Introduction

Like many other bacteria, Pseudomonads swim and swarm using rotating flagella powered by membrane ion gradients to relocate to environments optimal for their metabolism1. Bacteria can control flagellar-driven motility in a number of ways, including through control of flagella assembly2,3 and taxis signal-transduction pathways that alter the direction of flagella rotation4. One of the most recently identified means of control is through the second messenger cyclic di-GMP [bis-(3′–5′)-cyclic dimeric guanosine monophosphate; c-di-GMP]. In addition to motility control, this near ubiquitous bacterial second messenger is also involved in co-ordinating developmental processes, regulation of virulence determinants, and the transition to biofilm formation5,6,7.

In Pseudomonads, low intracellular levels of c-di-GMP are associated with a motile (flagellated) planktonic mode of growth, while elevated levels sequentially trigger slowing down of flagella motility for surface attachment, and production of adhesins and biofilm components for a consequent sessile lifestyle8,9,10. Diguanylate cyclases (DGCs) and phosphodiesterases (PDEs) – which control the dynamic changes in the intracellular levels of this second messenger, are abundant in most bacteria. DGCs contain a GGDEF motif within their catalytic A-site and synthesise c-di-GMP from two molecules of GTP; conversely, PDEs degrade c-di-GMP – either to linear pGpG [EAL-motif proteins] or to two molecules of GMP [HD-GYP motif proteins]. Although DGCs and PDEs mediate opposing functions, they are often linked together in multi-domain proteins. In most cases, however, one of the domains has lost its catalytic capacity and instead has been adapted to regulate the function of the protein11. So far only a few proteins have been shown to be bona fide dual functional proteins that possess both c-di-GMP synthesising and degradative activities. One such protein is the motility associated c-di-GMP signalling protein PP2258 of Pseudomonas putida12. Lack of PP2258 (or its over-expression) results in decreased motility due to elevated c-di-GMP levels, implying that c-di-GMP dependent signalling from PP2258 is involved in motility control of P. putida12,13.

The enzymatic processes that make and break c-di-GMP are fairly well understood. However, the upstream signals that control the activities of DGCs and PDEs, and the downstream target effector proteins (and RNAs) that respond to c-di-GMP signalling have not been as extensively elucidated14. One common domain of c-di-GMP responsive effector proteins is the PilZ domain. This type of domain undergoes conformational changes upon c-di-GMP binding, which leads to alterations in protein-protein interactions and allosteric affects. However, differences exist between individual PilZ domains when it comes to binding mode, stoichiometry, and quaternary structure15,16. In the enterics Escherichia coli and Salmonella, the PilZ domain protein YcgR co-localizes with flagella and acts as a brake in response to c-di-GMP by interacting with components of the flagella motor17,18,19. In these organisms, the non-rotating part of the motor – the stator – is formed by membrane complexes of the MotA/MotB proteins that provide the ion-translocating channels that energise the rotor. The stator lies above the FliG/FliM/FliN rotary switch complex, which drives rotation of the flagellum and also sets the rotational direction. The carboxy-PilZ domain of YcgR binds c-di-GMP with a 1:1 ratio and binding of c-di-GMP causes YcgR to adopt a more condensed conformation without altering its monomeric status20, while it’s amino-terminal domain is involved with interactions with its targets – MotA, FliG and/or FliM – to result in reduced torque and motility17,18,19.

Pseudomonas aeruginosa and other Pseudomonads have an additional set of Mot proteins – MotC/MotD21,22 – which in the case of P. aeruginosa appear to be the predominant complexes involved in swarming and swimming motility23. Pseudomonads also encode a distant relative of YcgR – FlgZ – that affects swimming motility of P. fluorescens and P. putida24. Most recently, the P. aeruginosa PA14 FlgZ counterpart has been shown to interact with MotC (rather than MotA) to control its swarming motility23. The structure and c-di-GMP binding properties of the P. putida KT2440 FlgZ counterpart – PP4397 – have previously been determined; unlike YcgR, PP4397 undergoes a dimer-to-monomer transition upon c-di-GMP binding in vitro25. Hence, in addition to low sequence identity, FlgZ/PP4397 exhibit disparate biochemical properties to those of YcgR.

Despite detailed structural studies, little is known about how expression of P. putida PP4397 is controlled, what upstream protein(s) can control its c-di-GMP responsive activity, and if it is a genuine functional counterpart of YcgR. In this work we provide evidence that expression of PP4397 is controlled by two alternative σ-factors – σ54 and the flagella specific σFliA – to allow coupling of PP4397 expression to other flagella motility related genes. Furthermore, we show that despite very limited amino acid sequence identity and different in vitro properties to those of YcgR, FlgZ/PP4397 lies downstream of PP2258 in c-di-GMP responsive motility control in P. putida and can be functionally replaced by YcgR in this process.

Results and Discussion

pp4397/flgZ is co-transcribed with flagellar associated genes dependent on σ54 and σFliA

In silico searches of genome sequenced P. putida KT2440 identified PP4397/FlgZ as the protein exhibiting highest identity to YcgR (17% and 24% identity with E. coli and Salmonella, respectively). Despite this low homology, similar to YcgR, PP4397/FlgZ also possesses a type I c-di-GMP binding PilZ and PilZN-like domain (Figs 1 and S1). The assembly and operation of Pseudomonad flagella depends on more than 50 genes arranged in clusters that are controlled by a four-tiered transcriptional regulatory cascade26. As depicted in Fig. 2A (upper), enteric ycgR lies in a monocistronic operon, while the P. putida flgZ/pp4397 gene lies within a cluster of genes encoding proteins involved in flagellar biosynthesis (Fig. 2A, lower). We therefore wanted to clarify if pp4397 is co-transcribed with any of these genes.

Figure 1
figure 1

PP4397 exhibits limited sequence identity with YcgR but maintains key residues involved in c-di-GMP binding. (A) Schematic illustration of the domains and % identity of YcgR and PP4397/FlgZ as detailed in the text. (B) The amino acid sequence of PP4397 is shown with nine highly conserved residues of PilZ domains (highlighted in bold and underlined) that encompass the RXXXR and (D/N)XSXXG motifs of c-di-GMP binding-proficient type I PilZ domains34. Residues, which when substituted by alanine, essentially abolish c-di-GMP binding by PP4397/FlgZ25 are highlighted in grey. A complete alignment of PP4397 and YcgR as in15,25 using ESPript35 is shown in Fig. S1, while the extensive homology with FlgZ proteins of representative Pseudomonads is shown in Fig. S2.

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Figure 2
figure 2

Transcription of PP4397 is dependent on both σ54 and σFliA. (A) Comparison of the genomic context of E. coli ycgR and P. putida flgZ/pp4397 genes. Upper schematic, illustration of E. coli MG1655 flgA, flgMN and ycgR genes (shown in black) with their cognate promoters indicated. Divergently transcribed genes are shown in grey. Lower, similar schematic of the P. putida flgA, flgM, flgN-like pp4396 and flgZ/pp4397 genes shown in black and the divergently transcribed cheV-3 and pp4398 genes shown in grey. The locations of primer pairs used for analysis of co-transcription of genes as depicted in panel B are indicated. (B) Agarose gels of PCR reactions using the indicated primer pairs (panel A) on P. putida genomic DNA, cDNA, or control samples where no reverse transcriptase was added to the cDNA reaction mixture (-RT). (C) In vivo transcription from PflgM in wild type P. putida KT2440 (1) and its FliA null (2) and RpoN null (3) counterparts carrying mono-copy transcriptional fusions to the promoter-less luxAB reporter genes (PP3733 to PP3735, Table S1). Graphed values are the average +/− standard deviation of six independent determinations from cultures grown to the stationary phase in LB (OD600 ~5.0 for wild-type and FliA null; ~2.1 for RpoN null). (D) Single-round in vitro transcription assays using 10 nM supercoiled DNA templates harbouring the σFliA –dependent P. putida Paer2 promoter (4; pVI1011) or PflgM (5; pVI2368) in the presence of 10 nM σFliA-RNAP. Inset shows images from one of two independent experiments used to obtain the graphed average values (PflgM upper; Paer2 lower). A comparison of the Paer2 and PflgM promoter sequences with the optimal consensus27 for P. putida σFliA is shown to the right.

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Reverse transcription polymerase chain reaction (PCR) assays with RNA isolated from P. putida KT2440 and primer pairs spanning pp4397 and adjacent genes showed that pp4397 is co-transcribed with the σFliA anti-σ-factor gene flgM and pp4396 – a gene encoding an FlgN family protein (Fig. 2B). An appropriately located σFliA-dependent promoter is found immediately upstream of the flgM-flgN-flgZ/pp4397 genes cluster (see Fig. 2A). Hence, this data is consistent with the idea that pp4397 lies within a tri-cistronic flgM-flgN-pp4397 operon, but does not exclude the possibility that internal promoters within flgM and/or flgN may also contribute to transcription of pp4397. Recent analysis of transcription of pp4397/flgZ counterparts in P. fluorescens F113 and P. putida KT2440 could only document a partial dependence on σFliA as assessed using FliA null strains10,24. Therefore, we extended the analysis to include the upstream flgA gene that encodes a protein involved in flagellar P-ring formation. As shown in Fig. 2B, transcription of the flgM-flgN-pp4397 operon also appears to be mediated by read-through transcription from a σ54-dependent promoter located upstream of the flgA gene.

To further substantiate the above findings, we performed in vivo and in vitro transcription assays. In vivo transcription of pp4397/flgZ was monitored using a luxAB (luciferase) transcriptional fusion generated downstream of pp4397 in wild-type, FliA null, and RpoN (σ54) null P. putida backgrounds. Consistent with co-dependence on both these σ-factors, transcriptional output was decreased but not abolished in both of the null strains as compared to wild-type when grown Luria-Bertani (LB) broth (Fig. 2C). Functionality of the identified σFliA promoter located upstream of the flgM-flgN-pp4397 tri-gene cluster (PflgM) was verified by single-round in vitro transcription assays with σFliA-RNA polymerase reconstituted from purified P. putida components (Fig. 2D). As anticipated by its high identity to the optimal consensus for P. putida FliA dependent transcription, the near consensus flgM promoter produces high levels of transcripts as compared to the previously analysed suboptimal σFliA-dependent promoter for aer227.

The difference in σ-dependence for the flgA promoters in Pseudomonads as compared to enterics is due to differences in the hierarchical expression of flagellar genes in these two species. Transcription of genes needed early, e.g. flgA, are dependent on σ54 in Pseudomonas and σ70 in E. coli and Salmonella26,28. Thus, while the genomic context of pp4397/flgZ is different from the moncistronic context of E. coli ycgR gene, transcriptional control of the flgA and flgMN counterparts is conceptually similar, with a promoter upstream of flgA generating read-through transcription of downstream genes within a σFliA-dependent operon. Given that the flgZ gene is highly conserved in sequence and synteny in all sequenced Pseudomonads24, including P. fluorescens F113, P. aeruginosa PA14, and P. aeruginosa PAO1 (Fig. S2), co-dependence on both σFliA and σ54 as found here for P. putida KT2440 is likely the case for these and other Pseudomonads. It is interesting to note that this regulatory arrangement would ensure σ54-dependent transcription of the flgM-flgN-pp4397 genes even in the absence of σFliA. Therefore, it would also result in σFliA-independent production of PP4397/FlgZ that serves to slow-down flagella (as detailed below) and production of the anti-σFliA-factor FlgM to block new de novo flagella production – two steps needed for preparation to enter the biofilm mode of growth9.

Lack of PP4397 results in altered swimming and swarming motility

Having established that PP4397 is co-ordinately regulated with genes of the flagella regulon, we next addressed its involvement in c-di-GMP responsive control of flagella-mediated motility. To test potential involvement of PP4397 in motility control, we first generated a null mutant of P. putida KT2701 (a streptomycin resistant derivative of the genome sequenced P. putida KT2440) in which the majority of the pp4397/flgZ gene was replaced by a tetracycline resistance cassette. When tested on rich (LB) 0.3% soft agar swimming motility plates or 0.5% agar swarming motility plates, the Δpp4397::Tc strain displayed only a slightly enhanced motility phenotype as compared to the wild type (Fig. 3).

Figure 3
figure 3

The reduced motility phenotypes of PP2258 null P. putida are partially rescued by the absence of PP4397. (A) Relative swimming motility of P. putida KT2701 wild type (WT) and the indicated null derivatives (Table S1) on 0.3% soft agar LB plates. Representative swim rings are shown above. (B) Relative swarming motility of the same strains on 0.5% agar plates. Representative swarm zones are shown to the right. Graphed values in all cases are averages with standard deviations calculated from three independent colonies and were normalized by setting the values of the wild type strain as 1. P-values shown for relevant comparisons were calculated with two-tailed student t-test (***P < 0.001; **P < 0.01).

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The above data is consistent with previous findings for E. coli and Salmonella in which a YcgR null counterparts only exhibited increased motility in a high c-di-GMP background – a condition that normally results in decreased motility20. Therefore, we also introduced the Δpp4397::Tc allele in P. putida KT2701 Δpp2258::Km, a previously analysed strain known to have elevated c-di-GMP levels and a swimming motility defect on soft agar plates despite having a wild type number of polar flagella12.

Both the Δpp2258::Km and the Δpp2258/Δpp4397 double mutant strains exhibit prolonged lag phases upon outgrowth from overnight cultures on rich media (LB), but once they attain exponential growth, have doubling times (41.3 +/− 2.9 min) similar to, but slower, than the wild type and the Δpp4397::Tc strains (36.2 +/− 1.7 min; see Fig. S3A). While the exact level of c-di-GMP in the Δpp2258::Km PP2258 null strain is unknown, elevated c-di-GMP levels in this strain and the Δpp2258/Δpp4397 double null strain are insufficient to provoke altered biofilm production or dispersal phenotypes (Fig. S3B–D), as judged using a microtitre dish-based assays that employs serial dilution to recapitulate biofilm growth and dispersal kinetics29. As detailed in Fig. S3, in both cases biofilm production and dispersal rates appear similar to wild type, despite a delay as a consequence of growth kinetics.

In contrast, the reduced motility seen for the Δpp2258::Km strain in both swimming and swarming abilities was significantly rescued in the double mutant [compare PP2258 null with the PP2258/PP4397 double null in Fig. 3A,B]. Even though exponentially growing cells were used for inoculation of the motility assay plates (see Methods) reduced growth rates as a consequence of elevated c-di-GMP levels probably, at least in part, underlies why full motility comparable to the wild-type strain could not be achieved. Taken together, the data in Figs 3 and S3 consolidate a role for PP4397/FlgZ of P. putida in swimming and swarming motility (but not biofilm production or dispersal), and provide the first evidence that PP4397/FlgZ functions downstream of PP2258 in response to modulation of c-di-GMP levels.

The c-di-GMP binding property of PP4397 is required to mediate motility control

To verify that the phenotype for the Δpp2258/Δpp4397 double null strain was not attributable to indirect effects on upstream genes within the flgM-flgN-flgZ/pp4397 operon, this strain was complemented with plasmids carrying either a native version of the pp4397/flgZ gene or a C-terminally FLAG-tagged version under control of the IPTG inducible lacIQ/Ptac promoter. The lacIQ/Ptac system of the expression plasmid is leaky and produced sufficient PP4397/FlgZ to reverse the effect of lack of PP4397 in the double Δpp2258/Δpp4397 strain – i.e. expression of PP4397 or PP4397-FLAG in the double mutants resulted in a reduced motility phenotype approximating that of the PP2258 null strain (Fig. 4). The motility phenotypes shown in Fig. 4 were unaffected by addition of 0.5 mM IPTG (data not shown). These results confirm that the motility rescue phenotype of the double mutant is due to lack of PP4397/FlgZ.

Figure 4
figure 4

The function of PP4397 in motility control is dependent on its c-di-GMP binding ability. Relative swimming motility of the double PP2258/PP4397 null derivative of P. putida KT2701 on 0.3% soft agar LB plates supplemented with carbenicillin. (A) Strains harbouring either a vector control (Vec. Cont.; pVI2300) or lacIQ/Ptac expression plasmids for derivatives of PP4397: c-di-GMP binding proficient wild type PP4397 (WT, pVI2301), or c-di-GMP binding deficient mutants R127A (pVI2302) or G162A (pVI2303). (B) Strains harbouring either a vector control (Vec. Cont.; pVI2300) or lacIQ/Ptac expression plasmids for FLAG-tagged derivatives of PP4397: c-di-GMP binding proficient wild type PP4397 (WT, pVI2304), or c-di-GMP binding deficient mutants R127A (pVI2305) or G162A (pVI2306). Graphed values are averages with standard deviations calculated from three independent colonies. Experiments were normalized by setting the values of the double mutant harbouring the vector control as 1 and P-values calculated with two-tailed student t-test (***P < 0.001). Images of representative swim rings are shown above the graphed values. The insert in panel B shows Western analysis of the FLAG-tagged PP4397 derivatives present in 10 µg of crude extract from the same cells. The cropped Western analysis image is derived from the same gel and is shown alongside molecular size markers in Fig. S4A.

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Biochemical analysis of PP4397 has shown that alanine substitutions of arginine 127 (R127A) or glycine 162 (G162A) both abolish the capacity of PP4397 to bind c-di-GMP. Arginine 127 is directly involved in binding of c-di-GMP, while glycine 162 is conserved among PilZ domain proteins and is probably needed for correct folding25. To confirm that c-di-GMP binding is required for PP4397 to exert its phenotypic effects, equivalent expression plasmids for native and C-terminally FLAG-tagged PP4397-R127A and PP4397-G162A derivatives were generated and tested as described for wild type PP4397. Neither of these c-di-GMP binding defective derivatives mediated a reduced motility phenotype (Fig. 4), even though they were expressed at the same levels as the wild type protein (Figs 4B and S4A). Thus, these results demonstrate that PP4397 is c-di-GMP responsive in vivo, and that c-di-GMP binding by PP4397/FlgZ is a prerequisite for its effects on P. putida motility.

YcgR, like PP4397, restores a motility defect in P. putida Δpp2258/Δpp4397

To ascertain if PP4397 and YcgR showed cross-species functionality, plasmids expressing PP4397-FLAG and E. coli YcgR-FLAG under control of an araC/PBAD promoter were introduced into E. coli MG1655-ΔyhjH/ΔycgR (which has elevated c-di-GMP levels due to the lack of the PDE YhjH) and P. putida Δpp2258/Δpp4397 (which also has elevated c-di-GMP levels due to the lack of PP2258). Relative swimming motilities were assayed on LB soft 0.3% agar swimming motility plates containing 0 to 1.0% L-arabinose. As anticipated, motility of the E. coli ΔyhjH/ΔycgR strain was greatly reduced by expression of YcgR-FLAG induced with either 0.2% or 1% L-arabinose, but not by expression of PP4397-FLAG (Fig. 5A). However, Western analysis revealed that expression levels of PP4397-FLAG were notably lower than those of YcgR-FLAG, which likely underlies the inability of PP4397 to cause an altered motility phenotype in this strain (expanded Western Fig. S4B). In marked contrast, both YcgR-FLAG and PP4397-FLAG greatly reduced motility of P. putida Δpp2258/Δpp4397 when expressed at similar levels (induced with 1% L-arabinose Figs 5B, and S4B). No reduction in motility was observed with c-di-GMP binding-deficient derivatives of either protein (PP4397-R127A-FLAG and its corresponding YcgR-R118A-FLAG counterpart, data not shown). These results lend strong support to the idea that despite their limited identity (Fig. 1), PP4397/FlgZ and YcgR are functional c-di-GMP responsive counterparts that act to control motility in P. putida and E. coli, respectively.

Figure 5
figure 5

Cross-species complementation - both PP4397 and E. coli YcgR restore a motility defect to the double PP2258/PP4397 null derivative of P. putida. (A) Relative swimming motility of E. coli MG1655-ΔyhjH/ΔycgR carrying araC/PBAD expression plasmid for P. putida PP4397-FLAG (pVI2370) or E. coli YcgR-FLAG (pVI2373), assayed on 0.3% soft agar LB motility plates containing the indicated amount of L-arabinose as inducer. Graphed values are averages with standard deviations calculated from three independent colonies and were normalized by setting the values of the double mutant harbouring the vector control as 1. P-values, calculated with two-tailed student t-test (***P < 0.001), are relative to the vector control. Images are of corresponding protein expression levels as revealed by separation of 10 µg of soluble protein and subsequent Western analysis. Cropped images are derived from the same experiment processed in parallel on the same gel, and are shown alongside molecular size markers in Fig. S4B. (B) Relative swimming motility of P. putida KT2701-Δpp2258/Δpp4397, carrying araC/PBAD expression plasmid for PP4397-FLAG (pVI2370) or YcgR-FLAG (pVI2373) as under panel A. Full images of the corresponding Western analysis for the cropped images are shown alongside molecular size markers in Fig. S4B.

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PP4397-EYFP locates to the cytosolic compartment

E. coli are peritrichous, with flagella distributed throughout their surface, and fluorescently tagged YcgR has previously been found to localize to puncta on the cells together with the flagellar apparatus17,19. Similar puncta have been observed for P. fluorescens when fluorescently tagged FlgZ was overexpressed in cells with elevated c-di-GMP23. In the case of P. aeruginosa, which possesses a single polar flagellum, mono-copy fluorescently tagged FlgZ exhibits co-polar localization with the motility apparatus, and could be observed in a higher percentage of cells when c-di-GMP levels were elevated24. P. putida KT2701 used here possess a bundle of 6 to 10 flagella located at a single pole27 and, therefore, co-localization of its FlgZ/PP3497 counterpart with flagella would be anticipated to result in a polar localization.

To determine if PP4397, similarly to YcgR and other FlgZ counterparts, co-localizes with the flagellar machinery, PP4397-EYFP fusions were introduced into P. putida, both in mono-copy in its native location on the chromosome, and in multi-copy on an araC/PBAD expression plasmid (as used in the motility assays in Fig. 5). Functionality of the PP4397-EYFP fusion, designed to have the same intervening residues as the YcgR-EYFP fusion, was confirmed by its maintenance of the reduced motility phenotype of the PP2258 null strain (Fig. S5). Western analysis was performed on cells harvested at the same time as cells were fixed for microscopy to facilitate correlation between images and corresponding protein expression levels.

In contrast to a mono-copy polar localization control (Aer2-EYFP13), mono-copy PP4397-EYFP was expressed at a higher level and localized to the cytoplasmic compartment in P. putida (Fig. 6, compare B to C). This apparent cytoplasmic localization was maintained in strains lacking PP2258 (Fig. 6, compare C and D) – i.e. under elevated c-di-GMP levels that results in altered swimming and swarming motility (Figs 3 and S5). This contrasts data for P. fluorescens and P. aeruginosa23,24, where cytosolic FlgZ counterparts could be visualized as puncta or at the pole under conditions where cellular c-di-GMP levels were elevated.

Figure 6
figure 6

Localization of fluorescent proteins in P. putida and E. coli strains. Cells shown are representative of >6 fields viewed in two or three independent experiments. Upper panels (A to D) and cognate western analysis are of strains cultured on LB. (A) P. putida KT2701 (negative control; cells examined n = 735). (B) P. putida KT2701::aer2-eyfp (positive control, mono-copy chromosomal fusion; cells examined n = 342 of which 64% exhibited polar localization). (C) P. putida KT2701::pp4397-eyfp (mono-copy chromosomal fusion; cells examined n = 219). (D) P. putida KT2701::pp4397-eyfp/Δpp2258 (PP2258 null with elevated c-di-GMP; cells examined n = 649). Western analysis of EYFP-tagged proteins expressed from mono-copy chromosomal translational fusions present in 50 and 25 µg of crude extract. Cell were harvested for Western analysis at the same time as fixing for imaging (after 2 to 2.5 hrs of growth; OD600 0.5 to 0.7), which contrasts those shown for the motility assays in Fig. 5 (harvested after 5 hr of growth; OD600 ~3.5). Note that Aer2-EYFP, although clearly visible at the pole of the cell in panel B, is expressed at much lower levels than PP4397-EYFP and is not detected at the exposure shown. The cropped image is derived from the same experiment processed in parallel on the same gel, and are shown alongside molecular size markers in Fig. S6A. Lower panels [E to H] and cognate western are of strains cultured on LB in the presence of 1% L-arabinose. Boxed images are differentially exposed cells for comparison of the presence or lack of puncta. (E) P. putida KT2701-Δpp2258/Δpp4397 (double PP2258/PP4397 null strain) carrying the multi-copy araC/PBAD pp4397-eyfp expression plasmid (pVI2374). Cells examined: n = 271, 0% with puncta. (F) P. putida KT2701-Δpp2258/ΔPP4397 carrying the multi-copy araC/PBAD ycgR-eyfp expression plasmid (pVI2375). Cells examined n = 282, 0% with puncta. (G) E. coli MG1655-ΔyhjH/ΔycgR (double YhjH/YcgR null strain) carrying the multi-copy araC/PBAD pp4397-eyfp expression plasmid (pVI2374). Cells examined: n = 343, 0% with puncta. (H) E. coli MG1655-ΔyhjH/ΔycgR carrying the multi-copy araC/PBAD ycgR-eyfp expression plasmid (pVI2375). Cells examined: n = 252, 25% with puncta [1 to 3 per cell]. Western analysis of EYFP-tagged proteins expressed from multi-copy translational fusions present in 25 and 12.5 µg of crude extract from P. putida (left) and E. coli (right). Cropped images are derived from the same experiment processed in parallel on the same gel, and are shown alongside molecular size markers in Fig. S6B.

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Cytosolic localization was also observed with PP4397-EYFP expressed from a multi-copy plasmid under inducing (1% L-arabinose) conditions (Fig. 6, compare C and E), which also alter P. putida motility (Figs 3 and S6C). While present at lower levels, multi-copy expression of YcgR-EYFP likewise showed a cytoplasmic location in P. putida (Fig. 6, compare E and F) and had a corresponding reduced effect on motility (Fig. S6C). This contrasts its punctate localization in E. coli, where it is expressed at similar levels as PP4397 (Fig. 6, compare F and H).

Taken together, the data in Figs 5 and 6 suggests that interaction between PP4397 (and likely YcgR) with the flagella motility apparatus of P. putida is weaker and/or more transient than that of YcgR with the motility apparatus of E. coli; and further, that a constant strict association with the flagella motor is not required for functionality. Although the interaction target of PP4397/FlgZ is unknown, based on the findings with the highly homologous FlgZ counterpart of P. aeruginosa23, it appears likely that one predominant target would be MotC and that functional replacement by YcgR relies on regions bearing common features between P. putida MotC and E. coli MotA proteins. Determining the interaction partner(s) for PP4397 is the subject of future studies.

Concluding Remarks

As for other bacteria, artificial increase of c-di-GMP levels by expression of native or heterologous DGCs results in reduced flagella-mediated motility in P. putida KT244012. Here we identify the PilZ domain containing PP4397/FlgZ protein as the effector relay protein that responds to elevated c-di-GMP levels resulting from lack of the signalling protein PP2258. Because P. putida harbours multiple c-di-GMP turnover proteins, it is likely that other c-di-GMP signalling pathways could also feed in to fine tune flagella performance through PP4397/FlgZ. Amongst the forty two P. putida c-di-GMP turnover proteins, PP2258 is the only one currently identified to possess both c-di-GMP degrading (PDE) and synthesising (DGC) activities12. However, the mechanism that controls the two opposing activities of PP2258 is unknown. One possibility is suggested by the genetic context of the pp2258 gene, which is located in a bicistronic operon downstream of aer1 that encodes a polar-localized receptor13. Both PP2258 and Aer1 possess PAS domains that are renowned for facilitating protein-protein interactions. Because the PAS domain of PP2258 is critical for its DGC activity12, it appears plausible that direct or indirect interaction between Aer1 and PP2258 could trigger a switch in its activities. Our current dissection of the signal transduction cascade from PP2258 to PP4397/FlgZ should greatly facilitate future work to determine if Aer1 controls PP2258 c-di-GMP signalling to ultimately control the ability of PP4397/FlgZ to act as an active hand-brake on the flagella motor.

Methods

Bacterial strains, growth conditions and general procedure

E. coli and P. putida strains (Table S1) were grown at 37 °C and 30 °C, respectively. E. coli DH530 was used for construction and maintenance of expression plasmids. The specialised replication-permissive E. coli S17λpir host, which expresses the Pir protein essential for replication of R6K31 was used for maintenance and conjugation of R6K-based suicide plasmids. P. putida strains used are all based on the genome sequenced KT244032 or a spontaneous streptomycin resistant derivative of KT2440 (KT270133). Plasmids (Table S2) were constructed by standard molecular techniques, as detailed in supporting information, and were introduced into P. putida by either electroporation or conjugation. Strains were cultured in Luria-Bertani (LB) broth (AppliChem GmbH) or on agar solidified plates supplemented with appropriate antibiotics. Concentrations used for E. coli were carbenicillin (Cb) 100 µg/ml, kanamycin (Km) 50 µg/ml, and tetracycline (Tc) 5 µg/ml, while those for P. putida were Cb 1 mg/ml, Km 50 µg/ml, and Tc 50 µg/ml.

PCR determination of the genome organisation of pp4397

Generation of cDNA from total RNA isolated from P. putida was as previously described13. After cDNA synthesis, mRNA was removed by 15 min incubation at 37 °C in the presence of 0.23 M NaOH and then neutralized by adding HEPES to a final concentration of 625 mM. The cDNA was subsequently buffer exchanged to 10 mM Tris-HCl (pH 8.5) using High Pure PCR product purification kit (Roche) before being subjected to PCR using the primer sets listed in Table S3 and depicted in Fig. 2B.

Generation of P. putida strains lacking PP4397

The pp4397 gene replacement cassette (Δpp4397::Tc) was introduced into the chromosome of P. putida KT2701 and its PP2258 null derivative13 via conjugation of pVI2299 (Table S2) from E. coli S17λpir and subsequent double-site recombination as previously described13. Growth in medium containing Tc and 10% sucrose was used to select for recombinants. Diagnostic PCR of the resulting strains was used to confirm loss of the native intact pp4397 gene and the presence of a fragment encompassing novel junctions of the Tc gene replacement and DNA upstream and downstream of the gene fragment of the suicide plasmid.

Generation of P. putida mono-copy chromosomal transcriptional and translational fusions

Fusions were introduced into the chromosome of P. putida strains via single site recombination as previously described13. Suicide plasmids carrying 3′-regions of target genes with cognate transcriptional fusions to either the promoter-less luxAB genes or in-frame translational fusions to eyfp, were introduced by conjugation as described above. Recombinants were selected using the antibiotic resistance marker(s) of the vector. Since the suicide plasmids carry only 3′-portions of the target genes, the resulting strains contain one functional (fused) copy and one inactive truncated copy of the gene separated by plasmid DNA. Diagnostic PCR was used to confirm correct recombination using primers homologous to DNA upstream of the gene fragment on the suicide plasmid and the DNA of the fusion partner.

In vivo luciferase transcriptional reporter assay

P. putida strains harboring mono-copy transcriptional fusions to luxAB were cultured in LB supplemented with appropriate antibiotics. To ensure balanced growth, overnight cultures were diluted in pre-warmed media and cultured into the exponential phase prior to a second dilution (to OD600 ~ 0.04) and initiation of the experiment. Growth and luciferase activity were monitored every 45 minutes for >9 hrs. Light emission was determined using 100 μl of culture after addition of decanal (1:2000 dilution) using a Infinite M200 (TECAN) luminometer.

In vitro transcription assays

Single-round transcription assays were performed at 30 °C using P. putida KT2440-derived core RNA polymerase (10 nM) and σFliA (40 nM) as previously described27 with 10 nM supercoiled pTE103-based plasmids as DNA templates (Table S2). Assays (20 μl) were performed in T-buffer (35 mM Tris-Ac pH 7.9, 70 mM KAc, 5 mM MgAc2, 20 mM NH4Ac, 1 mM DTT and 0.275 mg/ml BSA). For holoenzyme formation, core RNA polymerase and σFliA were pre-incubated for 5 minutes prior to addition of template DNA and a further 20 minutes incubation to allow open-complex formation. Transcription was initiated by the addition of NTPs (final concentration: ATP, 500 μM; GTP and CTP, 200 μM each; UTP, 80 μM and [α-32P]-UTP (5 μCi at >3,000 Ci/mmol) in the presence of heparin (0.1 mg/ml) to prevent re-initiation. After a further 10 minutes at 30 °C, reactions were terminated by adding 5 μl of a stop/load mix (150 mM EDTA, 1 M NaCl, 14 M urea, 3% glycerol, 0.075% (w/v) xylene cyanol, 0.075% (w/v) bromophenol blue) and transcripts analysed on 7 M urea/5% (w/v) polyacrylamide sequencing gels. Radioactivity was quantified using a Storm 860 imaging system (Molecular Dynamics).

Motility swimming and swarming plate assays

E. coli and P. putida strains were inoculated in LB supplemented with appropriate antibiotics and grown overnight. The next day, cultures were grown into early exponential phase, diluted to an OD600nm of 0.1 and grown once again for 5 hours. Cultures were then adjusted to an OD600nm of 0.3 and 5 μl were spotted on 0.3% soft agar LB plates for swimming assays and 0.5% agar LB plates for swarming assays. The resulting ring sizes were recorded after 6 h (E. coli) or 15 h (P. putida) of growth. Cells for western analysis were harvested at the same time as the dilutions prior to plating i.e. (after 5 hr of growth, OD600 = ~3.5).

Western analysis

Cell pellets were washed and resupended in ice-cold sonication buffer (20 mM Tris-HCl pH 7.5, 0.2 mM NaCl, 1 mM EDTA) containing protease inhibitors (Complete EDTA-free protease inhibitor tablet; Roche). Cells were disrupted by sonication and samples subsequently clarified by centrifugation. Protein concentrations of the resulting crude extracts were determined with PIERCE BCA protein assay (Thermo Scientific). Soluble protein samples were separated by 12% SDS-PAGE and transferred to PVDF membranes (Amersham Hybond-P) by electro-transfer. FLAG-tagged and EYFP-tagged proteins were detected using monoclonal mouse M2 anti-FLAG (Kodak) and anti-GFP (Invitrogen) antibodies, respectively. Antibody-decorated bands were revealed using polyclonal secondary goat anti-mouse antibodies conjugated with HRP and ECL Plus Western Blotting Reagents (GE Healthcare). Results were recorded using AGFA Curix Ultra UV-G medical X-ray film (Figs 4 and S4A) or LAS 4000 imaging system (Fujifilm; Figs 5, 6, S4B and S6).

Fluorescence microscopy

E. coli and P. putida strains were grown to exponential phase in LB, pelleted, washed and then fixed using paraformaldehyde (final concentration of 3%). Cells were adjusted to OD600nm of 0.8 in 50% PBS (68.5 mM NaCl, 1.35 mM KCl, 5 mM Na2HPO4, 0.9 mM KH2PO4; Applichem) containing 1 mg/ml BSA. Culture aliquots (3 µl) were air dried on glass slides prior to coating with Mowiol 4–88 (Calbiochem) mounting media (10% Mowiol 4–88 [w/v], 25% glycerol, 0.1 M Tris-HCl pH 8.5). Cells were imaged using an Eclipse 90i (Nikon) microscope equipped with a Hamamatsu ORCA-ER CCD camera, using oil immersion and a 100x objective with a numerical aperture of 1.30. Cells for western analysis were harvested at the same time as for fixing, i.e. mid exponential phase (OD600 = ~0.5 to 0.7), after 2 to 2.5 hours of growth.

Data availability statement

All data generated or analyzed during this study are included in this article and its Supplementary Information file.