holoenzyme is able to specifically bind the DNA fragment containing the σ28 recognition sequence.

cheOp3: identification of transcriptional start sites.

The region immediately upstream of cheA4 in cheOp3 (Fig. (Fig.5A)5A) was analyzed for known promoter recognition sequences. The 144 bp between cheA4 and the upstream orf is exceedingly GC-rich (82%), making it difficult to identify consensus promoter sequences. However, a GG at position −45 and a GC at position −33 (relative to the translational start of cheA4) were identified (Fig. (Fig.5B).5B). These specific bases and, importantly, the spacing between them are consistent with them representing the −24 and −12 recognition sequences of a σ54 regulated promoter (3). No putative σ70 recognition site was found.

FIG. 5.
(A) Diagram showing the architecture of cheOp3. Each gene is represented by large gray arrows and is labeled individually. The region contained within the lacZ construct pACM76 is indicated. (B) Diagram illustrating the promoter region of cheOp3. The ...

Multiple attempts to map the transcriptional start site upstream of the cheA4 of cheOp3 using mRNA isolated from WS8N failed. The 45 bp of DNA between the putative −24 GG promoter sequence and the translational start of cheA4 is 93% GC and is likely to have significant secondary structure.

cheOp3: analysis of mutants in the putative promoter regions.

An unmarked deletion mutant of the putative promoter region upstream of cheOp3 was constructed based on homology to known promoter recognition sequences and designated JPA457 (Fig. (Fig.5B).5B). The swarm diameters of this mutant JPA457 were comparable to the nonchemotactic strain JPA1301, suggesting that the region deleted contained the promoter sequences for cheOp3 (Fig. (Fig.6a).6a). The growth rates of all of the mutant strains were normal (data not shown).

FIG. 6.
(a) Comparison of the swarm diameters of the wild-type strain WS8N, the putative cheOp3 promoter mutant strain JPA457 and the nonchemotactic strain JPA1301 to 100 μM propionate under microaerobic conditions. The error bars represent the standard ...

Expression of CheB2 from cheOp3 in the deletion mutant JPA457 was compared to the wild type by immunoblotting with a specific antibody. The expression of CheB2 was abolished in JPA457 (Fig. (Fig.6b),6b), indicating a promoter between cheA4 and an orf (designated orfE) immediately upstream. It also excludes readthrough from any promoter upstream of orfE. These data suggest that the promoter sequences for cheOp3 expression have been deleted in the mutant strain.

cheOp3: RT-PCR.

To confirm the presence of a promoter in the region between orfE and cheA4, a RT-PCR was performed with an orfE Ω insertion mutant JPA466 in which all transcription is interrupted. Using primers that would amplify a short region of the reverse transcript of tlpT mRNA and a one-step reaction mix containing reverse transcriptase and a thermostable DNA polymerase, we assayed for the presence of tlpT cDNA. The presence of a band of the expected size indicated that a promoter downstream of the Ω insertion and immediately upstream of cheA4 initiates the transcription of cheOp3 genes (Fig. (Fig.7).7). A primer set specific for tlpC, the last gene in cheOp2 was used to control for mRNA quality and for the fidelity of the PCR. To control for contaminating DNA, PCRs were performed on RNA samples using the DNA polymerase alone. No bands were obtained in these controls.

FIG. 7.
RT-PCR using mRNA from the wild-type strain WS8N (lane 3) and the orfE omega insertion mutant JPA466 (lane 6) using primers to the reverse transcript of tlpT from cheOp3 showing the presence of a promoter immediately upstream of cheA4. The 298-bp band ...

cheOp3 is regulated by σ54 RpoN3.

During a random Tn mutagenesis screen, a nonchemotactic mutant (JPA1706) was identified. The DNA flanking the Tn insertion was sequenced and found to be identical to that of RpoN3. The expression of CheB1 encoded in cheOp2 and of CheW4 encoded in cheOp3 in this strain was compared to the wild type by immunoblotting with antibodies specific to these proteins (Fig. (Fig.8a).8a). Whereas the expression of CheB1 was not affected by the Tn insertion, the expression of CheW4 was abolished.

FIG. 8.
(a) Semiquantitative immunoblot of strain WS8N and the Tn mutant JPA1706 with an antibody specific to the cheOp3-encoded CheW4. The expression of CheW4 is abolished in the mutant strain. In comparison, a semiquantitative immunoblot with an antibody specific ...

To further investigate the role of RpoN3 in cheOp3 expression, the Smr promoterless lacZ translational fusion plasmids pACM76 and pACM77, which contained the 5′ of cheA4 and upstream sequences in the forward and reverse orientations, respectively, were introduced into the Tn mutant JPA1706. Expression of lacZ in these strains was compared to the wild type (Fig. (Fig.8b).8b). β-Galactosidase production was reduced to background levels in the Tn mutant. These data suggest that RpoN3 regulates cheOp3 expression. Moreover, taken together with the RT-PCR data, the analysis of the cheOp3 putative promoter mutant JPA457 and the identification of putative −24 and −12 promoter sequences, we suggest that RpoN3 recognizes sequences immediately upstream of cheA4.


R. sphaeroides contains two che loci that are essential for chemotaxis: cheOp2 and cheOp3 (6, 18). This study provides supportive evidence that expression of the operons is independently regulated by different sigma factors. In addition to a basal level of expression from a σ70-dependent promoter, we have shown that the expression of cheOp2 is also regulated by σ28, whereas cheOp3 is controlled by one of the four σ54 homologues in R. sphaeroides.

Flagellum synthesis is an energetically expensive process and the chemosensory machinery is worthless in the absence of a functional flagellum. The use of the σ28 alternative sigma factor and a cognate anti-sigma factor enables bacteria to coordinately regulate gene expression, such that the late flagellar genes and genes encoding chemotaxis proteins are not manufactured until the hook basal-body complex is in place (10). Consequently, flagellar synthesis is ordered and also directly coupled to chemotaxis. The expression of cheOp2 in R. sphaeroides is under dual control (Fig. (Fig.1B).1B). It seems that whereas the constitutive activity of the σ70 promoter ensures a basal level of cheOp2 mRNA, additional transcripts are initiated from the overlapping σ28 promoter. This may allow the coupling of chemosensing to flagellar synthesis and the growth cycle in R. sphaeroides. This is probably particularly important in a bacterium with a single flagellum that needs to coordinate the growth of a new flagellum with the development of the membrane-associated polarly localized chemosensory pathway expressed from cheOp2 (28), possibly ensuring the correct rate of expression for the current growth rate.

The sensory role of the cytoplasmic cluster is less clear. It seems probable that it responds to intracellular signals reflecting the current metabolic state, possibly integrating signals from the transmembrane chemoreceptors. The results presented here suggest that the expression of the cytoplasmic chemosensory cluster is regulated not by cell growth but by a specific σ54 that may respond to a cytoplasmic signal reflecting the metabolic state. R. sphaeroides is unusual in that it has four genes encoding σ54. Each of these controls specific regulons, and they are not apparently interchangeable (16). Published work has suggested that RpoN2 is involved in regulating flagellum synthesis (16). The present study suggests that RpoN3 regulates the expression of cheOp3. σ54 sigma factors are structurally and functionally distinct from all other sigma factors (29). They require the presence of an activator protein or an EBP that binds to an enhancer sequence located at least 100 bp upstream of the transcriptional start and catalyzes open complex formation through ATP hydrolysis. This requires looping of the DNA, a process often facilitated by an integration host factor, which binds DNA between the enhancer and the promoter. We have identified putative binding sites for an integration host factor approximately 50 bp upstream of the σ54 promoter sequence (Fig. (Fig.5B).5B). Interestingly, the gene coding the EBP for RpoN2, which regulates flagellar gene expression, is also located upstream of cheOp3. Why the flagellar and cytoplasmic chemosensory pathways need to be regulated by different σ54 homologues is interesting and suggests the regulation of CheOp3 is independent of the flagellum but linked to the intracellular metabolic state.

Alternative sigma factors permit control of the expression of proteins that are either not required at the same time or in the same amount. Such regulation enables the bacterium to adapt rapidly to changes in its environment. The independent regulation of cheOp2 and cheOp3 is consistent with the hypothesis of a distinct, if overlapping, role for each che pathway.


This study was supported by the BBSRC.

We thank T. J. Donohue for supplying core RNA polymerase from R. sphaeroides, O. Isathitphaisarn for technical assistance, and Scott Godfrey for help with the transposon screen.


[down-pointing small open triangle]Published ahead of print on 8 September 2006.


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