ABSTRACT
The flagellar systems of Escherichia coli and Salmonella enterica exhibit a significant level of genetic and functional synteny. Both systems are controlled by the flagellar specific master regulator FlhD4C2. Since the early days of genetic analyses of flagellar systems it has been known that E. coli flhDC can complement a ΔflhDC mutant in S. enterica. The genomic revolution has identified how genetic changes to transcription factors and / or DNA binding sites can impact the phenotypic outcome across related species. We were therefore interested in asking: using modern tools to interrogate flagellar gene expression and assembly, what would the impact be of replacing the flhDC coding sequences in S. enterica for the E. coli genes at the flhDC S. entercia chromosomal locus? We show that even though all strains created are motile, flagellar gene expression is measurably lower when flhDCEC are present. These changes can be attributed to the impact of FlhD4C2 DNA recognition and the protein-protein interactions required to generate a stable FlhD4C2 complex. Furthermore, our data suggests that in E. coli the internal flagellar FliT regulatory feedback loop has a marked difference with respect to output of the flagellar systems. We argue due diligence is required in making assumptions based on heterologous expression of regulators and that even systems showing significant synteny may not behave in exactly the same manner.
IMPORTANCE
The bacterial motility organelle known as the flagellum is shared across many bacterial species. Escherichia coli and Salmonella enterica have underpinned our appreciation of how bacteria express and assemble the bacterial flagellum for over half a century. We show that even though the E. coli and S. enterica flagellar systems look genetically identical, they input regulatory signals into the flagellar system differently. Our conclusions are based on experiments where we carefully transfer the master flagellar regulator from E. coli into the S. enterica chromosome and measure a range of outputs relating to flagellar gene expression, assembly and functional output.
INTRODUCTION
The flagellum in the enteric bacteria, Escherichia coli and Salmonella enterica, has been studied extensively for over fifty years and provides the canonical example for bacterial motility. These studies have revealed not only the complex structure of the enteric flagellum but also its role in host colonization, pathogenesis, and cellular physiology (1-4). In addition, these studies have identified many of the complex regulatory processes that coordinate the assembly and control of this exquisitely complex biological machine (3-5).
The flagellum in E. coli and S. enterica are structurally very similar and are often tacitly assumed to be effectively identical aside from differences in the filament structure. However, in the case of regulation, these assumptions are based more on sequence similarity rather than on actual experimental data (5) (6). Indeed, a number of studies have shown that these two systems are regulated in entirely different manners in response to environmental signals despite strong gene synteny. For example, many common E. coli strains are motile only during growth in nutrient-poor conditions whereas many common S. enterica strains are motile only during growth in nutrient-rich conditions (7). In addition, E. coli is more motile at 30°C than at 37°C whereas motility S. enterica is generally insensitive to these temperature differences (8). E. coli flhDC are transcribed from a single transcriptional start site that is responsive to OmpR, RcsB and CRP regulation, to name only a few regulatory inputs (8). In contrast S. enterica flhDC transcription is significantly more complex with up to 5 transcriptional start sites, albeit with only a subset being responsible for the majority of flhDC transcription (9).
Part of the problem is that different questions have been asked when studying the regulation of motility in these two bacterial species. Most studies in E. coli have focused on the environmental signals and associate regulatory process that induce bacterial motility. In particular, they have focused on the processes that regulate the expression of the master flagellar regulator, FlhD4C2 (8). Most studies in S. enterica, on the other hand, have focused on the regulatory processes that coordinate the assembly process following induction (4). In particular, they have focused on the downstream regulatory processes induced by FlhD4C2 (3).
Despite differences in regulation, the protein subunits of master flagellar regulators, FlhC and FlhD, exhibit high sequence similarity sharing 94 and 92% identity, respectively, between E. coli and S. enterica. Given that modifications to transcription factors and/or promoter structure can lead to divergence in regulatory circuits (10), we were interested in how FlhD4C2 functions in different genetic backgrounds? Previously, it was shown that E. coli flhDC can complement a ΔflhDC mutant in S. enterica, suggesting that these proteins are functions identical in the two bacterial species (11). However, it is not clear whether they are regulated in the same manner. We, therefore, investigated the impact of replacing the native master regulator in S. enterica with the one from E. coli. Defining the impact of known FlhD4C2 regulators such as ClpP, YdiV, FliT and FliZ on the two complexes suggest that these two species have adapted in how they perceive FlhD4C2. We argue that these phenotypic differences arise from adaptations E. coli and S. enterica have made during evolution to expand or modify cellular function with respect to movement within specific environmental niches.
RESULTS
Orthologous flhDC from E. coli can functionally complement flhDC in S. enterica
Given the similarities between the flagellar systems in S. enterica and E. coli, we sought to determine whether the FlhD4C2 master regulator is functionally equivalent in these two species of bacteria. To test this hypothesis, we replaced the flhDC genes in S. enterica flhDCSE) with the flhDC genes from E. coli (flhDCEC). The reason that we performed these experiments in S. enterica rather than E. coli was that the flagellar system is better characterized in the former, particularly with regards to transcriptional regulation. To avoid plasmid associated artefacts associated with the ectopic expression of flhDC, we replaced the entire S. enterica flhDC operon with the flhDC operon from E. coli at the native chromosomal locus (Figure S1).
We first tested whether flhDCEC was motile as determined using soft-agar motility plates. As shown in Figures 1A and B, these strains formed rings similar to the wild type. These results demonstrate that flhDCEC is functional in S. enterica. However, motility plates measure both motility and chemotaxis and do not provide any insights regarding possibly changes in the number of flagella per cell. To determine the impact flhDCEC had upon flagellar numbers we used a FliM-GFP fusion as a proxy for flagellar numbers (Figure 1C). When this fluorescent protein fusion is expressed in cells, it forms spots associated with nascent C-rings that loosely correlate with the number of flagella (12-14). By counting the number of spots per cell, we can determine the number of flagella made per cell. As shown Figure 1C, flhDCEC did not change flagellar numbers as compared to the wild type. These results demonstrate flhDCEC induces flagellar gene expression at similar levels as the wild type.
flhDC requires a specific transcription rate to maintain optimal flagellar numbers
The flagellar network in S. enterica contains a number of feedback loops to ensure that the cells regulate the number of flagella produced (4). One possibility is that these feedback loops mask any differences in FlhD4C2EC activity. To test this hypothesis, we replaced the native PflhD promoter with the tetracycline-inducble PtetA/tetR promoters. We then measured flagellar gene expression using a luciferase reporter system (15). In this case, a consistent and significant change in flagellar gene expression was observed when comparing FlhD4C2EC to FlhD4C2SE activity (Figure 2). Maximal expression of PflgA and PfliC, chosen to reflect flagellar gene expression at different stages of flagellar assembly (5), for both complexes was observed between 10 and 25 ng/ml of anhydrotetracycline, when flhDC transcription was from PtetA (Figure 2A and B). In contrast, PtetR, the weaker of the two tetracycline inducible promoters, reached a maximal output between 50 to 100 ng/ml anhydrotetracycline. In both scenarios the output for FlhD4C2EC control was lower than for the native FlhD4C2SE complex (Figure 2A and 2B).
We also measured the number of FliM-GFP foci at different anhydrotetracycline concentrations. PtetR::flhDC expression generated on average of approximately two FliM-foci per cell at 25 ng/ml of anhydrotetracycline for both FlhD4C2 complexes (Figure 2C). In contrast, 5 ng/ml induction of the PtetA::flhDCEC strain was sufficient to generate typical FliM-foci numbers (approx. 8 flagellar foci per cell). Even with the strong decrease in average foci per cell at these levels of induction, the number of basal bodies observed is sufficient to allow motility at comparable levels in the motility agar assay (Figure S2).
Replacement of flhC but not flhD in S. enterica with the E. coli orthologs affects motility
The hetero-oligomeric regulator FlhD4C2 is unusual in bacteria as the majority of transcriptional regulators are believed to be homo-oligomeric complexes. To determine the relative contributions of the two subunits, we individually replaced the flhC or flhD genes from S. enterica with their ortholog from E. coli (Figure S1). When we tested the two strains using motility plates, we found that motility was inhibited in the strain where flhCEC replaced the native S. enterica flhC (Figure 3A; blue bars), with an 88% reduction in swarm diameter when compared to WT S. enterica. The introduction of flhDEC compared to flhDCEC or flhDCSE produced swarms of a comparable size (Figure 3A; blue bars).
Using the dose-dependent inducible PtetA promoter(16) we observed that PtetA expression of flhCEC led to reduced PflgA transcription and strongly reduced PfliC transcription (Figure 4). Strains expressing flhDEC in S. enterica showed a mild increase in PflgA gene expression and a similar response for PfliC, although these changes were not significant (P = 0.32) (Figure 4). These data suggest that the combination of FlhDSE and FlhCEC generates an inefficient FlhD4C2 complex, resulting in reduced motility.
Orthologous FlhC and FlhD interaction is species specific and a key determinant of promoter recognition by the FlhD4C2 complex
The results above demonstrate that flhCEC is not functionally identical to flhCST. One possibility is that that FlhCEC is impaired in FlhD4C2 for DNA-binding. Alternatively, the stability of the FlhD4C2 complex is reduced in the flhCEC strain, leading to reduced FlhD4C2 activity. To test these hypotheses, we purified all combinations of the FlhD4C2 complex using affinity (Ni+ and heparin) chromatography (Figure 5A). In each complex, FlhD was tagged with a carboxy-terminal hexa-histidine to facilitate affinity purification. Such expression constructs have previously been used successfully to purify the FlhD4C2 complex (17, 18). Using either Ni+ affinity or heparin purification, we observed complete complex retrieval for three combinations (Figure 5A). FlhC recovery was less efficient in the FlhDSE/FlhCEC complex. In contrast, no FlhDSE/FlhCEC complex was recovered via Heparin purification, used to mimic DNA during protein purification of DNA-binding proteins (Figure 5A). This suggests that the FlhDSE/FlhCEC complex is less stable, resulting on a lower yield of complex retrieval.
We next used the EMSA assays to test all four protein complexes for their ability to bind the S. enterica PflgAB promoter region. Quantification of the DNA shifts showed that complexes containing the orthologous FlhCEC reduced the PflgAB promoter binding profile, compared to FlhCSE complexes (Figure 5B). This is consistent with FlhC being the DNA binding subunit of the complex and the variation in FlhD4C2 activated promoter-binding sites between S. enterica and E. coli (19). Therefore, these results suggest that FlhC is a key determinant of DNA binding ability. Furthermore, the reduction in FlhCEC motility and flagellar gene expression in S. enterica is a result of the FlhDSE/FlhCEC complex being unstable, ultimately reducing the cellular concentration of the FlhD4C2 complex.
FlhD4C2EC responds to proteolytic regulation
S. enterica and E. coli both regulate the FlhD4C2 complex through ClpXP-mediated proteolytic degradation. Proteolytic degradation of FlhD4C2 plays a fundamental role in facilitating rapid responses to environmental changes that require motility (20, 21). The FlhD4C2 complex has a very short half-live of approximately 2-3 minutes (22). Proteolytic degradation of FlhD and FlhC is regulated in E. coli and S. enterica by YdiV (23).
However, ydiV is not expressed under standard laboratory conditions in model E. coli strains, suggesting that ClpXP activity is modulated in a species-specific manner (7).
Previous work has shown that YdiV delivers FlhD4C2 complexes to ClpXP for degradation (24). We have assessed the impact on motility for ΔclpP and ΔydiV mutations (Figure 3). The ΔclpP and ΔydiV mutants exhibited improved motility and flagellar gene expression, including the FlhDSE/FlhCEC strain (Figure 3A and B). These results suggest that proteolytic degradation mechanism of FlhD and FlhC, and its regulation, is common to E. coli and S. enterica.
To complement the motility assays, we investigated how ΔclpP and ΔydiV mutations impact the number of FliM-foci in cell. Both ΔclpP and ΔydiV mutants showed an increased number of FliM-foci compared to the wild type (Figure 6 A-C). For flhCEC strain, FliM-foci were observed in 13% of the population where individual cells exhibited just one or two foci. However, the ΔclpP or ΔydiV mutants increased the flagellated population of the flhCEC strains to 51 and 46 % respectively, albeit with the majority still possessing only a single FliM focus (Figure 6 B and C).
FliT and FliZ regulation of FlhD4C2 complexes
FlhD4C2 activity has an additional level of regulation in S. enterica via the flagellar-specific regulators FliT and FliZ. FliT functions as an export chaperone for the filament cap protein, FliD, and is a regulator of FlhD4C2 activity (17, 25). FliT disrupts the FlhD4C2 complex but is unable to disrupt a FlhD4C2:DNA complex. Therefore, FliT modulates availability of FlhD4C2 complexes for promoter binding (17). In contrast, FliZ is a negative regulator of ydiV expression and thus increases the number of FlhD4C2 complexes in S. enterica (26, 27).
In motility assays of ΔfliT mutants, we observed a difference between our different flhDC strains. Motility is increased in a ΔfliT mutant background in S. enterica ((28) and Figure 3A). However, when flhDCEC and flhDEC replaced the native genes, a reduced swarm size was observed (Figure 3A). Furthermore, quantification of PfliC activity agreed with the motility profile for ΔfliT mutants, where flhDCEC and flhDEC containing strains had reduced promoter activity compared to wild type (Figure 3B). This suggests that the FlhD4C2 complexes are being perceived differently by FliT in S. enterica. The results for ΔclpP and ΔydiV mutants suggests that this is not due to protein stability, as all complex combinations reacted in a comparable fashion (Figure 3).
In contrast, the loss of fliZ resulted in a consistent reduction in motility, except for the flhCEC strain. However, as the flhCEC strain was already impaired in motility, it is possible that the resolution of the motility assay was unable to identify differences in ΔfliZ mutant. Flagellar gene expression activity did, however, suggest a 2-fold drop in PfliC expression in the flhCEC ΔfliZ strain as compared to the otherwise wild-type (Figure 3B).
Analysis of FliM-foci distribution in ΔfliT mutant reinforced the observed discrimination of flhDCEC and flhDEC gene replacements. Calculating the average foci per cell, S. enterica ΔfliT mutants showed an increased average number of foci per cell from 2.9 to 6.3, while the flhDEC (fliT+: 3.4 versus ΔfliT: 4.2) and flhDCEC replacements (fliT+: 3.6 versus ΔfliT: 2.7) exhibited no significant changes (Figure 7A). Interestingly, in a ΔfliZ mutant background, the FliM-foci analysis was able to differentiate flhDCEC and flhDEC from the native S. enterica flhDC strain. Both replacements exhibited an increase in the average foci compared to S. enterica ΔfliZ (Figure 7A).
These data suggest that there is a fundamental difference in how the FlhD4C2 complexes in E. coli and S. enterica respond to, at least, FliT regulation. There are two explanations for this: a) the E. coli combinations are being regulated via an unidentified mechanism in S. enterica or b) that they are insensitive to FliT regulation. Both arguments predict that in E. coli FlhD4C2 may respond differently to FliT regulation. Comparing S. enterica and E. coli does indeed identify a difference in the response to a ΔfliT mutant. While a ΔfliT mutant in S. enterica leads to a consistent increase in FliM-foci, no significant difference is noted for an E. coli ΔfliT mutant compared to E. coli wild type (Figure 7B). This suggests that the regulatory impact of FliT is very different in these two flagellar systems and the role FliT plays in S. enterica is potentially adaptive and species specific.
DISCUSSION
Two model flagellar systems that form the foundation of the flagellar field are those from the enteric species E. coli and S. enterica. These two systems have led to key discoveries in relation to many aspects of flagellar structure, type 3 secretion, flagellar cell biology and the regulation of flagellar assembly. Textbook explanations suggest that most flagellar systems are being activated, regulated and built according to the models for E. coli and S. enterica. Modifications of transcriptional regulatory circuits contribute to the phenotypic diversity we see in closely related gene sets and we are only now able to investigate this in depth due to the tools available. Here we have taken a simple step and asked how do orthologous FlhD4C2 complexes function in the closely related species E. coli and S. enterica?
At the onset of our work it was known that FlhD4C2 from E. coli could sustain motility in S. enterica(11). Our work was focussed on understanding and defining the species-specific differences in the regulon of two orthologous genes. Here we took advantage of the well-defined flagellar assembly tools to measure outputs such as, motility, flagellar assembly per cell and flagellar gene expression. Bioinformatic analysis identifies only an 8 and 6% identity difference between FlhD and FlhC in E. coli and S. enterica respectively, suggesting that these proteins function in an analogous fashion. It is well established that related taxa usually rely on orthologous regulators to coordinate response to a given signal (10).
The fine detail of the differences in the FlhD4C2 complexes only became apparent when we began to focus on their effect on flagellar gene expression and flagellar assembly. In all of our assays FlhD4C2EC exhibited a reduction in flagellar gene expression compared to FlhD4C2SE. Biochemical analysis of isolated complexes showed that FlhCEC had weaker DNA binding ability to the PflgAB promoter region from S. enterica, consistent with previous investigations into FlhD4C2 DNA binding activity (19). The isolation of FlhD4C2 complexes from our strains suggested that a key aspect of the phenotypes we observed, was the stability of the complexes formed.
With respect to flhDC transcription we show a discrepancy in flagellar numbers defined by FliM-foci when using PtetA/PtetR::flhDC expression. This was somewhat surprising as all constructs exhibited good swarming ability on motility agar plates (Figure S2). Original studies on the regulation of PtetA/PtetR from Tn10 have shown that these two promoters have differing activities but both respond to TetR regulation. We show that even though maximal activity of PflgA and PfliC can reach 40-50% of PtetA::flhDC expression for PtetR strains, this results in an average of 2 flagella per cell. This suggests that even though the majority of the literature states that E. coli and S. enterica produce between 4 and 8 flagella per cell, only 1 or 2 per cell is needed for an optimal output of the system with respect to motility agar assays.
It has been shown that FliT interacts with FlhC and that in S. enterica the output of this circuit is to destabilize FlhD4C2 complexes that are not bound to DNA. Our data suggests that this level of regulation does not impact E. coli FlhC. The nature of the adaptability needed by the favourable conditions to drive motility in E. coli may have led to the FliT regulatory input becoming less critical. Similarly, the impact of FliZ regulation becomes apparent for FlhDEC containing complexes when we assess flagellar numbers. FliZ regulates the transcription of ydiV in S. enterica (27). It is plausible that the impact in changing ydiV regulation is the source of this differentiation, especially as YdiV is proposed to interact with FlhDSE. Furthermore, we know that ydiV is not expressed in model E. coli strains, strengthening the argument that FlhDEC has adapted to the absence of YdiV or vice versa FlhDSE to YdiV.
Importantly our analysis shows that even though these two systems are genetically similar, investigation of FlhD4C2 activity identifies subtle but key differences into how the FlhD4C2 complex is modulated in two closely related species. We argue that this is a valid example of the caution needed in the age of synthetic biology to exploit heterologous systems in alternative species or chassis’. Our data shows that even systems showing significant synteny may not behave in exactly the same manner and due diligence is required in making assumptions based on heterologous expression.
MATERIALS AND METHODS
Bacterial Strains and Growth conditions
S. enterica and E. coli strains used in this study have been previously described elsewhere (12, 15, 17, 28). This study used S. enterica serovar Typhimurium strain LT2 as the chassis for all experiments. E. coli genetic material was derived from MG1655. All strains were grown at either 30°C or 37°C in Luria Bertani Broth (LB) either on 1.5% agar plates or shaken in liquid cultures at 160 rpm (17). Antibiotics used in this study have been described elsewhere (29). Motility assays used motility agar (17) incubated at 37°C for 6 to 8 hours. Motility swarms were quantified using images captured on a standard gel doc system with a ruler in the field of view and quantified using ImageJ to measure the vertical and horizontal diameter using the average as the swarm size. All motility assays were performed in triplicate using single batches of motility agar.
Genetic Manipulations
For the replacement of flhDC coding sequences the modified lambda red recombination system described by Blank et al (2011) was used (30). Deletion of clpP, ydiV, fliT and fliZ was performed using the pKD system described by Datsenko and Wanner (2000) (31).
PtetA / PtetR replacements of the Pf lhDC region was also performed using Datsenko and Wanner (2000) with the template being Tn10dTc (32). For Blank et al (2011) replacement experiments we used autoclaved chlortetracycline instead of anhydrotetracycline as described for the preparation of Tetracycline sensitive plates (33). All other gene replacements were performed as previously described (17). All primers used for these genetic manipulations are available on request.
Quantification of flagellar gene expression
Flagellar gene expression assays were performed using the plasmids pRG39::cat (PfliC) and pRG52::cat (PflgA) (15). Both plasmids were transformed into strains using electroporation. Gene expression was quantified as described previously and analysis was based on a minimum of n = 3 repeats for each strain tested (15).
Quantification of FliM-GFP foci
FliM-GFP foci were quantified using Microbetracker on images captured using a Nikon Ti inverted microscope using filters and exposure times described previously (14). Strains were grown to an OD600 of 0.5 to 0.6 and cells immobilised using a 1 % agarose pad containing 10 % LB (14, 17). For each strain a minimum of 5 fields of view were captured from 3 independent repeats. This allowed analysis of approximately 1000 - 1500 cells per strain. For the comparison of FliM foci in E. coli ΔfliT to S. enterica ΔfliT shown in Figure 7B the chemostat growth system described by Sim et al (2017) was used. For this experiment the growth rate of both strains was similar to batch culture in LB at 37°C where the media used was a MinE base with 0.1% Yeast extract and 0.2% glucose (14, 17).
Purification of FlhD4C2 complexes
Purification of proteins complexes was based on previously described methods (17). Wild type FlhD4C2SE was purified using pPA158. The other 3 complexes were purified from plasmids generated using the New England Biolabs NEBuilder DNA Assembly kit on the backbone of pPA158. The E. coli strain BL21 was used for all protein induction experiments prior to protein purification using either a pre-equilibrated 5ml His-trap column or a 5ml heparin column (GE Healthcare). Proteins were visualised using Tricine-based SDS polyacrylamide gel electrophoresis and standard commassie blue staining (17).
Electrophoretic mobility shift assay (EMSA)
All EMSA assays were performed using Ni++ (his-trap) purified proteins as this allowed analysis of all four complexes (Figure 5A). Buffer exchange from elution buffer to a 100mM Tris-HCl, 300 mM NaCl 1mM DTT (pH 7.9) buffer was performed through 10 cycles of protein concentration in VivaSpin columns with 20 ml buffer reduced to 5 ml per round of centrifugation at 4500 rpm. A protein concentration range of 100 to 700 nM was used with 80 ng / ml of a PCR product containing PflgAB from S. enterica. After incubation bound and unbound DNA were resolved using 5% acrylamide gels made with 1x TBE buffer.
Quantification of gel images was performed using ImageJ.
ACKNOWLEDGEMENTS
PDA would like to recognize the internal financial support of ICAMB during this study. The stipend and research costs for the PhD of AA was provided by The Ministry of Higher Education and Scientific Research (Iraq). We would like to thank the financial support of the Newcastle University Faculty of Medicine for providing the John W illiam Luccok and Ernest Jeffcock Research PhD Studentship to MS for this study. PAH would like to acknowledge the support of iUK/BBSRC (grant: BB/N023544/1), NERC (grant: NE/M001415/1), the University of Strathclyde and the Microbiology Society for funding. We would also like to thank all lab members for feedback on the project during the experimental and writing phases.