The TonB_m-PocAB System Is Required for Maintenance of Membrane Integrity and Polar Position of Flagella in Pseudomonas putida

The ubiquitous Pseudomonas species are well adapted to survive in a wide variety of environments. Their success relies on their versatile metabolic, signaling, and transport ability but also on their high intrinsic tolerance to various stress factors. This is why the study of the stress-surviving mechanisms of Pseudomonas species is of utmost importance. The stress tolerance of Pseudomonads is mainly achieved through the high barrier property of their membranes. Here, we present evidence that the TonB-ExbBD-like TonB_m-PocAB system is involved in maintaining the membrane homeostasis of Pseudomonas putida, and its deficiency leads to lowered stress tolerance and conditional cell lysis.

ABSTRACT

TonB-ExbB-ExbD-like energy transduction systems are widespread among Gram-negative bacteria. While most species have only one copy of tonB-exbBD genes, the Pseudomonas species possess more TonB-ExbBD homologues. One of them, the TonB3-PocA-PocB complex, was recently shown to be required for polar localization of FlhF and, thus, the flagella in Pseudomonas aeruginosa. Here, we show that the orthologous TonB_m-PocA-PocB complex is important for polar localization of FlhF and flagella in Pseudomonas putida as well. Additionally, the system is necessary for maintaining membrane integrity, as the inactivation of the TonB_m-PocAB complex results in increased membrane permeability, lowered stress tolerance, and conditional cell lysis. Interestingly, the functionality of TonB_m-PocAB complex is more important for stationary than for exponentially growing bacteria. The whole-cell proteome analysis provided a likely explanation for this growth phase dependence, as extensive reprogramming was disclosed in an exponentially growing tonB_m deletion strain, while only a few proteomic changes, mostly downregulation of outer membrane proteins, were determined in the stationary-phase ΔtonB_m strain. We propose that this response in exponential phase, involving, inter alia, activation of AlgU and ColR regulons, can compensate for TonB_m-PocAB’s deficiency, while stationary-phase cells are unable to alleviate the lack of TonB_m-PocAB. Our results suggest that mislocalization of flagella does not cause the membrane integrity problems; rather, the impaired membrane intactness of the TonB_m-PocAB-deficient strain could be the reason for the random placement of flagella.

IMPORTANCE The ubiquitous Pseudomonas species are well adapted to survive in a wide variety of environments. Their success relies on their versatile metabolic, signaling, and transport ability but also on their high intrinsic tolerance to various stress factors. This is why the study of the stress-surviving mechanisms of Pseudomonas species is of utmost importance. The stress tolerance of Pseudomonads is mainly achieved through the high barrier property of their membranes. Here, we present evidence that the TonB-ExbBD-like TonB_m-PocAB system is involved in maintaining the membrane homeostasis of Pseudomonas putida, and its deficiency leads to lowered stress tolerance and conditional cell lysis.

INTRODUCTION

The cell envelope of Gram-negative bacteria contains two membranes separated by a periplasmic space (1). Due to cell envelope architecture, no energy is produced in the outer membrane, which means that transport proteins that require energy need to import it from the cytoplasmic membrane. The energy transfer is carried out by the TonB-ExbB-ExbD complex in the inner membrane that harnesses the proton motive force and directly interacts with the TonB-dependent transporters in the outer membrane (2, 3). Once thought to be restricted only to iron and vitamin B₁₂ transport, the TonB-ExbBD complex promotes the transport of a wide range of substrates that are too large or scarce to enter the cell by diffusion (4).

The TonB-ExbBD system is intensively studied in Escherichia coli, but the exact mechanism of its functioning in energy transduction has remained unclear. TonB and ExbD each have a transmembrane N terminus and are predominantly located in the periplasm, whereas ExbB has three transmembrane domains and a large cytoplasmic domain. ExbB forms a proton channel (5) and provides the structure for the complex (6). Depending on the membrane condition, ExbB proteins form either a pentameric or a hexameric channel with different conductance for protons (7). ExbD is predicted to be the carrier of protons during their translocation from periplasm to cytoplasm (8), but ExbD also interacts with TonB and appears to determine the right conformation of TonB during energy transmission (9, 10). TonB expands across the periplasm and transduces the harvested energy to the outer membrane transporters (11, 12) that have a TonB box domain. This domain is proposed to extend into the periplasm and become available for interaction with TonB after the binding of substrate to the transporter (13).

The TonB-ExbBD complexes are widespread among Gram-negative bacteria. Most species, including E. coli, have one copy of tonB-exbBD genes, but the number of tonB homologues can vary by up to nine per genome (14). The ubiquitous soil and rhizosphere bacterium Pseudomonas putida possesses two TonB homologues encoded by PP_4994 and PP_5308. PP_5308 (tonB) is in an operon with exbB and exbD, and their corresponding proteins appear to have a function similar to that of their E. coli homologs, as the P. putida strains defective in tonB-exbBD genes are impaired in siderophore transport and deficient in iron acquisition (15, 16). Moreover, the exbB of P. putida can complement the lack of exbB in E. coli (15). Besides its importance in iron transport, the TonB-ExbBD complex is required for P. putida for the tolerance of several antibiotics, p‐hydroxybenzoate, and toluene (16, 17) and affects its fitness in colonizing corn seeds and roots (18).

PP_4994 has not been studied in P. putida, but its conserved homologues can be found in most Pseudomonas species (19). The orthologue of PP_4994 in P. aeruginosa, TonB3, likely forms an inner membrane-associated complex with homologues of ExbB and ExbD, named PocA and PocB, respectively (20). Interestingly, P. aeruginosa’s TonB3-PocAB complex does not appear to have a role in promoting iron transport (21). Instead, it is involved in swimming and twitching ability (20, 22). The TonB3-PocAB complex, although not polarly localized itself, is needed for the polar localization of flagella and type IV pili (20). Deleting any one of the tonB3-pocAB genes in P. aeruginosa reduces swimming due to the random localization of flagella, which is caused by the random localization of otherwise polarly localized FlhF protein.

In this study, we investigate the role of PP_4994-encoded TonB and PP_1898-1899-encoded PocAB in P. putida. We show that like its orthologue in P. aeruginosa, the TonB-PocAB complex of P. putida is needed for correct placement of flagella and FlhF. For this shared role in motility, we name the PP_4994-encoded protein TonB_m. Interestingly, our results also indicate that the TonB_m-PocAB complex has a separate task in maintaining membrane integrity. The TonB_m-PocAB-deficient strains have permeable membranes and are sensitive to several chemicals. Curiously, the effects of TonB_m-PocAB’s deficiency depend on the physiological state of the inoculum as bacteria originating from stationary phase are significantly more compromised than the exponentially growing TonB_m-PocAB knockout mutants. The comparison of whole-cell proteomes of the ΔtonB_m strain and wild-type P. putida revealed extensive proteomic changes occurring in the exponentially growing but not in the stationary-phase ΔtonB_m strain. We hypothesize that the observed changes compensate for the lack of tonB_m in exponentially growing cells.

RESULTS

Inactivation of the TonB_m-PocAB complex decreases swimming and results in membrane defects.

The ability of bacteria to bind Congo red (CR) dye has been used as a marker of membrane defects and cell lysis (23). Thus, we screened a transposon mutant library of the P. putida wild-type strain for CR binding mutants to find genes that could be important in membrane homeostasis. In accordance with previous studies (23, 24), the current screen repeatedly detected the CR binding mutants with transposon insertion into the colRS operon. ColR is the response regulator of the ColRS two-component regulatory system that responds to the excess of metals, such as zinc, iron, manganese, and cadmium (25). In the absence of this system, P. putida has problems with maintaining cell membrane integrity not only in the presence of excess metals (25, 26) but also on glucose minimal medium (23, 24). The second most common cause of pink colonies was disruption of PP_4994, which codes for an orthologue of P. aeruginosa’s TonB3. For one pink colony, the transposon had disrupted PP_1898, a probable orthologue of P. aeruginosa’s pocA gene (19). Identifying orthologues of genes that regulate flagellum localization in P. aeruginosa (20) in our membrane stress screen was intriguing, because assuming that PP_4994 and PocA of P. putida have functions similar to those of their orthologues, it is unclear why wrong localization of flagella and decreased motility result in membrane stress (23, 24).

To get further insight into the role(s) of PP_4994 and PocAB in P. putida, the PP_4994 (tonB_m), PP_1898 (pocA), and PP_1899 (pocB) single deletion strains as well as the whole TonB_m-PocAB complex deletion strain (ΔtonB_m ΔpocAB) were constructed. As the decreased swimming ability of P. aeruginosa in the absence of the TonB3-PocAB system is caused by the nonpolar localization of FlhF (20), a protein that determines the place of flagellum formation (27), the flhF (PP_4343) deletion strain was constructed as well.

To test if tonB_m, pocA, and pocB are required for motility in P. putida, the swimming ability of all the deletion strains was analyzed. Compared to the ΔflhF strain, which had a severe motility defect, the swimming ability of ΔtonB_m, ΔpocA, ΔpocB, and ΔtonB_m ΔpocAB strains was less affected but nevertheless clearly inhibited (Fig. 1A). The ΔtonB_m ΔflhF double deletion strain displayed swimming defects similar to that of the ΔflhF strain, which is in good correlation with FlhF lying downstream of the TonB3-PocAB system in regulation of flagellum localization in P. aeruginosa (20). Complementation of the ΔtonB_m strain with ectopically expressed tonB_m restored normal motility (Fig. 1A).

FIG 1.

Effect of TonB_m-PocAB system’s deficiency in swimming ability (A), flagellum localization (B and C), FlhF positioning (D), and Congo red binding (E). The cells of P. putida wild type (wt), tonB_m, pocA, pocB, and flhF single deletion strains, tonB_m pocAB triple deletion strain, tonB_m flhF double deletion strain, and tonB_m deletion strain complemented with the tonB_m gene (tac tonB_m) and wild-type, ΔtonB_m, ΔflhF, and ΔtonB_m ΔpocAB strains carrying the flhF-gfp fusion under the control of the inducible P_tac promoter were grown at 30°C. The expression of TonB_m and FlhF-GFP was induced with 0.5 mM IPTG. (A) Swimming ability. The cells were grown on LB medium containing 0.3% agarose for 18 h. (B) Localization of flagella. Exponentially growing cells (OD₅₈₀ of ∼0.5) were stained with silver and examined using oil immersion light microscopy. (C) Quantification of flagellum localization. Flagellum localization was considered random (nonpolar) if at least one flagellum deviated from polar positioning. Relative proportions of cells with polar and nonpolar flagella (means with 95% confidence intervals from at least four independent experiments) are presented. Two hundred to 800 cells were examined for each strain. Statistically significant differences from the wild type (*, P < 0.001) and difference from the uninduced tac tonB_m strain (¤, P = 0.000003) are indicated. (D) Localization of FlhF-GFP. The cells were grown overnight in LB medium supplemented with IPTG and examined using oil immersion light microscopy. (E) Congo red (CR) binding. The cells were grown on glucose minimal medium supplemented with 0.0005% CR for 72 h.

To determine if the swimming defect of TonB_m-PocAB- and FlhF-deficient strains is caused by incorrect localization of flagella, the position of flagella was ascertained by microscopy. More than 90% of exponentially growing wild-type P. putida cells had all of their flagella (1 to 5 flagella) located at the center of the cell pole (Fig. 1B and C). However, in the TonB_m-PocAB single and triple deletion strains, the uniform polar placement of flagella was lost and approximately 80% to 90% of cells had at least one incorrectly placed flagellum (Fig. 1B and C). The ΔflhF and ΔtonB_m ΔflhF mutants had a random distribution of flagella similar to that of TonB_m-PocAB-deficient strains (Fig. 1C), and their number of flagella per cell was decreased. Already without isopropyl-β-d-thiogalactopyranoside (IPTG) induction, the tonB_m complementation strain showed more cells with only polar flagella (35%), which indicates that the tac promoter in the lacI^q-P_tac-tonB_m cassette was leaky. When the expression of TonB_m was induced with 0.5 mM IPTG, about 80% of the cells of the tonB_m complementation strain had polar flagella (Fig. 1C). To analyze the localization of FlhF, the flhF-gfp translational fusion was constructed and introduced into the chromosome of P. putida wild-type, ΔtonB_m, ΔflhF, and ΔtonB_m ΔflhF strains. Fluorescence microscopy showed that FlhF-green fluorescent protein (FlhF-GFP) is polar in wild-type and ΔflhF strains but locates randomly in ΔtonB_m and ΔtonB_m ΔflhF strains (Fig. 1D). We also observed that the FlhF-GFP foci were brighter in wild-type and ΔflhF bacteria than in the tonB_m-deficient bacteria, where more FlhF-GFP seemed to localize in the cytoplasm. Thus, our data indicate that the P. putida TonB_m-PocAB system has a role similar to that of TonB3-PocAB in P. aeruginosa.

To investigate whether the motility defect and random localization of flagella correlate with the CR-binding phenotype, the motility-deficient strains were tested on glucose medium supplemented with CR. Interestingly, while strains with the TonB_m-PocAB-deficient system stained pink, the ΔflhF strain resembled wild-type P. putida (Fig. 1E). The control experiment with the tonB_m complementation strain revealed that the CR binding phenotype of the ΔtonB_m strain is suppressed by the overexpression of TonB_m. These data show that the motility defect and wrong localization of flagella do not account for the CR binding of the ΔtonB_m, ΔpocA, and ΔpocB strains and that in the absence of TonB_m-PocAB system, cells must have additional problems not related to the misplacement of flagella.

The absence of TonB_m-PocAB complex results in glucose-specific cell lysis.

Given that the Congo red binding indicates a severe membrane defect (23, 24), we next tested the motility mutants for membrane leakage. We introduced the β-galactosidase expression plasmid into the wild-type and mutant strains and measured the β-galactosidase activity from the supernatant of glucose-grown bacteria. High extracellular β-galactosidase activity was detected in the case of ΔtonB_m, ΔpocA, ΔpocB, ΔtonB_m ΔpocAB, and ΔtonB_m ΔflhF strains, indicating cell lysis (Fig. 2A). Contrary to this finding, the ΔflhF mutant displayed no β-galactosidase leakage.

FIG 2.

TonB_m-PocAB-deficient strains have increased membrane permeability. (A) β-Galactosidase activities measured from the supernatant of glucose- and LB-grown P. putida wt, tonB_m, pocA, pocB, and flhF single deletion, tonB_m pocAB triple deletion, and tonB_m flhF double deletion strains. All strains carry pKTlacZS/C plasmid. Data (means with 95% confidence intervals) from at least three independent experiments are presented. (B) Flow cytometry analysis of glucose- and LB-grown bacteria stained with SYTO9 and propidium iodide (PI). Relative proportions of the subpopulations of intact, PI-permeable, and dead cells (means with 95% confidence intervals) from at least four independent determinations are presented. (C) β-Galactosidase activities measured from the supernatant of glucose-grown P. putida wt, tonB_m deletion, oprB1-deficient tonB_m, and pocA, pocB, flhF, and tonB_m flhF deletion strains. All strains carry pKTlacZS/C plasmid. Data (means with 95% confidence intervals) from at least three independent experiments are presented.

In order to test whether the lysis of the ΔtonB_m, ΔpocA, and ΔpocB strains is dependent on the growth medium, the β-galactosidase leakage assay was performed with lysogeny broth (LB)-grown bacteria as well. However, as neither of the LB-grown strains differed from the wild type (Fig. 2A), the cell lysis of ΔtonB_m, ΔpocA, and ΔpocB strains seems to be specific to growth on glucose.

Previous results with colR-deficient P. putida have shown that glucose-specific cell lysis is a subpopulation phenotype (24, 28). Therefore, to get insight into the population structure of TonB_m-PocAB-deficient strains, flow cytometry analysis of bacteria stained with SYTO9 and propidium iodide (PI) was performed. Three populations could be detected: (i) undamaged cells that stained with SYTO9 only; (ii) cells that stained with both SYTO9 and PI (indicated as PI-permeable); and (iii) cells that stained with PI and SYTO9 but had lower side scatter. The latter subpopulation has been shown to correlate with cell lysis and therefore has been defined as dead cells (28). The single-cell analysis of the glucose solid medium-growing bacteria revealed that colonies of the ΔtonB_m, ΔpocA, ΔpocB, and ΔtonB_m ΔflhF strains contained more PI-permeable and dead cells than the wild type, whereas the deletion of flhF did not influence either the membrane permeability or death of bacteria (Fig. 2B). Interestingly, analysis of the mutants grown on LB solid medium also showed that the ΔtonB_m, ΔpocA, ΔpocB, and ΔtonB_m ΔflhF colonies contain more PI-permeable cells, but the amount of dead cells was similar to that of the wild type (Fig. 2B). The LB-grown ΔflhF strain had no difference from the wild type. These results show that while the deficiency in the TonB_m-PocAB system increases membrane permeability regardless of the growth medium, the membrane damage is more pronounced on the glucose minimal medium. The unaffected membrane permeability of the ΔflhF strain indicates that wrong placement of flagella does not affect membrane permeability or cause cell lysis.

The inability to tolerate the increased expression of the sugar channel protein OprB1 in the outer membrane has been described as the reason for glucose-specific cell lysis in the colR-deficient mutant (23). To test if the glucose-induced OprB1 could be the reason for cell lysis of glucose-grown ΔtonB_m, ΔpocA, ΔpocB, and ΔtonB_m ΔflhF cells as well, oprB1-deficient derivatives were constructed from the deletion strains and β-galactosidase leakage was analyzed. Data shown in Fig. 2C indicated that disruption of oprB1 eliminated the glucose-dependent cell lysis of TonB_m-PocAB-deficient strains. This indicated that, similar to the colR-deficient mutant, glucose-induced expression of OprB1 was involved in the glucose-specific cell lysis of TonB_m-PocAB mutants.

TonB_m-PocAB system affects stress tolerance and generation time in a growth phase-dependent manner.

Since the flow cytometry analysis revealed that the TonB_m-PocAB system contributes to membrane integrity, we hypothesized that the stress tolerance of these strains is affected. To test this, the growth of the ΔtonB_m, ΔpocA, and ΔpocB strains and wild-type P. putida was compared on LB medium supplemented with different chemicals: metal salts (LiCl, NaCl, MnSO₄, FeSO₄, CoCl₂, NiSO₄, CuSO₄, ZnSO₄, RbCl, CdSO₄, and K₂Cr₂O₂), antibiotics (colistin, polymyxin B, benzylpenicillin, tetracycline, and rifampin), DNA-damaging chemicals (mitomycin C and 4-nitroquinoline 1-oxide), compounds producing reactive oxygen species (paraquat and H₂O₂), and EDTA. The ΔflhF strain was used as a control to rule out the effect of mislocalization of flagella on stress tolerance. During the experiments we observed that the results were dependent on the age of the inoculum; thus, the stress tolerance of cells from both stationary and exponential growth phases was analyzed. When the exponential-phase cells were tested, difference in growth could be observed only on media containing MnSO₄ or 4-nitroquinoline 1-oxide (nitroquinoline) (Fig. 3A). However, when inoculated cells originated from the stationary growth phase, the TonB_m-PocAB deletion strains revealed increased sensitivity to CuSO₄, MnSO₄, colistin, benzylpenicillin, nitroquinoline, and mitomycin C (Fig. 3B), and their growth was slightly decreased by ZnSO₄, NaCl, and EDTA (data not shown). These results indicated that the TonB_m-PocAB complex affected P. putida’s stress tolerance, but the magnitude of the effect depended extensively on the growth phase. Given that the ΔflhF strain resembled that of the wild type under all conditions, the increased stress sensitivity could not be caused by mislocalization of flagella. Although the TonB_m-PocAB system was not involved in iron tolerance, we also investigated the growth of mutants under iron-limited conditions, but as no differences from the wild-type were detected (data not shown), the TonB_m-PocAB complex is likely not involved in iron acquisition.

FIG 3.

TonB_m-PocAB system affects stress tolerance and generation time in a growth phase-dependent manner. (A and B) Stress tolerance of exponential-phase (A) and stationary-phase (B) P. putida wt and tonB_m, pocA, pocB, and flhF deletion strains. Exponential-phase cells were grown for 3 h (OD₅₈₀ of ∼1) and stationary-phase cells overnight before inoculating the strains onto LB medium supplemented with different chemicals. The cells were then grown for 20 h at 30°C. Approximately 50,000, 5,000, 500, and 50 cells were inoculated per spot. Bp, benzylpenicillin; NQ, nitroquinoline; MMC, mitomycin C. (C and D) Growth curves and minimal generation times of bacterial cultures inoculated either with exponential-phase (C) or stationary-phase (D) cells. P. putida wt, tonB_m, pocA, and pocB deletion strains and colR-deficient and tonB_m colR double deficient strains were grown for 3 h (exponential phase) or overnight (stationary phase) before inoculating the cells into LB medium. The strains were grown at 30°C on a microtiter plate. Means from eight parallels of one measurement with 95% confidence intervals are presented. Average minimal generation times (mgt) from three independent measurements for each strain also are shown. Statistically significant differences from the wild type are indicated (*, P = 0.012; **, P < 0.0001).

To further analyze the growth phase effects of TonB_m-PocAB-deficient P. putida, the LB medium was inoculated with bacteria of different ages and growth curves were recorded. While the maximum growth rate of the wild type remained the same independent of the inoculum used, the growth rates of TonB_m-PocAB-deficient strains were clearly affected by the growth stage of the inoculum. When exponential-phase cells were used for inoculation, the growth curves and minimal generation times of ΔtonB_m, ΔpocA, and ΔpocB strains resembled those of the wild type (Fig. 3C). However, when the culture was started with stationary-phase cells, the minimal generation time for the ΔtonB_m and ΔpocAB strains was, on average, about 10 min longer (Fig. 3D) and had a statistically significant difference from the wild type. This furthermore demonstrates that the effect of TonB_m-PocAB deficiency depends on the age of the P. putida cells.

MexCD-OprJ efflux system is downregulated in ΔtonB_m strain in stationary phase.

Considering that the TonB_m-PocAB system’s homologue in E. coli, the TonB-ExbBD complex, connects the two membranes by mediating the energy of cytoplasmic membrane to outer membrane transporters (2), we next analyzed different membrane fractions of the TonB_m-PocAB-deficient strains from both exponential and stationary growth phases. While the patterns of lipopolysaccharides and periplasmic and cytoplasmic membrane proteins of wild-type and TonB_m-PocAB deletion strains were similar (data not shown), the outer membrane protein (OMP) pattern of mutants revealed noticeable differences from the wild type (Fig. 4).

FIG 4.

The absence of intact TonB_m-PocAB complex causes changes in the outer membrane protein pattern. Outer membrane proteins were prepared from P. putida wt and tonB_m and pocA deletion strains, separated by SDS-PAGE, and visualized by silver staining. Cells were grown in LB medium for 3 h (OD₅₈₀ of ∼1; exponential phase) or 17 h (stationary phase). All lanes contain 1 μg of protein. Major differences between wild-type and tonB_m-pocAB-deficient strains are indicated by arrows.

To get more detailed insight into the protein profile of the TonB_m-deficient strain, a whole-cell proteome analysis of exponential- and stationary-phase ΔtonB_m and wild-type P. putida cells was performed. Extensive growth phase-dependent proteome rearrangements were detected in both strains: 1,426 proteins out of 2,652 in the wild-type strain (see Table S1 in the supplemental material) and 1,326 proteins out of 2,673 in the ΔtonB_m strain (Table S2) had a statistically significant and at least 2-fold difference between the growth phases. Interestingly, when the stationary-phase data of the wild type was compared to the corresponding ΔtonB_m data, only six proteins were found to be significantly differentially expressed (Fig. 5 and Table S3). Besides these six proteins, we also considered nine differentially expressed proteins that were present in all the samples of one strain but were not detectable in any of the parallels of the other (so-called on-off-regulated proteins) and had at least a 2-fold difference in their expression level when imputed values were used. Notably, 11 of these 15 were membrane proteins, most of them being downregulated in the ΔtonB_m strain (Fig. 5 and Table S3). The greatest differences were the 20-, 82-, and 65-fold decreases in the amount of MexC, MexD, and OprJ, respectively, which together form a multidrug efflux system that exports several antimicrobial agents in P. aeruginosa (29). The level of PhaK, an outer membrane channel protein that facilitates the uptake of phenylacetic acid (30), had dropped about 9-fold. This is probably caused by the 4-fold increased level of PhaN, a transcriptional repressor of the phenylacetic acid pathway (30).

FIG 5.

Volcano plot showing differences in protein expression between stationary-phase tonB_m-deficient P. putida and the wild type. Filled symbols represent 15 proteins considered differently expressed in the ΔtonB_m strain. Diamonds indicate on-off-regulated proteins. Proteins located above the horizontal line have statistically significant changes in their expression values.

In wild-type cells, MexCD-OprJ efflux pump components are upregulated in stationary phase (Table S1), whereas in the ΔtonB_m strain, MexC is clearly downregulated and MexD and OprJ are not even detectable in stationary phase (Tables S2 and S3). This remarkable downregulation of MexCD-OprJ efflux pump proteins in the ΔtonB_m strain led us to hypothesize that it was the reason for the decreased stress tolerance of the ΔtonB_m strain. To test this possibility, we deleted the whole mexCD-oprJ operon from both wild-type and ΔtonB_m strains and tested stress tolerance. The deletion of mexCD-oprJ had no effect on the tolerance of CuSO₄, MnSO₄, nitroquinoline, colistin, benzylpenicillin (Fig. 6), polymyxin B, and mitomycin C (data not shown) in either of the strains, suggesting that MexCD-OprJ did not contribute to the efflux of these compounds and that the downregulation of MexCD-OprJ complex was not the cause of elevated stress sensitivity of the ΔtonB_m strain. On the other hand, if the MexCD-OprJ complex was related to the stress tolerance of the ΔtonB_m strain, then its overexpression should have alleviated the increased sensitivity.

FIG 6.

MexCD-OprJ effux pump is not related to the stress susceptibility of tonB_m deletion strain. Overnight-grown P. putida wt, mexCD-oprJ, tonB_m, and nfxB deletion strains and tonB_m mexCD-oprJ and tonB_m nfxB double deletion strains were grown on LB medium supplemented with different chemicals for 20 h at 30°C. Approximately 50,000, 5,000, 500, and 50 cells were inoculated per spot.

In P. aeruginosa, the expression of the mexCD-oprJ operon is controlled by two repressors, NfxB (31) and EsrC (32). The loss of NfxB leads to overexpression of mexCD-oprJ genes (33), whereas EsrC is active as a repressor only in the presence of NfxB and its effect on mexCD-oprJ is modest compared to that of NfxB (32). The homologue of NfxB in P. putida is PP_2820, and, expecting its absence to raise the expression of mexCD-oprJ, we constructed ΔnfxB (PP_2820) deletion strains of wild-type and ΔtonB_m strains. In P. aeruginosa, the upregulation of MexCD-OprJ due to the loss of NfxB results in increased ciprofloxacin resistance (34). In accordance with that, the ciprofloxacin resistance of the ΔnfxB and ΔtonB_m ΔnfxB strains was considerably higher than that of their parent strains (Fig. 6), suggesting that MexCD-OprJ was upregulated in the P. putida ΔnfxB strain. To assess if increased expression of MexCD-OprJ would relieve the reduced stress tolerance of the ΔtonB_m strain, we next tested the ΔnfxB strain tolerance to several compounds. The results revealed that while the lack of nfxB decreased the benzylpenicillin tolerance of both wild-type and ΔtonB_m strains (Fig. 6), it did not influence the tolerance to other chemicals. These results support the contention that low levels of MexCD-OprJ are not sufficient to explain the decreased stress tolerance of the TonB_m-PocAB-deficient strain.

Proteome analysis revealed extensive changes in exponentially growing ΔtonB_m cells.

Contrary to the few differences in stationary phase, the proteome analysis of exponential-phase cells showed at least a 2-fold change of 126 proteins in the ΔtonB_m strain compared to level for the wild type (Fig. 7 and Table S4). One hundred eight of these changes were statistically significant, but we also included 18 on-off proteins (Table S4). It is remarkable that most of the proteins, 102 out of 126, were upregulated in the ΔtonB_m strain (Fig. 7 and Table S4). Only the outer membrane proteins that predominantly belonged to the transport and secretion category were mainly downregulated. About a third of all the changed proteins were associated with either amino acid and protein metabolism (17 proteins) or with general metabolism (22 proteins), suggesting that a considerable metabolic reprogramming was induced in the exponentially growing ΔtonB_m strain. Another large group comprised 29 proteins related to transport and secretion. Half of these proteins were downregulated in the ΔtonB_m strain and half, including 10 proteins of the type VI secretion system K1 (35), were upregulated (Table S4). It is also noteworthy that 13 proteins associated with stress and defense responses and 10 transcriptional regulators or histidine kinases were responding to TonB_m deficiency, and again, most of them were upregulated. Surprisingly, only five proteins (OprQ, OpdP, TolC, PP_0765, and PP_4978) overlapped the differentially expressed proteins of stationary phase.

FIG 7.

Volcano plot showing differences in protein expression between exponentially growing tonB_m-deficient P. putida and the wild type. Filled symbols represent 126 proteins considered differently expressed in the ΔtonB_m strain. Diamonds indicate on-off proteins. Proteins belonging to the AlgU regulon are shown in blue, and proteins regulated by ColR are in red. A name tag indicates that the protein was also differently expressed in the zinc-exposed colR-deficient strain or is ColR regulated (also in red). Proteins located above the horizontal line have statistically significant changes in their expression values.

AlgU and ColR regulons are activated in ΔtonB_m strain.

The proteome data of exponential-phase bacteria revealed that several alginate biosynthesis proteins as well as the alginate regulator AmrZ were upregulated in the ΔtonB_m strain (Fig. 7 and Table S4). These results were verified by analysis of amrZ-lacZ and algD-lacZ transcriptional fusions in the ΔtonB_m strain, showing an increased promoter activity (Fig. 8A and B). Alginate is an exopolysaccharide that contributes to biofilm formation and stress tolerance under water-limiting conditions in P. putida (36). In P. aeruginosa, alginate is produced in response to cell wall stress due to the activation of sigma factor AlgU (37). Analysis of the proteome response of the ΔtonB_m strain revealed that 31 out of 126 differentially expressed proteins were orthologues of the P. aeruginosa AlgU regulon (Fig. 7, marked with blue) (37). This suggests that the AlgU regulon is activated in the exponentially growing ΔtonB_m strain.

FIG 8.

AlgU and ColR regulon genes respond to tonB_m deficiency. (A and B) β-Galactosidase activities measured in P. putida wt and tonB_m deletion strains carrying the amrZ or algD transcriptional fusion with lacZ in the plasmid p9TT_BlacZ. (C) β-Galactosidase activities measured in P. putida wt, tonB_m deletion, and colR-deficient tonB_m deletion strains carrying the oprQ, PP_0903, PP_5152, or PP_0900 transcriptional fusion with lacZ. Bacteria were grown in LB medium for 3 h (OD₅₈₀ of ∼1) at 30°C. Data (means with 95% confidence intervals) from at least four independent experiments are presented.

Besides activation of the AlgU regulon, three proteins of the ColR regulon, OprQ, PP_0903, and PP_5151 (25), responded to TonB_m deficiency (Fig. 7, indicated in red) in a direction indicating that the ColRS signaling is active in the ΔtonB_m strain. To test this possibility, the ColR-responsive oprQ-lacZ, PP_0903-lacZ, and PP_5152-lacZ transcriptional fusions (PP_5152 is the first gene in the two-gene PP_5152-PP_5151 operon) were analyzed in exponentially grown wild-type, ΔtonB_m, and ΔtonB_m colR double mutant strains (Fig. 8C). The expression of oprQ is known to be repressed, and the expression of PP_0903 and PP_5152 activated by ColR (25, 38). In accordance with that and verifying the changes seen in the proteome data (Fig. 7), the promoter activity of oprQ was lower and the activities of PP_0903 and PP_5152 were higher in the ΔtonB_m than the wild-type strain (Fig. 8C). Given that none of the three promoters responded to the lack of tonB_m in the ΔtonB_m colR strain (Fig. 8C), ColR is clearly responsible for the altered expression of those genes in the ΔtonB_m strain. To further confirm if ColRS signaling is active in TonB_m-PocAB-deficient cells, the expression of the ColR-responsive PP_0900, which was not detectable in the proteome, was also analyzed. Corroborating the prior results, the PP_0900-lacZ fusion was induced in TonB_m-deficient cells in a ColR-dependent manner (Fig. 8C), confirming the activation of the ColRS system in response to TonB_m-PocAB deficiency.

Activation of the ColR regulon is beneficial for the ΔtonB_m strain.

The proteome of colR-deficient P. putida recently has been analyzed (26), and the comparison of these previous data to the current data revealed a remarkable overlap between the proteome responses of exponentially grown ΔtonB_m and zinc-stressed colR mutant strains (26). The same 61 proteins, including, for example, several transport and stress response proteins, were differentially expressed in both mutant strains, and all in the same manner (Fig. 7 [the overlapping 61 proteins are indicated with name tags] and Table S4). Considering this partial overlap of the proteome responses and several shared membrane stress-indicating phenotypes (glucose-induced Congo red binding and cell lysis) of ΔtonB_m and colR mutant cells (Fig. 1 and 2) (23), it was hypothesized that the activation of ColRS signaling could alleviate the membrane damage of the ΔtonB_m mutant. To test this, the growth parameters and stress tolerance of the ΔtonB_m colR double mutant was analyzed. The growth curve of the combined mutant in LB medium displayed a prolonged lag phase that was particularly apparent when stationary-phase cells were used as an inoculum (Fig. 3C and D). The minimal generation time of the ΔtonB_m colR strain was, on average, about 5 min longer than that of other strains when exponential-phase cells were analyzed (Fig. 3C). The growth rate reduction of the double mutant was even more pronounced when stationary-phase inocula were tested (Fig. 3D). The ΔtonB_m colR double mutant was also more compromised in some stress tolerance tests. Exponential-phase ΔtonB_m colR cells were less tolerant to MnCl₂ than the tonB_m single mutant (Fig. 9A), and the stationary-phase ΔtonB_m colR double mutant tolerated less CuSO₄, MnCl₂, and benzylpenicillin than the ΔtonB_m strain (Fig. 9B). Given that colR deficiency clearly intensifies the phenotypes of the ΔtonB_m strain, the activation of the ColR regulon seems to be beneficial to P. putida lacking TonB_m. The accumulation of the effects of tonB_m and colR deficiency suggests that the origin of the problems in these strains is different, which implies that although the lack of TonB_m-PocAB and ColRS produces similar responses, the two systems operate in separate regulatory pathways.

FIG 9.

Stress tolerance of exponential-phase (A) and stationary-phase (B) P. putida wt, tonB_m deletion, colR-deficient tonB_m deletion, and colR-deficient strains. Exponential-phase cells were grown for 3 h (OD₅₈₀ of ∼1) and stationary-phase cells overnight before inoculating the strains onto LB medium supplemented with different chemicals. The cells were then grown for 20 h at 30°C. Approximately 50,000, 5,000, 500, and 50 stationary-phase cells were inoculated per spot.

DISCUSSION

Prior to the current work, the homologue of the TonB_m-PocAB complex, TonB3-PocAB, was shown to be required for swimming and twitching in P. aeruginosa (20, 22). In the absence of tonB3, pocA, or pocB, the swimming ability of P. aeruginosa is impaired because the polar localization of flagella is lost due to the random positioning of FlhF (20), which marks the assembly point of new flagella (27). The present study suggests that the TonB-PocAB complex has the same function in other Pseudomonas species, as P. putida tonB_m, pocA, and pocB deletion strains have randomly distributed flagella and decreased swimming ability as well. However, our results indicate that besides the role in motility, the TonB_m-PocAB complex is needed for the maintenance of membrane integrity. We show that the misplacement of flagella is not responsible for the membrane defects of TonB_m-PocAB mutants and rather suggests that the impaired membrane homeostasis is the main cause for the random localization of flagella as well as for other deficiencies.

The subpopulation lysis of TonB_m-PocAB mutants on glucose medium, evidenced by the leakage of cytoplasmic β-galactosidase (Fig. 2A), clearly shows that these mutants have a fragile membrane. The finding that glucose-specific cell lysis was abolished by the deletion of oprB1 indicates that TonB_m-PocAB-deficient strains cannot tolerate OprB1 porin. OprB1 is a carbohydrate-selective porin that has a relatively high affinity for glucose (39) and is significantly upregulated when glucose becomes limiting (23). In nutrient-rich LB medium, glucose transport is inhibited, as Pseudomonas species prefer to use organic acids and amino acids as a carbon source instead (40, 41). A similar glucose-specific and OprB1-dependent cell lysis has been previously described for a P. putida colR-deficient strain (23, 24). The lysis of the colR mutant was shown to result from the intolerance of the accumulation of OprB1 in the outer membrane, the cell’s normal response to glucose limitation (23). Given that the TonB_m-PocAB-deficient strains have similar hypersensitivities to the OprB1 porin, we conclude that the outer membrane of the TonB_m-PocAB mutants is compromised. Furthermore, single-cell analysis revealed that, independent of the carbon source, the populations of TonB_m-PocAB mutants contain significantly more PI-permeable cells than the wild type (Fig. 2B). As propidium iodide can pass through undamaged outer membrane via porins but cannot cross intact cytoplasmic membrane (42), the increased permeability to PI implies that not only the outer membrane but also the inner membrane of the ΔtonB_m mutant is deficient.

The stress tolerance assays suggested that the fitness effects of TonB_m-PocAB’s disruption were more pronounced when stationary-phase bacteria were analyzed (Fig. 3A and B). Considering that the growth characteristics of the ΔtonB_m strain also depended on the growth phase of the inoculum under unstressed conditions (Fig. 3C and D), the TonB_m-PocAB complex seems to be more important for stationary- than for exponential-phase bacteria. The proteome analysis provided a likely explanation for this growth phase dependence, namely, extensive differences, including upregulation of many metabolism-related stress response and regulatory proteins, were observed in the exponentially growing ΔtonB_m strain, while only a few proteomic changes were detected in the stationary-phase ΔtonB_m strain. This suggests that exponentially growing cells are actively dealing with the stress caused by the absence of functional TonB_m-PocAB, while stationary-phase cells lack this reprogramming, perhaps due to energy limitation, and cannot cope with the effects of TonB_m-PocAB deficiency. Thus, we assume that the ability of exponentially growing cells to compensate for the absence of the TonB_m-PocAB system allows them to preserve the wild-type-like growth rate in rich medium and start growth in the presence of stress.

Besides growth phase differences, the proteome data confirmed that the TonB_m-PocAB complex is necessary for maintaining membrane integrity. First, most of the proteins with changed expression in stationary-phase ΔtonB_m cells were membrane proteins that were upregulated in the wild-type but not in the ΔtonB_m strain. This hints that the cell envelope of the TonB_m-PocAB-deficient strain is sensitive to membrane protein upregulation, somewhat analogously to the sensitivity of OprB1 in glucose medium. Second, as the expression of 31 AlgU regulon proteins was found to be changed in the exponentially growing ΔtonB_m strain, AlgU seems to be activated in the ΔtonB_m strain. AlgU is an envelope stress response sigma factor that controls the expression of large numbers of genes (37). Under normal growth conditions, AlgU is bound to the anti-sigma factor MucA, which anchors it to the cytoplasmic membrane and prohibits it from regulating gene expression (43). Under cell envelope stress, induced either by cell wall-acting antibiotics or other compounds that disrupt bacterial membranes (37, 44) or by the overexpression of certain outer membrane proteins (43), MucA is degraded and AlgU is released into the cytoplasm (37). Therefore, the activation of the AlgU regulon indicates that the TonB_m-PocAB-deficient cells must experience envelope stress. Third, a large part of the differentially expressed proteins in exponentially growing ΔtonB_m strain (61 out of 126) overlapped the proteome response previously observed in colR-deficient P. putida treated with ZnSO₄ (26). This included many AlgU-regulated and stress-related proteins, the expression of which could be explained by the activation of AlgU in both strains, but also several other proteins, such as pyoverdine synthesis and type VI secretion system proteins. Analogously to AlgU regulon genes, the pyoverdine genes in P. aeruginosa are controlled by extracytoplasmic-function (ECF) sigma factors such as PvdS, FpvI, and SigX (45, 46). Given that activation of ECF sigma factors depends on transmembrane signaling, the upregulation of pyoverdine synthesis proteins may result from membrane stress as well. Since the ΔtonB_m strain and the colR mutant possess several common traits indicating their membrane deficiency, the overlapping proteome response can be considered an indicator of a similar type of envelope stress of the two mutants. However, we should emphasize that compared to TonB_m-PocAB-deficient strains, the inactivation of ColRS signaling results in significantly milder phenotypes: colR mutant displays a lower level of glucose-dependent lysis and higher stress tolerance (except for metals like zinc, iron, and cadmium) and has no swimming deficiency (Fig. 9 and data not shown).

In addition to the AlgU regulon, the ColR regulon is activated in the ΔtonB_m strain as well. This response most likely can somewhat alleviate the envelope stress, because the ΔtonB_m colR double mutant displays a stronger growth defect (Fig. 3C and D) and has lower stress tolerance than the strain deficient only in tonB_m (Fig. 9A and B). This suggests that the ColRS and TonB_m-PocAB systems have some overlap in their roles in membrane homeostasis. The activation of the ColR regulon in LB-growing ΔtonB_m cells was somewhat surprising, particularly considering that ColS is a sensor that recognizes the excess of certain metals (25). However, there are two-component systems like PhoP/PhoQ that can sense different stimuli and also detect physical properties of the membrane (47–49). The current study suggests that, besides metals, ColS also senses membrane integrity and ColRS signaling can be triggered by membrane damage.

Both TonB_m-PocAB- and FlhF-deficient mutants have randomly placed flagella (Fig. 1B), but only TonB_m-PocAB deficiency results in membrane stress, indicating phenotypes like glucose-dependent lysis (Fig. 2A), increased membrane permeability (Fig. 2B), and lowered stress tolerance (Fig. 3). This shows that abnormal placement of flagella per se is not causing any membrane defects. Notably, in P. aeruginosa, the positioning of not only FlhF but also CheA, a chemotaxis histidine kinase with unipolar localization, becomes random in TonB3-PocAB-deficient strains (20), indicating that the TonB3-PocAB complex has a more general role in determining polar localization of different protein complexes. Interestingly, a somewhat similar role in the maintenance of membrane integrity as well as in the polar positioning of certain proteins has been reported for the Tol-Pal complex (50, 51). The transenvelope Tol-Pal complex consists of an outer membrane lipoprotein, Pal, a periplasmic protein, TolB, and inner membrane-situated TolA, TolQ, and TolR that are paralogous to TonB_m, PocA, and PocB, respectively. The Tol-Pal complex is part of the cell division machinery. It localizes to the division plane in early predivisional cells, assists proper invagination of the outer membrane, and remains at the new pole until division is completed (51, 52). Besides that, the Tol-Pal complex interacts with chemoreceptors and is required for maintaining the polar positioning of chemoreceptor clusters (50). It is proposed that the Tol-Pal complex physically restricts the departure of the chemoreceptor clusters from the poles after cell division. However, not all polarly localized proteins in E. coli require Tol-Pal for their maintenance in the pole (50). Given that the TonB3-PocAB complex itself is not polarly localized (20), the mechanism of how it determines the polar placement of FlhF and flagella should be indirect and likely does not resemble the mechanism described for the Tol-Pal complex. While the requirements for polar positioning of FlhF are not known, several characteristics of the cell pole can be considered. For example, it has been hypothesized that FlhF detects membrane curvature or recognizes specific proteins or lipids (e.g., cardiolipin and phosphatidylethanolamine) that are enriched at cell poles (53–56). It cannot be ruled out that the TonB-PocAB complex controls the polar placement of FlhF via a specific regulatory mechanism. Still, in light of the current results, we propose that the yet-undetermined polar marker for FlhF is altered due to the membrane defect of the TonB_m-PocAB mutant, which then results in the characteristic mislocalization of flagella.

TonB-like proteins connect the inner and outer membrane, as they are situated in the inner membrane and interact with outer membrane proteins. For instance, TonB of E. coli interacts with TonB-dependent outer membrane transporters to mediate siderophore uptake (12, 57). While it is reasonable to assume that the TonB_m-PocAB complex has a similar role in bridging the two membranes in P. putida, our attempts to detect the potential interaction partners of TonB_m have been unsuccessful so far (data not shown). Hopefully, further studies will reveal the putative interaction partners of TonB_m and disclose the true mechanism of how TonB_m-PocAB maintains membrane integrity and polar positioning of flagella.

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions.

The bacterial strains and plasmids used are listed in Table 1. All strains are derivatives of P. putida PaW85 (58), which is isogenic to the fully sequenced KT2440 (59). Bacteria were grown in lysogeny broth (LB) or on minimal medium (60) containing 0.2% glucose. When selection was necessary, the growth medium was supplemented with benzylpenicillin (800 μg/ml), kanamycin (50 μg/ml), or streptomycin (200 μg/ml) for P. putida and kanamycin (50 μg/ml) or streptomycin (20 μg/ml) for E. coli. E. coli was incubated at 37°C and P. putida at 30°C. Bacteria were electrotransformed according to the protocol of Sharma and Schimke (61).

TABLE 1.

Strains and plasmids

Strain or plasmid	Genotype or characteristics	Source or reference
Strains
Escherichia coli CC118 λpir	Δ(ara-leu) araD ΔlacX74 galE galK phoA20 thi-1 rpsE rpoB argE(Am) recA1 λpir lysogen	73
Pseudomonas putida
PaW85	Wild type, isogenic to KT2440	58
colR strain	PaW85 colR::Km (Kmr)	74
ΔtonBm strain	PaW85 ΔPP_4994	This study
ΔpocA strain	PaW85 ΔPP_1898	This study
ΔpocB strain	PaW85 ΔPP_1899	This study
ΔtonBm ΔpocAB strain	PaW85 ΔPP_4994 ΔPP_1898-PP_1898	This study
ΔflhF strain	PaW85 ΔPP_4343	This study
ΔtonBm ΔflhF strain	ΔtonBm ΔflhF	This study
ΔtonBm tac-tonBm strain	ΔtonBm with genomic lacIq-Ptac-tonBm expression cassette (Smr)	This study
wt/flhF-gfp strain	Wild type with genomic lacIq-Ptac-flhF-gfp expression cassette (Gmr)	This study
ΔtonBm/flhF-gfp strain	ΔtonBm with genomic lacIq-Ptac-flhF-gfp expression cassette (Gmr)	This study
ΔflhF/flhF-gfp strain	ΔflhF with genomic lacIq-Ptac-flhF-gfp expression cassette (Gmr)	This study
ΔtonBm ΔflhF/flhF-gfp strain	ΔtonBm ΔflhF with genomic lacIq-Ptac-flhF-gfp cassette (Gmr)	This study
ΔtonBm oprB1 strain	ΔtonBm and PP_1019::Sm (Smr)	This study
ΔpocA oprB1 strain	ΔpocA and PP_1019::Sm (Smr)	This study
ΔpocB oprB1 strain	ΔpocB and PP_1019::Sm (Smr)	This study
ΔtonBm ΔflhF-oprB1 strain	ΔtonBm ΔflhF PP_1019::Sm (Smr)	This study
ΔmexCD-oprJ strain	PaW85 ΔPP_2817-2819	This study
ΔtonBm ΔmexCD-oprJ strain	ΔtonBm ΔPP_2817-2819	This study
ΔnfxB strain	PaW85 ΔPP_2820	This study
ΔtonBm ΔnfxB strain	ΔtonBm ΔPP_2820	This study
ΔtonBm colR strain	ΔtonBm colR::Km (Kmr)	This study
Plasmids
pUTTn5Sm-lacItac	Delivery plasmid for miniTn5Sm-lacItac (Apr Smr)	75
pRK2013	Helper plasmid for conjugal transfer (Kmr)	64
pKTlacZS/C	Transcriptional fusion of Tn4652 tnpA with lacZ in pKTlacZ	76
pEMG	Plasmid for homologous recombination, lacZa with two flanking I-SceI sites (Kmr)	63
pEMG/ΔtonBm	pEMG containing chimeric DNA fragment for deleting PP_4994 (Kmr)	This study
pEMG/ΔpocA	pEMG containing chimeric DNA fragment for deleting PP_1898 (Kmr)	This study
pEMG/ΔpocB	pEMG containing chimeric DNA fragment for deleting PP_1899 (Kmr)	This study
pEMG/ΔpocAB	pEMG containing chimeric DNA fragment for deleting PP_1898 and PP_1899 (Kmr)	This study
pEMG/ΔflhF	pEMG containing chimeric DNA fragment for deleting PP_4343 (Kmr)	This study
pEMG/ΔmexCD-oprJ	pEMG containing chimeric DNA fragment for deleting ΔPP_2817-2819 operon (Kmr)	This study
pEMG/ΔnfxB	pEMG containing chimeric DNA fragment for deleting PP_2820 (Kmr)	This study
pSW (I-SceI)	Plasmid coding for I-SceI endonuclease for allelic exchange experiments (Bpr)	77
p704L/oprB1::Sm	pGP704L with EcoRI fragment of oprB1::Sm from pKS/oprB1::Sm (Apr Smr)	78
pBRlacItac	Expression vector containing lacIq repressor-controlled Ptac promoter (Ampr)	79
pBRlacItac/tonBm	pBRlacItac containing tonBm in a 1,044-bp HindIII-XbaI fragment under the Ptac promoter (Ampr)	This study
pUCNotKm	Cloning vector (Kmr)	80
pUCNotKm/tactonBm	pUCNot containing lacIq-Ptac-tonBm expression cassette (Ampr)	This study
pBK-miniTn7-ΩSm	pUC19-based delivery plasmid for miniTn7-ΩSm (Ampr Smr)	81
pminiTn7tac-tonBm	pBK-miniTn7-ΩSm containing lacIq-Ptac-tonBm expression cassette (Ampr Smr)	This study
pKTlacItac	Expression vector	82
pKTlacItac/flhF-gfp	Plasmid for expression of FlhF-GFP fusion protein (Ampr)	This study
pGP-mini-Tn7-ΩGm	Delivery plasmid for mini-Tn7-ΩGm (Ampr Gmr)	83
pGPminiTn7Gm/lacItac-flhF-gfp	Delivery plasmid for miniTn7Gm-lacItac-flhF-gfp (Ampr Gmr)	This study
pUXBF13	Plasmid coding for the Tn7 transposition proteins (Ampr, mob+)	84
p9TTBlacZ/algD	PP_1288 promoter fused with lacZ in p9TTBlacZ (Ampr Cmr)	78
p9TTBlacZ/amrZ	PP_4470 promoter fused with lacZ in p9TTBlacZ (Ampr Cmr)	26
p9TTBlacZ/oprQ	PP_0268 promoter fused with lacZ in p9TTBlacZ (Ampr Cmr)	78
p9TTBlacZ/903	PP_0903 promoter fused with lacZ in p9TTBlacZ (Ampr Cmr)	38
p9TTBlacZ/5152	PP_5152 promoter fused with lacZ in p9TTBlacZ (Ampr Cmr)	25
p9TTBlacZ/900	PP_0900 promoter fused with lacZ in p9TTBlacZ (Ampr Cmr)	38

P. putida transposon mutant library screening for identification of Congo red binding mutants.

Wild-type P. putida was subjected to mutagenesis using a Tn5-based minitransposon, miniTn5Sm-lacItac. Plasmid pUTTn5Sm-lacItac was conjugatively transferred from the E. coli CC118 λpir strain into P. putida PaW85 with the aid of the helper plasmid pRK2013. Transconjugants with random chromosomal insertions of the minitransposon were selected on 0.2% glucose minimal plates supplemented with streptomycin, Congo red (0.0005%), and 2 mM phenol. We searched for pink colonies among the white ones. Pink colonies were analyzed by arbitrary PCR and sequencing. PCR products were generated by two rounds of amplification as described elsewhere (62). In the first round, oligonucleotides prtac, specific for the tac promoter, and the arbitrary Arb6 were used as primers. Second-round PCR was performed with the primers OEint and Arb2. Screening of about 35,000 transposon mutants yielded five independent transposon insertions into the tonB_m gene and one into the pocA gene.

Construction of plasmids and strains.

For the generation of deletion strains, the pEMG-based plasmids were constructed according to a protocol described elsewhere (63). The upstream and downstream regions (about 500 bp) of the gene(s) to be deleted were amplified separately and then joined into an approximately 1-kb fragment by overlap extension PCR. Oligonucleotides used in PCR amplifications are listed in Table 1. For construction of the plasmid pEMG/ΔtonB_m, the PCR fragment was cut with BamHI and EcoRI. For construction of the pEMG/ΔpocA, pEMG/ΔpocB, pEMG/ΔpocAB, pEMG/ΔflhF, and pEMG/ΔmexCD-oprJ plasmids, the PCR fragments were cut with XbaI and EcoRI. For construction of pEMG/ΔnfxB, EcoRI and SacI were used. The cut fragments were then ligated into the corresponding sites of the plasmid pEMG. The obtained pEMG plasmids were delivered to P. putida PaW85 or its deletion strains by electroporation, and after 2.5 h of growth in LB medium the bacteria were plated onto LB agar supplemented with kanamycin. Kanamycin-resistant cointegrates were selected and electrotransformed with the I-SceI expression plasmid pSW(I-SceI). To resolve the cointegrate, the plasmid-encoded I-SceI was induced with 1.5 mM 3-methylbenzoate overnight. Kanamycin-sensitive colonies were selected and the deletions were verified by PCR. The plasmid pSW(I-SceI) was eliminated from the deletion strains by growing them overnight in LB medium without antibiotics.

For complementation of the ΔtonB_m strain, the tonB_m gene amplified with the oligonucleotides 4994alg and 4994lopp was first cloned under the control of the tac promoter and lacI^q repressor in pBRlacItac. The lacI^q-P_tac-tonB_m cassette was excised from pBRlacItac/tonB_m with BamHI and subcloned into BamHI-opened pUCNotKm, resulting in pUCNotKm/tactonB_m. Finally, the TonB_m expression cassette was inserted as a NotI fragment into the minitransposon delivery vector pBK-miniTn7-ΩSm. The obtained pminiTn7Sm/tactonB_m was introduced to the E. coli CC118 λpir strain and conjugatively transformed into the P. putida ΔtonB_m strain with the help of pRK2013. The chromosomal presence of the lacI^q-P_tac-tonB_m cassette was verified by PCR.

For construction of C-terminal fluorescent fusion to FlhF, the flhF gene was PCR amplified using primers flhFEco and flhFXho. The EcoRI-XhoI-cleaved flhF fragment was then used to replace the colR gene in the colR-gfp translational fusion in plasmid pKTlacItac/colR-gfp (laboratory collection). The lacI^q-P_tac-flhF-gfp cassette then was inserted into SmaI-KpnI-opened miniTn7 delivery plasmid pGP-miniTn7-ΩGm. The plasmid pGPminiTn7Gm/lacItac-flhF-gfp was introduced into the P. putida wild-type, ΔtonB_m, ΔflhF, and ΔtonB_m ΔflhF strains by coelectroporation together with the helper plasmid pUXBF13. The presence of the lacI^q-P_tac-flhF-gfp cassette in the attTn7 site was verified by PCR.

For construction of oprB1-deficient strains, p704L/oprB1::Sm plasmid was introduced into the E. coli CC118 λpir strain and conjugatively transformed into P. putida ΔtonB_m, ΔpocA, ΔpocB, and ΔtonB_m ΔflhF strains with the help of pRK2013 (64). oprB1 deficiency was verified by PCR.

Motility assay.

The motility assay was conducted on petri dishes containing 20 ml of LB medium supplemented with 0.3% agarose. Motility was assessed by inoculating 1 μl of overnight culture into the medium and measuring the diameter of growth after incubating the petri dishes at 30°C for 18 h.

Microscopy analysis of flagellum localization.

Bacteria were grown in 5 ml LB medium at 30°C until the optical density at 580 nm (OD₅₈₀) was ∼1. The cells were centrifuged at 1,700 × g for 45 s, washed twice with distilled water, and gently resuspended in 1 ml distilled water. Ten-microliter drops of bacterial dilutions (100 cells/μl) were slowly dried on a microscopy slide. The dried drops were covered with stain [0.31% FeCl₃, 6.25% tannin, 92 mM AlK(SO₄)₂] for 4 min, washed with distilled water, and allowed to dry again. The dried drops were then covered with 5% AgNO₃ and 0.0125% NH₄OH solution for 3 min, washed with distilled water, and allowed to dry. Bacteria were observed using oil immersion light microscopy (×1,000 magnification).

Congo red binding assay.

Bacteria were grown overnight in 5 ml LB and diluted 100-fold, and 5-μl drops were inoculated onto glucose minimal medium plate containing 0.0005% Congo red dye. The plate was incubated at 30°C for 72 h.

β-Galactosidase leakage assay.

Strains containing the pKTlacZS/C plasmid were grown on solid glucose minimal or LB medium at 30°C for 20 h. Cells were suspended in 500 μl M9 buffer and centrifuged for 1 min at 12,000 × g, and the supernatant was used to measure β-galactosidase activity according to a previously described protocol (65).

Flow cytometry analysis.

Bacteria were grown overnight in 5 ml LB and diluted 100-fold, and 5-μl drops were inoculated onto agar plates with glucose minimal medium or LB. After 24 h of growth at 30°C, the cells were scraped off from the plates and suspended in 1 ml M9 buffer. The optical density of the cell suspension was diluted to an OD₅₈₀ of 0.015. The two components of the LIVE/DEAD BacLight kit (L7012; Invitrogen), 20 mM red fluorescent dye propidium iodide (PI) and 3.34 mM green fluorescent dye SYTO9, were mixed at a 1:1 volume ratio and then diluted 17.6-fold into filter-sterilized M9 buffer. For staining of bacteria, the diluted cell suspension was mixed with the freshly prepared reagent mixture at a 20:1 ratio. Samples were incubated at 30°C in the dark for 30 min, and approximately 10,000 events from every sample were analyzed with a FACSAria flow cytometer (BD Biosciences). Fluorescent dyes were excited at 488 nm. Forward and side scatter of the light and fluorescence emission at 530 and 616 nm were acquired for every event. Populations of intact, PI-permeable, and dead cells were defined as previously described (28). While PI-permeable cells differ from the intact subpopulation only by their PI-permeable membrane, the dead cells are more compromised, containing less DNA than other cells (28).

Stress tolerance plate assay.

To evaluate the stress tolerance, bacteria were grown overnight (stationary phase) or to an OD₅₈₀ of ∼1 (exponential phase) in 5 ml LB medium. Serial dilutions were spotted as 5-μl drops onto LB agar plates supplemented with different chemicals (specified in Results). Plates were incubated at 30°C for 20 h.

Growth curve and minimal generation time.

To start the assay always with cells in the same physiological state, 50-μl stocks made of overnight-grown bacteria were used. To investigate the growth curve of bacteria starting from stationary phase, cells were inoculated from stock into 5 ml LB medium and grown for 21 h (OD₅₈₀ of ∼4) at 30°C. For exponential-phase cells, 100 μl of the bacteria grown for 18 h were diluted into fresh LB medium and grown for 3 h (OD₅₈₀ of ∼1) at 30°C. The optical densities of microbial cultures at 580 nm were measured and the bacteria were diluted in LB medium for the OD₅₈₀ to be 0.1. Aliquots of 100 μl of the dilutions were transferred into microtiter plate wells, and the cells were grown at 30°C and 400 rpm inside a POLARstar Omega plate reader spectrophotometer. The OD₅₈₀ was measured every 7 min. Data were collected with the Omega data analysis software.

Minimal generation time was calculated from the slope of the most rapid exponential growth according to the formula G = t/3.3log(b/B), where G marks the generation time, t is the time interval in minutes, and B and b are OD₅₈₀ at the beginning and the end of the time interval, respectively.

Isolation of outer membrane proteins.

For the isolation of OMPs of stationary-phase cells, bacteria were grown in 30 ml LB medium at 30°C for 17 h. To obtain the OMPs of exponentially growing cells, bacteria were grown in 200 ml LB for 3 h at 30°C to an OD₅₈₀ of 0.9. Cells were collected by centrifugation at 5,000 × g at 4°C for 10 min, washed with 6 ml of 10 mM HEPES buffer (pH 7.4), and resuspended in 3 ml of the same buffer. Cells were then disrupted by ultrasonication, and cell debris was pelleted by centrifugation at 7,000 × g at 4°C for 15 min. Further OMP isolation was done as described previously (23). One-microgram samples of isolated OMPs were loaded onto 10% SDS-PAGE gel and stained according to a previous protocol (66).

Label-free proteomic analysis.

(i) Growth conditions and nano-LC-MS/MS analysis. Bacteria were pregrown overnight in liquid LB at 30°C, diluted into fresh LB medium (OD₅₈₀ of ∼0.1), and grown either up to an OD₅₈₀ of ∼1 (exponential phase) or for 17 h (stationary phase). Cells were harvested at 5,000 × g at 4°C for 10 min, resuspended in lysis buffer (4% SDS, 100 mM Tris, pH 7.5, 10 mM dithiothreitol), heated at 95°C for 5 min, and sonicated. The protein concentration was measured by tryptophan fluorescence, and 30 μg of cellular protein was loaded onto 30-kDa-cutoff Vivacon 500 ultrafiltration spin columns (Sartorius). Three independent samples of exponential- and stationary-phase P. putida wild-type and ΔtonB_m cells were digested for 4 h on filter with 1:50 Lys-C (Wako) and then overnight with 1:50 proteomics-grade dimethylated trypsin (Sigma-Aldrich) as described for the filter-aided sample preparation protocol (67). Peptides were desalted with C₁₈ StageTips (68), eluted, dried, and reconstituted in 0.5% trifluoroacetic acid. Nano-liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis was performed as described previously (69) using an Ultimate 3000 RSLCnano system (Dionex) and a Q Exactive mass spectrometer (Thermo Fisher Scientific) operating with top-10 data-dependent acquisition.

(ii) MS raw data processing. Raw data were identified and quantified with the MaxQuant 1.4.0.8 software package (70). Label-free quantification with MaxQuant LFQ algorithm (71) was enabled with default settings. Methionine oxidation and protein N-terminal acetylation were set as variable modifications, while cysteine carbamidomethylation was defined as a fixed modification. A search was performed against the UniProt (www.uniprot.org) P. putida KT2440 database (April 2015 version) using the tryptic digestion rule (cleavage after lysine and arginine without proline restriction). Identifications with minimally one peptide of at least seven amino acids were accepted, and transfer of identifications between runs was enabled. Protein quantification criteria were set to two peptides with a minimum of two consecutive MS1 scans per peptide. Peptide-spectrum match and protein false discovery rate were kept below 1% using a target-decoy approach. All other parameters were kept at default.

(iii) Quantitative protein profiling. Data analysis was performed with the Perseus software (72). The whole data set contained 3,359 different proteins. Parallel samples were grouped together, and groups were compared in pairs: (i) P. putida wild type from exponential (wt^E) versus stationary (wt^S) phase (2,652 proteins), (ii) ΔtonB_m strain from exponential (tonB^E) versus stationary (tonB^S) phase (2,673 proteins), (iii) wt^E versus tonB^E (2,406 proteins), and (iv) wt^S versus tonB^S (2,556 proteins). To be included in the analysis, a protein needed to be detected in all three parallels of one group. Thereafter, missing values were imputed using default settings. Mean protein abundances were compared between two groups using the independent-sample Student t test. The Benjamini-Hochberg multiple-testing correction was applied with the false discovery rate set to 0.05.