The Master Regulators of the Fla1 and Fla2 Flagella of Rhodobacter sphaeroides Control the Expression of Their Cognate CheY Proteins

ABSTRACT

Rhodobacter sphaeroides is an alphaproteobacterium that has two complete sets of flagellar genes. The fla1 set was acquired by horizontal transfer from an ancestral gammaproteobacterium and is the only set of flagellar genes that is expressed during growth under standard laboratory conditions. The products of these genes assemble a single, subpolar flagellum. In the absence of the Fla1 flagellum, a gain-of-function mutation in the histidine kinase CckA turns on the expression of the fla2 flagellar genes through the response regulator CtrA. The rotation of the Fla1 and Fla2 flagella is controlled by different sets of chemotaxis proteins. Here, we show that the expression of the chemotaxis proteins that control Fla2, along with the expression of the fla2 genes, is coordinated by CtrA, whereas the expression of the chemotaxis genes that control Fla1 is mediated by the master regulators of the Fla1 system. The coordinated expression of the chemosensory proteins with their cognate flagellar genes highlights the relevance of integrating the expression of the horizontally acquired fla1 genes with a chemosensory system independently of the regulatory proteins responsible for the expression of fla2 and its cognate chemosensory system.

IMPORTANCE Gene acquisition via horizontal transfer represents a challenge to the recipient organism to adjust its metabolic and genetic networks to incorporate the new material in a way that represents an adaptive advantage. In the case of Rhodobacter sphaeroides, a complete set of flagellar genes was acquired and successfully coordinated with the native flagellar system. Here we show that the expression of the chemosensory proteins that control flagellar rotation is dependent on the master regulators of their corresponding flagellar system, minimizing the use of transcription factors required to express the native and horizontally acquired genes along with their chemotaxis proteins.

INTRODUCTION

Chemotaxis is a complex response that enables bacteria to detect and swim toward or away from positive or negative stimuli in the surrounding environment. To achieve chemotaxis, a signal transduction system senses and controls the flagellar motor. Signals are sensed by specialized receptors that are assembled together with the chemotaxis proteins CheW and CheA to form complex arrays. In response to receptor occupancy, CheA phosphorylates the response regulator CheY, which in turn binds to the flagellar switch complex to control flagellar rotation. The response is terminated by lowering the levels of CheY-P, through the action of the phosphatase CheZ. The chemotaxis proteins CheB and CheR play a key role in adaptation by chemically modifying the chemotaxis receptors. The methylesterase CheB is stimulated upon phosphorylation by CheA, and CheR is a constitutive methyltransferase. The activity of these proteins resets the signaling state of the receptor (for reviews, see references 1–5).

This basic chemotaxis system is found in many species other than Escherichia coli and Salmonella enterica. However, many species have multiple copies of the chemotaxis genes. The products of these additional copies frequently are involved in chemotaxis, but in some species these copies have evolved to control other cellular functions such as development, biofilm formation, and twitching motility mediated by type IV pili (6, 7).

In several bacterial species, the chemotaxis genes are expressed coordinately with the flagellar genes. Therefore, the cytoplasmic components that control flagellar rotation are present only when the flagellum is formed. In the paradigmatic bacteria E. coli and S. enterica, the chemotaxis genes are expressed when the cytoplasmic and membrane components of the flagella are already formed (8–10). A similar situation has been observed in Pseudomonas aeruginosa and Xanthomonas campestris (11, 12). In the alphaproteobacteria Caulobacter crescentus and Sinorhizobium meliloti, chemotaxis genes are expressed under the control of the master activator protein of the flagellar genes (13, 14). Therefore, the chemotaxis genes are expressed early during flagellar biogenesis.

Rhodobacter sphaeroides is an alphaproteobacterium with two different flagellar systems, multiple copies of the chemotaxis genes, and four different rpoN genes, encoding functionally different σ⁵⁴ factors. σ⁵⁴-2 has been shown to be required to transcribe one of the flagellar sets (15–17).

This bacterium assembles a single, subpolar flagellum under standard laboratory growth conditions (18), and the products of the fla1 genes are responsible for the formation and functioning of this structure. The expression of these genes follows a hierarchical pattern. At the top of the hierarchy, the master protein FleQ, together with the RNA polymerase core (E) associated with the σ⁵⁴-2 factor, promotes the expression of the class II flagellar operon fleT-fliEFGHI. Subsequently, the hetero-oligomeric complex FleQ/FleT and Eσ⁵⁴-2 bring about the transcription of genes encoding the proteins that form the hook and the basal body (HBB), fliA (which encodes σ²⁸) and flgM (encoding the anti-sigma factor of σ²⁸). Once the HBB has been assembled, FlgM is exported out of the cell, and σ²⁸ is free to associate with E to transcribe the late flagellar genes, such as those encoding flagellin (FliC) and its scaffold protein, FliD (19).

The other flagellar system present in R. sphaeroides, Fla2, enables the formation of several polar flagella (16, 20). The fla2 genes are not expressed under standard laboratory growth conditions. So far, the Fla2 flagellar system has been studied in mutant strains that are unable to form Fla1 due to a mutation in a regulatory or structural gene and importantly contain a gain-of-function mutation in the histidine kinase CckA. In these strains, it has been determined that the expression of fla2 is dependent on the two-component system formed by the histidine kinase CckA, the phosphotransferase protein ChpT, and the response regulator CtrA (21).

Phylogenetic studies have shown that the fla2 genes constitute the native flagellar genes of R. sphaeroides, whereas the fla1 set was acquired by horizontal transfer from an ancestral gammaproteobacterium (16). In R. sphaeroides, the multiple copies of the chemotaxis genes are organized in four different operons (15) (Fig. 1).

FIG 1.

Schematic representation of the organization of the chemotaxis genes in R. sphaeroides. The multiple cheY genes are in red. Black outlines group the chemotaxis operons that control Fla1 or Fla2.

The control of Fla1 was shown to be mediated by the products of che operon 2 (cheOp2) and cheOp3 (15, 22–24), whereas the control of Fla2 is mediated by the chemosensory proteins encoded in cheOp1 (25, 26). Specifically, the chemotaxis response dependent on Fla1 is mainly controlled by CheY6 (encoded in cheOp3), as well as either CheY3 (encoded in cheOp2) or CheY4 (24). Fla2 is controlled by CheY2 and CheY5 (both encoded by cheOp1) (25).

Regarding the expression of these genes, cheOp3 expression reportedly depends on the σ⁵⁴-3 factor (27). This is unexpected, given that the expression of flagellar and chemotaxis genes in bacteria is usually coordinated and that σ⁵⁴-3 has not been implicated in the transcription of any fla1 gene. The expression of cheOp1 has yet to be studied. The other chemotaxis operon whose expression has already been analyzed is cheOp2, which is dependent on σ⁷⁰ and σ²⁸ (FliA) (27). Therefore, whether cheY4 and cheOp1 are expressed concurrently with fla1 and fla2, respectively, remains to be determined.

Another important aspect that remains to be determined is the identity of the activator protein that is responsible for interacting with Eσ⁵⁴-3 to promote cheOp3 expression; since Eσ⁵⁴ is unable to form open complex by itself, the presence of an activator protein of the bacterial enhancer-binding protein (bEBP) family is compulsory (28). In this context, it should be stressed that most bacteria have a single σ⁵⁴ factor (encoded by rpoN) that interacts with multiple bEBPs that recognize specific binding sites to achieve the transcription of a specific set of genes (29). In contrast, R. sphaeroides has four σ⁵⁴ factors, two of which, σ⁵⁴-1 and σ⁵⁴-2, are activated by specific bEBPs (i.e., NifA and FleQ/FleT, respectively) (17, 30). Potential bEBP-encoding genes were previously identified in silico in the genome of R. sphaeroides, revealing that aside from the bEBPs for σ⁵⁴-1 and σ⁵⁴-2, only two loci remain that encode proteins with bEBP family signatures that could be the activators for σ⁵⁴-3 and σ⁵⁴-4 (19).

In this work, we have determined that the expression of cheOp3 is dependent on σ⁵⁴-2 and not on σ⁵⁴-3, as was previously reported (27), and that the bEBP of the fla1 system (FleQ) is the activator protein for this operon. In addition, we found out that cheY4 expression is dependent on σ²⁸ (FliA) and, consequently, is also a part of the flagellar transcription hierarchy. Our results also show that cheOp1 expression is controlled by CtrA. Together, these results establish that the expression of the chemotaxis genes in R. sphaeroides is coordinated with the expression of their cognate flagellar system.

RESULTS

In this work, we used derivatives of the WS8N and AM1 strains to study the regulation of the flagellar and chemotaxis genes. WS8N is the wild-type strain that constitutively expresses the Fla1 flagellum but is unable to express the fla2 genes given that the two-component system CckA/ChpT/CtrA is inactive. The AM1 strain is a WS8N derivative that is unable to express the fla1 genes, given that it carries a mutation in the flagellar master activator fleQ and also carries a point mutation in cckA that activates the CckA/ChpT/CtrA two-component system (21, 25); therefore, this strain has a Fla1⁻ Fla2⁺ phenotype.

cheOp1 expression is dependent on the master regulator of the fla2 genes.

The expression of three chemotaxis proteins encoded in cheOp1 (Fig. 1) was evaluated by Western blotting using total cell extracts obtained from AM1 (25) and its derivative, EA1 (ΔctrA::aadA) (21). The proteins CheY2 and CheY5 were detected in AM1 but not in the absence of ctrA (Fig. 2A), indicating that cheOp1 expression is dependent on the presence of CtrA. To test for another protein encoded in cheOp1, we isolated a chromosomal fusion of mcpA with the Flag epitope in the AM1 strain. The resulting strain, JV1 (mcpA-Flag) and its derivative, JV2 (mcpA-Flag ΔctrA::hyg), were tested by immunoblotting using an anti-Flag antibody (Fig. 2A). Consistent with the hypothesis that CtrA is required to transcribe cheOp1, McpA-Flag was detected in total cell extracts of JV1 but not of JV2.

FIG 2.

Determination of the presence of proteins encoded in cheOp1 by immunoblotting and measuring the transcriptional activity of cheOp1 using a reporter gene. (A) Total cell extracts were obtained from cultures grown in Sistrom's minimal medium supplemented with 80 μM succinic acid and harvested at an OD₆₀₀ of 0.3. The notation at the top of the Western blots regarding the status of the cckA allele indicates that the WS8N and ΔfleQ (SP13) strains do not carry the gain-of-function mutation in CckA that enables the expression of the fla2 genes whereas the AM1 strain and its derivatives carry the mutations ΔfleQ and, importantly, cckA_L391F, which activate the CckA/ChpT/CtrA two-component system. Each lane is labeled according to the relevant mutation present in the strain. The antibody used for each blot is indicated at the top. MW indicates the lane where the molecular weight marker was loaded. Anti-CheY2 and anti-CheY5 antibodies were used at a 1:3,000 dilution, and anti-Flag antibody (Sigma-Aldrich) was used at a 1:10,000 dilution. (B) β-Glucuronidase activity promoted by the chromosomal fusion mcpB::uidA-aadA. AM1 and its ΔctrA derivative (LC7 strain) carrying the transcriptional fusion mcpB::uidA-aadA were grown in 80 μM succinic acid, and total cell extracts were used for the enzymatic reaction. Activity is reported in picomoles of 4-methylumbelliferone produced per minute per milligram of protein. The mean values and standard deviations of the results of three independent determinations are shown.

To rule out the possibility that an undetected mutation other than ΔctrA was responsible for the lack of CheY2, CheY5, and McpA-Flag, we introduced a plasmid that expresses CtrA (pRK_ctrA) into the EA1 and JV2 strains. In these complemented strains, the expression of CheY2, CheY5, and McpA-Flag was restored (see Fig. S1 in the supplemental material).

Consistent with the proposed model of activation of the CckA/ChpT/CtrA system (21), we did not observe the presence of either CheY2 or CheY5 in the wild-type strain WS8N or in the SP13 (ΔfleQ::kan) mutant (Fig. 2A), indicating that cheOp1 is expressed only when the CckA/ChpT/CtrA two-component system is active. To test the hypothesis that cheOp1 expression could be transcriptionally controlled by CtrA, a transcriptional fusion of mcpB to the promoterless uidA gene was made. This fusion was used to replace the mcpB⁺ gene in AM1 and ΔctrA::hyg (LC7) strains. uidA expression was recorded as the amount of β-glucuronidase present in total cell extracts. We noticed that the enzymatic activity was severely reduced in the absence of CtrA (Fig. 2B), implying that transcription of cheOp1 is directly or indirectly controlled by CtrA.

The control region of cheOp1 has a potential CtrA binding site relevant for its expression.

Given that the expression of several of the proteins encoded in cheOp1 was dependent on CtrA, we searched for potential CtrA binding sites upstream of the annotated genes in chromosome 1 using a CtrA position weight matrix (PWM) built from the regulatory regions of 54 genes identified to be controlled by CtrA in Caulobacter crescentus (31). From this analysis, the sequence TTAGN₇TTAT was found 67 bp upstream of the translational start site of RSWS8N_02465, the gene located upstream of cheY5 (Fig. 1 and 3A). The identified sequence is highly similar to the CtrA consensus binding sequence (TTAAN₇TTAA) previously characterized in C. crescentus and other species (32), suggesting that cheOp1 transcription may be activated by CtrA from this site. This finding implies that RSWS8N_02465 might be part of cheOp1. To evaluate this prediction, we performed a reverse transcription (RT)-PCR experiment using specific primers for RSWS8N_02465 and cheY5. Using total RNA from AM1, the main product amplified from this reaction had the expected size for a DNA fragment that expands from RSWS8N_02465 to cheY5 (Fig. 3B). Further supporting the idea that RSWS8N_02465 is part of cheOp1, it should be noted that the stop codon of RSWS8N_02465 overlaps the start codon of cheY5. RSWS8N_02465 encodes a 109-amino-acid protein with a STAS (sulfate transporter and anti-sigma factor antagonist) domain that has not been implicated in chemotaxis so far.

FIG 3.

Nucleotide sequence upstream of cheOp1, activity of the reporter gene uidA fused to different fragments from the regulatory region of cheOp1 in the pBBMCS53 plasmid, and RT-PCR data for cheY5 and its upstream gene. (A) Nucleotide sequence upstream of RSWS8N_02465. The first residues of the protein encoded by this gene are shown below the corresponding codons. The putative Shine-Dalgarno sequence is underlined, and nucleotides similar to the consensus binding site of CtrA, identified using a previously reported position weight matrix (31), are shown in bold and capital letters. The LOGO sequence of the CtrA-binding site is shown below; it was reported as part of a genomic analysis performed in C. crescentus (32). Asterisks above the sequence indicate nucleotides that match with the most frequent base in the LOGO sequence. An arrow indicates the 5′ end of the fragment cloned into pBBMCS53_cheOp1S. The 5′ end of the fragment cloned into pBBMCS53_cheOp1L includes 25 bp upstream of the sequence shown. (B) Agarose gel showing the RT-PCR products of the primer pair that amplifies the region encompassing CheY5 and RSWS8_02465. A 469-bp product was expected for this reaction. Lane 1, 5 μl from the RT-PCR; lane 2, 5 μl from the control reaction carried out in the absence of reverse transcriptase. The relevant molecular sizes from a 100-bp DNA ladder are shown on the left. (C) β-Glucuronidase activity of cell extracts from the AM1 and EA1 (ΔctrA::aadA) strains carrying the indicated plasmids. Cell cultures were grown in Sistrom's minimal medium supplemented with 80 μM succinic acid. Activity is reported in picomoles of 4-methylumbelliferone produced per minute per milligram of protein. The mean values and standard deviations (SD) of the results of three independent determinations are shown.

To evaluate if the upstream region of RSWS8N_02465 exhibits CtrA-dependent promoter activity, we cloned this region upstream of the reporter gene uidA in the pBBMCS53 plasmid. The resulting construct (pBBMCS53_cheOp1L) promoted high β-glucuronidase expression in the AM1 strain but not in EA1 (ΔctrA::aadA) (Fig. 3C). In addition, we cloned a shorter fragment from the upstream region of RSWS8N_02465, which excludes the putative CtrA binding site, into the pBBMCS53 plasmid. The resulting plasmid (pBBMCS53_cheOp1S) did not promote β-glucuronidase expression in either of the two strains (Fig. 3C). The lack of expression of the reporter gene in this construction could be explained by the loss of either the CtrA binding site or the −35 promoter region. In this context, it should be stressed that most of the CtrA-activated promoters in C. crescentus show overlapping of these two sequence motifs (32, 33).

cheOp3 expression is dependent on σ⁵⁴-2 and the master regulator of the fla1 genes.

CheY6 and either CheY3 or CheY4 are needed to control the chemotaxis response of the Fla1 flagellum (24, 34). CheY6 is encoded in cheOp3, which is located within the 56-kb fla1 gene cluster, and CheY3 is encoded in cheOp2, located elsewhere on chromosome I, whereas CheY4 is located on chromosome II (34) (Fig. 1).

cheOp3 expression was previously shown to depend on σ⁵⁴-3 (27); however, fla1 gene expression is dependent on σ⁵⁴-2, raising the possibility that the expression of cheOp3 is unlinked from the expression of Fla1. Given that Eσ⁵⁴ is unable to form open complex by itself, we decided to identify the bEBP responsible for activating Eσ⁵⁴-3. The in silico identification of bEBP-encoding genes in the R. sphaeroides genome revealed only five genes that encode proteins with the characteristic signatures of the σ⁵⁴ transcriptional activators (19): nifA, fleQ, fleT, RSWS8N_RS13335, and RSWS8N_RS04480. We previously reported the isolation of strains with mutations in the genes nifA, fleQ, and fleT (19, 30); therefore, we isolated strains with mutations in RSWS8N_RS13335 and RSWS8N_RS04480.

The expression of CheY6 (encoded in cheOp3) was tested in total cell extracts obtained from the wild-type strain, along with cell extracts from the strains with mutations in each of the five bEBP-encoding genes. As shown in Fig. 4A, CheY6 expression was abolished in the strain carrying the ΔfleQ::kan allele and was not affected by the absence of RSWS8N_RS13335, RSWS8N_RS04480, or nifA, indicating that CheY6 is expressed as a part of the fla1 transcriptional hierarchy.

FIG 4.

Determination of the presence of CheY6 in different strains, activity of the reporter gene uidA fused to the regulatory region of cheOp3 in the pBBMCS53 plasmid, and RT-PCR of cheY6 and its upstream gene. Total cell extracts obtained from cell cultures grown in Sistrom's minimal medium and harvested at an OD₆₀₀ of 0.6 were tested as described in Materials and Methods. (A) Total cell extracts were separated by gel electrophoresis, transferred to nitrocellulose, and probed with anti-CheY6 antibodies (1:3,000). WT, WS8N cell extract. Each lane is labeled according to the relevant mutation present in the strain. Square brackets group the samples according to their function. Fla-EBPs indicate the two enhancer binding proteins required for activation of the fla1 genes, rpoN indicates the genes encoding σ⁵⁴-1, -2, -3, or -4, and EBPs indicate the genes encoding proteins that belong to the enhancer binding protein family in the R. sphaeroides genome sequence. The signal identified as CheY was quantified by densitometry using ImageJ; the resulting value was normalized using an unspecific band on the blot. The values shown in parentheses are the average results of three independent experiments and are expressed as a percentage of the WT level. Figure S3 shows the graph of these values with SD. (B) β-Glucuronidase activity of the wild-type and mutant strains carrying the plasmid pBBMCS53_cheA4. Activity is reported in picomoles of 4-methylumbelliferone produced per minute per milligram of protein. The mean values and standard deviations of the results of three independent determinations are shown. (C) Agarose gel of the RT-PCR products of the primer pair that amplifies the region encompassing cheY6 and its upstream gene tlpT. A 391-bp product was expected for these primers. Lane 1, 5 μl from the RT-PCR; lane 2, 5 μl from the control reaction carried out in the absence of reverse transcriptase. The relevant sizes in base pairs of the DNA molecular size marker X (Roche Life Science) are shown on the left.

An approximately 60% reduction in CheY6 levels was noticed in the absence of FleT and FliA. In both of these strains, the FleQ activator is present and the σ²⁸ (FliA) factor is absent, suggesting that wild-type CheY6 levels could arise from the combined activity of a class I FleQ-activated promoter and a class IV σ²⁸-dependent promoter.

In contrast to the proposal that cheOp3 expression is dependent on σ⁵⁴-3 (27), we observed that the expression of CheY6 (encoded in cheOp3) was dependent only on σ⁵⁴-2 and that the absence of σ⁵⁴-3 did not affect CheY6 expression levels (Fig. 4A). Given that cheY6 and the upstream gene tlpT are located 204 bp apart, cheY6 may be transcribed independently of the upstream chemotaxis genes (Fig. 1). Therefore, to determine if the expression of the genes located upstream of cheY6 was also dependent on FleQ and σ⁵⁴-2, we tested the expression of a reporter gene under the control of the region located upstream of cheA4 which has been reported to carry the cheOp3 promoter (27). To this end, an 854-bp fragment of the upstream region of cheA4, starting from codon 48 of cheA4, was cloned upstream of the reporter gene uidA in pBBMCS53 (35). The resulting plasmid was introduced into the WS8N, ΔrpoN2::kan, and ΔfleQ::kan strains. As a control, this plasmid was also introduced into the ΔrpoN3 strain.

Total cell extracts of these strains were assayed for β-glucuronidase activity. A strong reduction in the amount of β-glucuronidase was detected for the ΔrpoN2::kan and ΔfleQ::kan mutants compared with that in the wild-type WS8N or ΔrpoN3::kan strains (Fig. 4B), suggesting that the expression of cheA4 is also dependent on σ⁵⁴-2 and FleQ. In addition, RT-PCR revealed that cheY6 is cotranscribed with tlpT, suggesting that these genes could form a single operon (Fig. 4C).

σ²⁸ is involved in the expression of cheY4.

The expression of CheY3 and CheY4 was analyzed in each of the mutant strains lacking either one of the bEBP or σ⁵⁴ factors. We observed that the expression of CheY4 was severely reduced in ΔfleQ::kan (SP13), ΔfleT::aadA (SP12), ΔrpoN2::kan (SP7), and ΔfliA::kan (SP15) strains (Fig. 5A). Since it has been shown that the expression of fliA is dependent on FleQ, FleT, and σ⁵⁴-2 (19), these results suggest that cheY4 expression is dependent on σ²⁸ (fliA) and that the absence of CheY4 in the ΔfleQ, ΔfleT, and ΔrpoN2 strains is the expected consequence of this regulation.

FIG 5.

Determination of the presence of CheY4 in different mutant strains by immunoblotting. Total cell extracts obtained from cell cultures grown in Sistrom's minimal medium and harvested at an OD₆₀₀ of 0.6 were tested as described in Materials and Methods. Anti-CheY4 antibodies were used at 1:3,000. WT denotes the WS8N cell extract. Each lane is labeled according to the relevant mutation present in the strain. Other labels are the same as described in the legend for Fig. 4A. (B) Acrylamide gel electrophoresis of the RT-PCR products corresponding to the primer pair that amplifies cheY4 (lanes 1 to 4) and the primer pair that amplifies flgH (lanes 5 and 6). Lanes: 1, 3, and 5, total RNA from WS8N; 2, 4, and 6, total RNA from the fliA::kan mutant strain. Control reactions were carried out in the absence of reverse transcriptase (lanes 3 and 4). Lane MW corresponds to PhiX174 DNA digested with HaeIII and was used as a molecular size marker (fragment sizes, 1,353, 1,078, 872, 603, 310, 281, 271, 234, 194, and 118 bp). The expected sizes for the cheY4 and flgH products are 361 and 639 bp, respectively).

To verify that the expression of cheY4 was dependent on the presence of σ²⁸, we carried out an RT-PCR assay using specific oligonucleotides for cheY4. The expected product of 361 bp was detected when the total RNA from WS8N was included in the reaction, whereas a faint product was detected when RNA from the fliA::kan strain was used (Fig. 5B, lanes 1 and 2, respectively). It should be noted that for this RNA, a similar signal was detected when no reverse transcriptase was included in the reaction mixture (Fig. 5B, lane 4), indicating that this slight signal is the product of a small amount of DNA in the sample. To be certain that both samples of RNA had the appropriate quality to yield an RT-PCR product, we performed a reaction using a pair of oligonucleotides that amplified a region of the flgH gene (Fig. 5B, lanes 5 and 6).

Given that our results suggest that CheY4 is expressed from a σ²⁸-dependent promoter, we searched upstream of cheY4 for a sequence similar to the σ²⁸ consensus promoter. We analyzed a 159-bp region from the stop codon of RSWS8N_17744 (RSP_3301) to the start codon of cheY4 by using a PWM built from 24 σ²⁸-dependent promoters (36). This analysis revealed a possible σ²⁸-promoter located 104 bp upstream of the start codon of cheY4 (Fig. S2). The sequence of this putative promoter is similar to that of previously reported σ²⁸-dependent promoters in R. sphaeroides (19).

A more complex situation was observed for CheY3, where we detected a reduction in the amount of CheY3 in the ΔfliA mutant strain but no reduction in the absence of FleQ or FleT (Fig. 6A); this result was unexpected given that the expression of fliA depends on FleQ and FleT (19). It has been shown that cheOp2 contains a regulatory region with two promoters, one dependent on σ⁷⁰ and the other dependent on σ²⁸ (27), which explains the reduction of CheY3 in the absence of σ²⁸ (FliA). However, the CheY3 expression level in the absence of FleQ and FleT (in these strains, fliA is not expressed [19]) is similar to that observed in the wild-type strain, suggesting that FleQ and FleT may have a negative effect on CheY3 expression.

FIG 6.

Determination of the presence of CheY3 in different strains by immunoblotting and plot profile of the primer extension experiment for cheOp2. (A) Total cell extracts obtained from cell cultures grown in Sistrom's minimal medium and harvested at an OD₆₀₀ of 0.6 were tested as described in Materials and Methods. Anti-CheY3 antibodies were used at 1:3,000. WT, WS8N cell extract. Each lane is labeled according to the relevant mutation present in the strain. Other labels are the same as described in the legend for Fig. 4A. (B) Total RNA was isolated from cell cultures of WS8N and SP13 (ΔfleQ::kan) strains grown in Sistrom's minimal medium. After hybridization of the RNA samples with the 5′-end-labeled oligonucleotide, cDNA synthesis was performed using AMV reverse transcriptase. The reactions were subjected to gel electrophoresis, and the images were visualized using a Typhoon scanner (GE Healthcare Life Science) and quantified using ImageJ. The plot profiles initiate immediately below the largest transcript detected (position −155 from the translational start site of RSWS8N_13355), denoted by an open circle. The transcripts that originated from the σ⁷⁰ and σ²⁸ promoters are indicated by vertical arrows. The asterisks correspond to the migration of the 151-, 140-, 118-, 100-, and 82-bp (from left to right) DNA fragments of PhiX174 that were obtained after digestion with HinfI (Promega) and 5′ end labeled.

To evaluate this possibility, we carried out a primer extension experiment using the oligonucleotide cheOp2B, which targets the first gene of cheOp2 (i.e., RSWS8N_13355). The reaction mixtures were subjected to gel electrophoresis, and the products were analyzed using ImageJ. The plot profile of the sample from the wild-type strain revealed the previously reported σ²⁸ and σ⁷⁰ promoters (27) (Fig. 6B). From two independent experiments, a 26% average increase was observed in the amount of the σ⁷⁰-dependent transcript in the ΔfleQ mutant strain compared with that in the wild-type strain. This increase is consistent with a negative role for FleQ on cheOp2 expression. Unexpectedly, we detected a low-abundance transcript in both strains that initiates approximately 155 nt upstream of the translational start of RSWS8N_13355. The amounts of transcript are similar for the two samples.

DISCUSSION

Our work shows that the expression of chemotaxis proteins in R. sphaeroides is coordinated with the expression of the cognate flagellar system. In many bacteria, chemotaxis genes are expressed under the control of the flagellar regulator proteins, and in E. coli and other related bacteria, their expression is dependent on σ²⁸ (FliA), indicating that these genes belong to the lower class of the flagellar hierarchy. In contrast, chemotaxis genes in other bacteria are expressed under the control of the master regulator of the flagellar hierarchy; therefore, the chemotaxis proteins are synthesized during the early steps of flagellar formation. These opposing patterns suggest that the timing of chemotaxis protein expression may be irrelevant for the function of the system or that this regulation may reflect different regulatory needs. Nonetheless, the coordinated expression of flagellar proteins and chemotaxis proteins seems to be a ubiquitous scenario, and this scheme prevails in the case of R. sphaeroides, even though one flagellar system was horizontally acquired, indicating a strong pressure to coordinate the presence of the flagellar and chemotaxis proteins. In agreement with this view, it has been proposed that the gene encoding σ⁵⁴-2 (rpoN2) is a duplication from the vertically inherited rpoN1 gene (37).

In R. sphaeroides, the expression of the fla2 genes is dependent on the transcriptional activator CtrA (21). We demonstrated that cheOp1 expression is also dependent on CtrA. Our data identified a 652-bp fragment that contains the cheOp1 promoter and revealed the possible site through which CtrA mediates transcriptional activation. Our results also showed that RSWS8N_02465 is part of cheOp1; the product of this gene has yet to be implicated in chemotaxis, but its presence in cheOp1 suggests its potential involvement in this response. Interestingly, a gene containing a STAS domain is also located upstream of cheY3 in cheOp3, strengthening the idea that the products of these genes might be involved in chemotaxis. In addition, our results show that the cheOp1, CheY5, CheY2, and McpA gene products are not expressed when the CckA/ChpT/CtrA two-component system is inactive (in the WS8N strain or its derivative, SP13 [ΔfleQ::kan]), further supporting the notion that cheOp1 is expressed only when the fla2 genes are expressed.

In contrast to the Fla2 flagella, where the expression of all the CheY proteins involved in its control are dependent on the master flagellar regulator CtrA, the expression of the chemotaxis proteins that control Fla1 appears to be more complex. The expression of cheOp3, in which CheY6 is encoded, was previously suggested to depend on the sigma factor σ⁵⁴-3 (27); however, our results show that cheOp3 expression is dependent on σ⁵⁴-2 and the master regulator FleQ. We speculate that a mistake in the nomenclature of the σ⁵⁴ factors might be behind this disparity. The relevant point that should be stressed is that the expression of cheOp3, which encodes CheY6, the only CheY able to bind the Fla1 flagellar motor, is controlled by the master flagellar regulator FleQ and the σ⁵⁴-2 factor.

We also observed a reduction in the amount of CheY6 in cell extracts from the ΔfleT::aadA (SP12) and ΔfliA::kan (SP15) strains (Fig. 4A), suggesting that cheOp3 should have an additional σ²⁸-dependent promoter and that cheOp3 might be expressed from both a FleQ-dependent promoter and from a σ²⁸ (FliA)-dependent promoter. If this hypothesis is correct, the observed reduction of CheY6 in the absence of FleT and FliA can be explained by the absence of σ²⁸ in these two strains, leaving only the FleQ promoter active. The fact that CheY6 expression depends on the master flagellar regulator FleQ and probably also on σ²⁸ (FliA) highlights the importance of keeping a high and sustained level of this protein as long as the Fla1 flagellum is present, consistent with the idea that CheY6 is the main chemotaxis regulator of Fla1 (34).

The response regulators CheY3 and CheY4, which act along with CheY6 to control Fla1, are also expressed through the action of the transcriptional factors that regulate the Fla1 gene hierarchy. We observed that CheY4 expression is dependent on σ²⁸ (FliA), and consequently we did not detect this protein in the absence of either σ²⁸ or the FleQ, FleT, and σ⁵⁴-2 proteins, which are required to transcribe fliA (19). Accordingly, no RT-PCR product of cheY4 was obtained from RNA of the cheY4::aadA strain, and a sequence similar to the σ²⁸ consensus promoter was identified upstream of cheY4.

The expression of CheY3, which is encoded in cheOp2, depends on σ⁷⁰ and σ²⁸, and a part of its expression is dependent on the σ²⁸ promoter (27). Our results also showed a reduction in the level of CheY3 in the ΔfliA mutant strain. However, the lack of a reduction in the ΔfleQ and ΔfleT strains suggests that these proteins have a negative effect on CheY3 expression. A primer extension experiment designed to detect expression by the two cheOp2 promoters showed a 26% increase in the amount of the σ⁷⁰-dependent transcript in the absence of FleQ, which could explain the Western blot results. The mechanism of this negative regulation remains to be determined, but the sustained expression of cheOp2 mediated by σ⁷⁰, as well as its possible increase in the absence of FleQ, makes this operon unique among the four chemotaxis operons, suggesting that the products of cheOp2 might be required to accomplish another physiological role.

Figure 7 presents a model summarizing the conclusions of this work. This model illustrates the transcriptional relationship between the expression of flagellar genes and their cognate chemosensory proteins. Notably, only one transcriptional regulator directly controls the expression of the proteins involved in the chemotaxis response of Fla2, whereas the chemosensory proteins that control Fla1 are directly or indirectly regulated by two different transcription factors, FleQ and σ²⁸. Whether there is a physiological reason for this multiple regulation remains to be elucidated, but obviously, a pathway controlled by several regulators offers multiple checkpoints for finely tuned responses. Nevertheless, it should be kept in mind that this global regulation may simply reflect the evolutionary history of these genes.

FIG 7.

Model for the transcriptional control of genes encoding the CheY proteins of R. sphaeroides through the master regulators of fla1 and fla2. Solid arrows represent the confirmed regulatory pathways established here and in reference 27. Broken lines represent the regulation inferred from our results. The brackets at the bottom indicate the chemotaxis operon to which each cheY gene belongs.

MATERIALS AND METHODS

Genome sequences, plasmids, bacterial strains, and growth conditions.

All plasmids and bacterial strains used in this work are listed in Table 1. The genome sequence analyzed in this work corresponds to R. sphaeroides WS8N (NCBI reference sequence NZ_CM001161.1, GI:332561612). R. sphaeroides WS8N (38) and its derivatives were grown chemoheterotrophically at 30°C in Sistrom's minimal medium (39). AM1 and its derivatives were grown photoheterotrophically in Sistrom's minimal medium with 80 μM succinic acid as the carbon source. Photoheterotrophic liquid cultures were grown under continuous illumination and in completely filled screw-cap tubes. Heterotrophic liquid cultures were incubated in the dark with orbital shaking at 200 rpm. Escherichia coli was grown in Luria-Bertani (LB) medium (40) at 37°C. When required, antibiotics were added at the indicated concentrations: for R. sphaeroides, kanamycin (25 μg/ml), tetracycline (1 μg/ml), spectinomycin (50 μg/ml), and hygromycin (20 μg/ml for liquid cultures and 150 μg/ml for plates); for E. coli, kanamycin (50 μg/ml), spectinomycin (50 μg/ml), ampicillin (100 μg/ml), and hygromycin (20 μg/ml for liquid cultures and 200 μg/ml for plates).

TABLE 1.

Strains and plasmids used in this study

Strain or plasmid	Description	Reference or source
Rhodobacter sphaeroides strains
AE1	AM1 derivative; ΔfleQ::kan cckAL391F ΔctrA::aadA	21
AM1	WS8N derivative; ΔfleQ::kan cckAL391F	25
JV1	AM1 derivative; ΔfleQ::kan cckAL391F mcpA-Flag	This work
JV2	JV1 derivative but ΔctrA::hyg	This work
JV3	AM1 derivative; mcpB::uidA-aadA	This work
JV4	AM1 derivative; mcpB::uidA-aadA ΔctrA::hyg	This work
LC7	AM1 derivative; ΔctrA::hyg	This work
RS1Y3	WS8N derivative; cheY3::aadA	25
RS1Y4	WS8N derivative; ΔcheY4::aadA	25
RS1Y6	WS8N derivative; ΔcheY6::aadA	25
RS2Y1	AM1 derivative; cheY1::aadA	25
RS2Y2	AM1 derivative; cheY2::aadA	25
RS2Y5	AM1 derivative; cheY5::aadA	25
SP7	WS8N derivative; ΔrpoN2::kan rpoN2 (RSWS8N_05555)	17
SP8	WS8N derivative; ΔrpoN1::aadA rpoN1 (RSWS8N_07950)	17
SP12	WS8N derivative; ΔfleT::aadA	19
SP13	WS8N derivative; ΔfleQ::kan	19
SP15	WS8N derivative; ΔfliA::kan	19
SP16	WS8N derivative; ΔnifA::kan	30
SP21	WS8N derivative; ΔrpoN3::kan rpoN3 (RSWS8N_19584)	17
SP22	WS8N derivative; ΔrpoN4::kan rpoN4 (RSWS8N_04590)	17
SP23	WS8N derivative; ΔRSWS8N_RS13335::hyg	This work
SP24	WS8N derivative; ΔRSWS8N_RS04480::kan	This work
WS8N	Wild-type strain; spontaneous Nalr	38
Escherichia coli strains
S17-1	recA endA thi hsdR RP4-2-Tc::Mu::Tn7	53
TOP10	Cloning strain	Invitrogen
Plasmids
pB45	Plasmid source of the hygromycin cassette	42
pBBMCS53	Transcriptional uidA fusion vector; Gmr	35
pBBMCS53_cheA4	pBBMCS53 carrying the cheOp3 promoter	This work
pBBMCS53_cheOp1L	pBBMCS53 carrying 652 bp upstream of cheOp1	This work
pBBMCS53_cheOp1S	pBBMCS53 carrying 591 bp upstream of cheOp1	This work
pCR2.1-TOPO	Cloning vector	Invitrogen
pFlgC1	Integration vector used to create a C-terminal fusion with the FLAG tag epitope; Spcr	45
pJQ200_ΔctrA::hyg	pJQ200mp18 carrying ΔctrA::hyg	This work
pJQ200_mcpB::uidA-aadA	pJQ200mp18 carrying mcpB::uidA-aadA	This work
pJQ_ΔRSWS8N_RS04480::kan	pJQ200mp18 carrying ΔRSWS8N_RS04480::kan	This work
pJQ_ΔRSWS8N_13335::hyg	pJQ200 mp18 carrying ΔRSWS8N_13335::hyg	This work
pJQ200mp18	Mobilizable suicide vector for R. sphaeroides; Gmr	41
pRK_ctrA	pRK415 derivative expressing ctrA	21
pTZ_ctrAupdown	pTZ19R Bam− carrying ctrAupdown	21
pTZ EcoRI− SacI−	pTZ19R without EcoRI and SacI sites	Laboratory collection
pTZ_Eco− SstI− ΔRSWS8N_RS04480::kan	pTZ19R EcoRI− SacI− carrying ΔRSWS8N_RS04480::kan	This work
pTZ_Eco− SacI− RSWS8N_RS04480	pTZ19R EcoRI− SacI− carrying RSWS8N_RS04480	This work
pTZ19R	Cloning vector; Apr	Thermo Scientific
pTZ19R Bam−	pTZ19R without BamHI site	Laboratory collection
pTZ19R BamHI−_ΔRSWS8N_RS13335::hyg	pTZ19R Bam− carrying ΔRSWS8N_RS13335::hyg	This work
pTZ19R BamHI−_RSWS8N_RS13335	pTZ19R Bam− carrying RSWS8N_RS13335	This work
pTZ18R_mcpB	pTZ18R carrying mcpB	26
pTZ18R_mcpB::uidA-aadA	ptZ18R carrying mcpB::uidA-aadA	This work
pUC4K	Plasmid source of the kanamycin cassette	Pharmacia
pWM5	Vector source of the uidA-aadA cassette	46

Oligonucleotides.

The oligonucleotides used in this work are listed in Table S1 in the supplemental material.

Isolation of mutant strains.

In general terms, to inactivate RSWS8N_RS13335 and RSWS8N_RS04480, which encode proteins with characteristic bEBP signatures, a suicide vector (41) carrying the target gene disrupted with an antibiotic resistance gene was used to perform allelic exchange. Specifically, to inactivate RSWS8N_RS13335, which encodes a 446-residue protein, the oligonucleotides 159-1 and 159-2 (both carrying an XbaI recognition site at the 5′ end), were used to PCR amplify a 2,541-bp product containing the complete coding region of RSWS8N_RS13335, as well as regions upstream and downstream of this gene. This product was cloned into pTZ19RBam⁻, and the resulting plasmid was digested with BamHI and partially digested with BglII to remove a 573-bp portion of RSWS8N_RS13335 (residues 229 to 420). The resulting fragment was ligated to a BamHI-digested, 1.4-kb hygromycin resistance cassette (hyg), obtained by PCR from the pB45 plasmid (42). The plasmid carrying the ΔRSWS8N_RS13335::hyg allele was digested with XbaI, and the fragment containing the mutant allele was subcloned into pJQ200mp18 (41). This plasmid was then introduced into E. coli S17-1 (43) and subsequently transferred to R. sphaeroides by conjugation (44). Since pJQ200mp18 cannot replicate in R. sphaeroides, the double-recombination event was selected directly on LB agar plates in the presence of hygromycin and 5% sucrose. The mutant was verified by PCR.

A similar strategy was used to inactivate RSWS8N_RS04480, which encodes a 645-residue protein. The oligonucleotides 187-1 and 187-2 (both carrying an XbaI recognition site at the 5′ end) were used to PCR amplify a 2,968-bp product containing the complete coding region of RSWS8N_RS04480, as well as regions upstream and downstream of the target gene. This product was cloned into pTZ18REcoRI⁻ SacI⁻, and the resulting plasmid was digested with EcoRI and SacI to remove a 1,098-bp segment of RSWS8N_RS04480 (residues 4 to 370). The resulting fragment was end repaired with T4 DNA polymerase and ligated with a kanamycin resistance cassette (kan) obtained from the pUC4K plasmid. The plasmid carrying the ΔRSWS8N_RS04480::kan allele was digested with XbaI, and the fragment containing the mutant allele was subcloned into pJQ200mp18. This plasmid was then introduced into E. coli S17-1 and subsequently transferred to R. sphaeroides by conjugation. The mutant strains were selected, and the gene replacement was verified by PCR.

To obtain strain JV1 (mcpA-Flag), a 764-bp PCR product corresponding to the 3′ end of mcpA without its stop codon but including an extra guanine was obtained using the oligonucleotides mcpAtagFw and mcpAtagRv; the McpA-Flag fusion was obtained by cloning the PCR product into the pFlgC1 plasmid (45). The resulting plasmid was introduced into R. sphaeroides by conjugation, and integration into the chromosome by single recombination was selected. The Spc^r transconjugants were tested by PCR and Western blotting by using monoclonal antibodies against the Flag epitope. Subsequently, the ctrA⁺ allele in the JV1 strain was replaced with the ΔctrA::hyg allele by using the plasmid pJQ200_ΔctrA::hyg. This plasmid was obtained by inserting a BglII fragment carrying the hyg cassette into the BamHI site of the pTZ_ctrAupdown plasmid (21).

To obtain the mutant strains JV3 (mcpB::uidA-aadA) and JV4 (mcpB::uidA-aadA ΔctrA::hyg), we used the plasmid pTZ18R_mcpB (26), which carries a 1,830-bp fragment of the coding region of mcpB plus 107 bp upstream of the start codon and 40 bp downstream of the stop codon. This plasmid was digested with NcoI, end repaired with T4 DNA polymerase, and ligated to a 4-kb SmaI fragment obtained from pWM5, which carries the uidA-aadA cassette (46). The fragment containing the mcpB::uidA-aadA allele was subcloned into pJQ200mp18. This plasmid was introduced into AM1 and LC7 strains to replace the chromosomal mcpB gene by following the procedure described above.

Plasmids used in this work.

A transcriptional fusion between the regulatory region of cheOp1 and the reporter gene uidA was obtained by cloning a 652-bp fragment of the upstream region of cheOp1 into the pBBMCS53 plasmid (35). This fragment was obtained by PCR using the oligonucleotides cheOpromFw and cheOpromRv. The resulting plasmid was named pBBMCS53_cheOp1L. The pBBMCS53 plasmid was designed to generate transcriptional fusions with the uidA gene, which encodes the enzyme β-glucuronidase (35). The plasmid pBBMCS53_cheOp1S was obtained by cloning a 591-bp PCR product carrying the regulatory region of cheOp1 (without the putative CtrA binding site identified in this work) into the pBBMCS53 plasmid. The PCR product in this plasmid was generated using the oligonucleotides cheOprommut and cheOpromRv. Both plasmids were introduced into AM1 and EA1 strains by conjugation.

Molecular biology techniques.

Standard methods were used to obtain chromosomal or plasmid DNA (40). Restriction and other DNA-modifying enzymes were acquired from New England BioLabs (NEB), Roche, or Invitrogen. For sequencing, plasmids were purified using the Illustra plasmidPrep mini spin kit (GE Healthcare Life Sciences). Chromosomal or plasmid DNA was amplified with the appropriate oligonucleotides using PrimeSTAR HS DNA polymerase (TaKaRa Bio Inc.) according to the manufacturer's recommendations.

RNA isolation and RT-PCR assays.

Total RNA was isolated from cells grown to an optical density at 600 nm (OD₆₀₀) of 0.5 in Sistrom's minimal medium using hot phenol (47). After purification, the samples were incubated with RNase-free DNase I using the DNA-free DNA removal kit (Thermo Fisher Scientific). Reverse transcription and PCR amplification were performed with the Access RT-PCR system (Promega) according to the manufacturer's instructions. A control reaction mixture lacking reverse transcriptase was run in parallel. The PCR products were analyzed by agarose or acrylamide gel electrophoresis. The oligonucleotides cheY5XbaFw and cheY5PstRev were used to test the region between RSWS8N_02465 and cheY5. CheY6UP and tlpTdown were used to test the region between cheY6 and tlpT. The oligonucleotides cheY4Fw and cheY4Rev were used to amplify an internal region of cheY4. FlgHFw and flgHRv were used as a control to amplify an internal region of flgH.

Primer extension.

Total RNA was purified as previously described (47). The oligonucleotide cheOp2B was 5′ end labeled with [γ-³²P]ATP. Then, 50 μg of total RNA was hybridized with the labeled oligonucleotide at 42°C in the presence of 50% formamide. The hybrid RNA-DNA was ethanol precipitated and resuspended in a 30-μl reaction volume. The cDNA was synthetized using avian myeloblastosis virus (AMV) reverse transcriptase. The products were subjected to electrophoresis on a 6% polyacrylamide–8 M urea gel. As a molecular marker, we used DNA from PhiX174 digested with HinfI (Promega) and 5′ end labeled with [γ-³²P]ATP. The images were visualized using a Typhoon scanner (GE Healthcare Life Science) and quantified using ImageJ software (48).

Western blot analysis.

Samples were obtained from exponentially growing cultures at an OD₆₀₀ of 0.6 from heterotrophically grown cultures and at an OD₆₀₀ of 0.3 for photoheterotrophic cultures grown in 80 μM succinic acid. Whole bacterial cells were lysed by boiling in a solution containing 2% SDS, 1% β-mercaptoethanol, and 50 mM Tris (pH 7.5). These samples were separated on 12% polyacrylamide gels. The proteins were electrophoretically transferred to nitrocellulose membranes (49). The membranes were then probed with antibodies raised in rabbits against CheY2, CheY3, CheY4, CheY5, and CheY6 (1:3,000) or probed with a commercial mouse antibody against the Flag epitope tag (1:10,000) (Sigma-Aldrich). Anti-rabbit or anti-mouse IgG antibodies (1:30,000) (Sigma-Aldrich) coupled to alkaline phosphatase were used for chemiluminescent detection with CDP-star as the substrate (Applied Biosystems). His-tagged CheY proteins were purified by following previously reported protocols (50). Rabbit immunizations were performed as reported elsewhere (49). Representative results of three independent experiments are shown in Fig. 2A, S1, 4A, 5A, and 6A.

β-Glucuronidase activity assay.

β-Glucuronidase assays employed 4-methylumbelliferyl-β-d-glucuronide (MUG) as the substrate along with sonicated cell extracts, as previously described (51). Samples of 100 μl were taken at three time points between 5 and 20 min and then mixed with 0.9 ml of stop buffer (0.2 M Na₂CO₃). Fluorimetric determinations were made on a Perkin-Elmer LS-5 apparatus (excitation, 360 nm; emission, 446 nm). The fluorimeter was calibrated using 4-methylumbelliferone (Sigma-Aldrich) standards. Specific enzyme activity in cell extracts was expressed as picomoles of methylumbelliferone per minute per milligram of protein. Protein content was determined with a Bio-Rad protein assay kit, using bovine serum albumin as the standard.

Identification of the CtrA and σ²⁸ binding sites.

The sequence upstream of cheY4 was analyzed using a position weight matrix (PWM) built using the σ²⁸ promoters from E. coli (36) with the tool matrix-scan (52) included in the Regulatory Sequence Analysis Tools (RSAT) (http://embnet.ccg.unam.mx/rsa-tools/). The sequence upstream of the first gene in the cheOp1 operon, RSWS8N_02465, was analyzed using the same tool, with a PWM built from the regulatory region of 54 genes identified to be controlled by CtrA in C. crescentus (31).