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. Author manuscript; available in PMC: 2020 Jul 13.
Published in final edited form as: Environ Microbiol. 2019 Mar 12;21(5):1659–1676. doi: 10.1111/1462-2920.14546

Transcriptional regulation of fatty acid cis–trans isomerization in the solvent-tolerant soil bacterium, Pseudomonas putida F1

Tatiana Kondakova 1, John E Cronan 1,2,*
PMCID: PMC7357427  NIHMSID: NIHMS1592192  PMID: 30702193

Summary

One key to the success of Pseudomonas spp. is their ability to reside in hostile environments. Pseudomonas spp. possess a cis–trans isomerase (Cti) an enzyme that converts the cis-unsaturated fatty acids (FAs) of the membrane lipids to their trans-isomers to rigidify the membrane and thereby resist stresses. Whereas the posttranslational Cti regulation has been previously reported, transcriptional cti regulation remains to be studied in more details. Here, we have studied cti transcriptional regulation in the solvent-tolerant strain Pseudomonas putida F1. Two cti transcriptional start sites (cti-279 and cti-77) were identified with cti-279 transcript being dominant. Expression of cti was found to increase with temperature increase, addition of the organic solvent, octanol and in the stationary growth phase. We found that cti expression was repressed by the cyclic-AMP receptor protein (Crp) and repression required the cyclic-AMP ligand of Crp. Production of trans-unsaturated FAs was found to decrease after 24 h of growth. Although this decrease was accompanied by an increase in cyclopropane FA content, this was not at the expense of trans-unsaturated FAs demonstrating the absence of competition between Cti and Cfa in FA modification.

Introduction

Pseudomonas spp. are ubiquitous bacteria that thrive in a wide variety of environments (Mena and Gerba, 2009; De las Heras and De Lorenzo, 2011). The remarkable ecological success of Pseudomonas spp. can be attributed to the metabolic versatility encoded by their large genomes and the presence of sophisticated mechanisms to adjust membrane fluidity. Adjustment of membrane fluidity is accomplished by changes in the fatty acid (FA) composition of membrane lipids. For instance, Pseudomonas spp. are capable to post-synthetically transform cis-unsaturated FA (cis-UFA) into their trans isomers (trans-UFA) without changing the position of the double bond (von Wallbrunn et al., 2003). Based on their stereochemistry trans-UFA mimic, saturated FA (SFA) imparts rigidity and a tight packing of the membrane lipids, whereas cis-UFA gives membranes having relatively high fluidity (Okuyama et al., 1990). Enzymatic isomerization of cis-UFA to trans-UFA is catalysed by Cti and does not depend on the de novo FA and protein synthesis, ATP or any other cofactor (Pedrotta and Witholt, 1999; Heipieper et al., 2003), presenting an efficient mean to rigidify the membrane in the response to changing environments. Cti, which located in the cellular periplasm (Pedrotta and Witholt, 1999), is a cytochrome c-type protein, bearing a heme-binding motif essential for the cis to trans isomerization reaction (Holtwick et al., 1999). Moreover, the highest levels of the trans-UFA were found 30 min after the initiated stress, demonstrating cis–trans isomerization to be a fast adaptive response (Heipieper et al., 1992; Eberlein et al., 2018).

Cyclopropane FA synthase Cfa also uses phospholipid bound cis-UFA as a substrate to replace the double bond with a methylene bridge to give cyclopropane FA (cyclo-FA) (Pohl et al., 1963; Wang et al., 1992). As trans-UFA, cyclo-FA stabilizes membranes against adverse conditions such as acidic conditions (Chang and Cronan, 1999; Poger and Mark, 2015). Cfa is present in most bacteria, whereas Cti is much rarer and to date is found only in a few proteobacterial species (Heipieper et al., 2010), among which Pseudomonas putida seems one of the more interesting to study due to its ability to grow in soils and sediments contaminated with high concentrations of heavy metals and organic contaminants (Ramos et al., 1995; Wu et al., 2011). P. putida is of particular interest in biotechnology because of its high potential in bioremediation and biodegradation processes (Loh and Cao, 2008; Samuel et al., 2014). Thus, several reports report that trans-UFA plays an important role in the adaptation of diverse P. putida strains to the increase in temperature, the presence of organic solvents and heavy metals, as well as osmotic stress and addition of membrane-active antibiotics (Heipieper et al., 1996; Isken et al., 1997; Neumann et al., 2003). It is generally accepted that cti is constitutively expressed in Pseudomonas spp. (Kiran et al., 2004) and proposed to be regulated only post-translationally by the membrane fluidity (Eberlein et al., 2018). However, Bernal and colleagues (2007) reported the upregulation of P. putida cti expression in response to the toluene addition, raising the question about a possible transcriptional cti regulation in this bacterium. Here, we report a detailed assessment of the cti transcriptional regulation in P. putida F1, a soil isolate of known genomic sequence that is resistant to organic solvents (Zylstra et al., 1988). We have characterized the cti promoter region and defined two cti transcriptional start sites (TSS). The effect of stress factors on cti expression was accessed. Hence, we found that cti expression is upregulated by temperature increases, solvent addition, as well as in stationary growth phase. We also found that the cyclic-AMP receptor protein (Crp)–cyclic-AMP complex represses cti expression and that cti is co-transcribed with downstream genes. In addition, this study resolved the question of competition between Cti and Cfa in bacterial membrane modification.

Results

Formation of trans-UFA occurs in P. putida F1 during growth

The production of trans-UFA in P. putida during growth has been documented (Diefenbach et al., 1992; Heipieper et al., 1996). However, several studies reported that some P. putida strains produce trans-UFA only in stress or nongrowing conditions, such as during sample preparation (Härtig et al., 2005; Loffhagen et al., 2007). To ask if P. putida F1 produces trans-UFA in the absence of stress, cultures were grown at the optimal growth temperature of 30°C for 24 h and treated with the powerful chaotrophic agent 10% trichloroacetic acid (TCA) to immediately stop bacterial metabolism (Supporting Information Fig. S1A). FA methyl esters (FAME) were using a CP-Sil 88 column which gave a well-separated peak corresponding to C16:1-trans (Supporting Information Fig. S1B). To test if the formation of C16:1-trans is not the artefact of TCA treatment, FAME of Escherichia coli K-12 MG1655, known to be unable to form trans-UFA, were tested using the same protocol. No trans-UFA was detected in MG1655 samples. Altogether, these data demonstrated that P. putida F1 produces trans-UFA during the growth rather than during the stress of handling.

The pattern of cti transcription in P. putida F1

We have dissected the cti promoter region in P. putida F1 to investigate the response of cti expression to environmental stimuli. Multiple studies in Pseudomonas spp. have sought to define the cti TSS (Junker and Ramos, 1999; Kiran et al., 2005; Bernal et al., 2007). However, these studies targeted Pseudomonas spp. strains of undetermined genome sequence and each identified a different cti promoter. To more systematically characterize cti promoter region in P. putida F1, we performed RLM-RACE analysis. Two cti TSS regions were found located at positions −77 and −279 upstream of the cti translational start site (+1) (Fig. 1A and C). Similar data were reported in P. putida DOT-T1E, where the cti TSS was placed at position −294 (Bernal et al., 2007). Hexamer sequences resembling those proposed to be recognized by the housekeeping σ70 RNA polymerase of Pseudomonas spp. (Ojangu et al., 2000; Domínguez-Cuevas and Marqués, 2004; Vakulskas et al., 2009) were found within the −10 and −35 regions of each of the in P. putida F1 TSS we detected (Fig. 1D).

Fig. 1.

Fig. 1.

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Characterization of the cti promoter region.

A. Sequencing profiles of the amplified fragments obtained by RLM-RACE experiments demonstrating the two cti transcriptional start sites (top: cti-77; bottom: cti-279). The 5′ ends of cti mRNAs are shown by orange lines, and the blue lines show the 5′RACE RNA adaptor sequence. The transcriptional start sites are shown by asterisk.

B. β-Galactosidase activity. The bacteria were grown in PMM for 25 h (OD600 = 2). N = 3.

C. Schematic representation of the cti promoter region. cti-77 and cti-279 transcriptional start sites are shown as black arrows. Their positions are relative to cti translational initiation start (+1). The ORFs are based on P. putida F1-annotated genome sequence. The schema is drawn in scale.

D. DNA sequence of the cti promoter region. Transcriptional start sites are shown as black arrows. The deduced −10 and −35 sequences are underlined.

E. Construction of translational cti′-′lacZ fusions. Black lines show long (CtiPP1) and short (CtiPP2) 5′ cti fragments. Their 5′ end positions shown under the lines are relative to the cti translational initiation start (+1). The fragments were fused in-frame with 3′ lacZ (light grey) to give pFCtiPP1 and pFCtiPP2 plasmids. The pFCtiPP1-based pFCtiPP1KO and pFCtiPP2KO fusions were constructed by 18-bp deletion within putative −10 region of P1-77 and P2-279 respectively. The pFCtiPP0 fusion used as a negative control contains an 88-bp deletion within cti RBS and translational start preventing lacZ expression. For all panels, mean ± SEM is shown. N indicates biological replicates corresponding to independent experiments. Statistical significance was determined by using one-way ANOVA (Turkey multiple comparison). ns, not significant.

To investigate the activity of the two cti promoters, P1 (TSS at −77) and P2 (TSS at −279), we constructed two translational fusions in which one of the promoters drove β-galactosidase expression (cti′–′lacZ fusions). The pFCtiPP1 ‘long’ promoter region fusion contained both P1-77 and P2-279, whereas the pFCtiPP2 fusion corresponded to the ‘short’ promoter region containing only P1-77 (Fig. 1E). The β-galactosidase activity of the long pFCtiPP1 fusion was 26-fold higher than that of the pFCtiPP2 fusion, which showed a very low activity (7.1 Miller units) (Fig. 1B), indicating that in the tested conditions expression from the P2-279 promoter is dominant. To test if the low β-galactosidase activity of pFCtiPP2 fusion was not an experimental artefact, we constructed the pFCtiPP0 plasmid, which had P1-77 and P2-279 promoters but lacked cti ribosome-binding site (RBS) and cti translational start site. This construct showed significantly lower activity (0.5 Miller units) than seen in pFCtiPP2, demonstrating that the P1-77 promoter is able to express the cti gene. We next constructed deletions within P1-77 and P2-279 promoters. Deletion of the putative P1-77 −10 region (pFCtiPP1KO) resulted in an insignificant decrease in β-galactosidase activity. In contrast, deletion of putative −10 region of the P2-279 promoter (pFCtiPP2KO) showed a significant drop in cti expression.

To ask if P1-77 and P2-279 are conserved in P. putida strains, we aligned cti sequences in all P. putida strains having genomes available in NCBI database (last accessed 27 December 2018). All P. putida strains possessed the cti coding sequences. In agreement with previously reported high level of sequence conservation of P. putida the core genome (Udaondo et al., 2016), cti and its 5’UTR were well conserved. For instance, cti of P. putida F1 and DOT-T1E strains (Udaondo et al., 2012), cti of which was previously described (Bernal et al., 2007), has 99% identity, although the lowest cti identity of 77% was found between the F1 and JBC17 strains. Whereas P2-279 promoter was highly conserved in all tested strains, P1-77 was missing in strains E46, E41, PC2, NX-1, IEC33019 and W619 strains (Supporting Information Fig. S2).

Altogether, these data indicate that the predominant cti expression initiates from the well-conserved P2-279 promoter and cti is moderately expressed without stressors addition.

Addition of octanol activates cti expression

The effect of addition of organic solvents to cultures of Pseudomonas spp. on trans-UFA production has been the subject of multiple studies (Junker and Ramos, 1999; Kiran et al., 2005; Heipieper and Fischer, 2010) and showed that addition of various organic solvents activated trans-UFA production. Thus, the ability of organic solvents to activate trans-UFA production was positively correlated with solvent hydrophobicity (Heipieper et al., 1995; Weber and de Bont, 1996; Ramos et al., 2002). Highly hydrophobic solvents partition preferentially into the membranes, where they were supposed to cause an increase of membrane fluidity, which would be offset by increased trans-UFA production. Based on these reports, a model of Cti regulation by membrane fluidity has been proposed (Heipieper et al., 2010). According to this model, an increase in membrane fluidity engendered by addition of organic solvents allows the Cti active centre to reach the double bond to catalyse trans-UFA production (Heipieper et al., 2010). However, studies of the effect of organic solvents on cti expression are few. To test if the addition of organic solvents modulates cti expression in P. putida F1, the pFCtiPP1 and pFCtiPP2 fusions were assayed in octanol-treated cultures (Fig. 2A). In P. putida F1, treatment with different concentrations of this solvent gave a moderate dose-dependent activation of the expression of the pFCtiPP1 and the pFCtiPP2 fusions (Fig. 2B). The cti expression from P2-279 and P1-77 was 1.4- and 1.6-fold higher in 2.5 mM octanol-treated cultures relative to the untreated cultures. These data agree with previously reported results (Bernal et al., 2007) and parallel others (Junker and Ramos, 1999; Kiran et al., 2005). The basal level of trans-UFA in untreated P. putida F1 carrying pFCtiPP1 was already high, due to the tetracycline needed to maintain the ′lacZ fusions (Isken et al., 1997) and did not change significantly with octanol treatment (Fig. 2C). However, when tetracycline was omitted, a dose-dependent increase of trans-UFA production was seen with addition of octanol in agreement with previously obtained results (Heipieper et al., 1995). Importantly, the concentrations of octanol used in this study allowed bacterial survival after 1 h of exposure (Fig. 2D).

Fig. 2.

Fig. 2.

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Addition of octanol activates cti expression and production of trans-UFA in P. putida F1.

A. Workflow. Prior to octanol treatment, bacterial cultures were grown in PMM at 30°C with vigorous shaking (250 r.p.m.) and constant aeration for OD600 of 0.6. After 1 h of treatment with different octanol concentrations, bacterial samples were taken for β-galactosidase activity and FAME assays.

B. β-Galactosidase activity of pFCtiPP1 (left y-axis) and of pFCtiPP2 (right y-axis). N = 3.

C. Percentages of trans-UFA from total bacterial FA. P. putida F1 + pFCtiPP1 cultures were grown in the presence of 20 μg ml−1 Tet. N = 3.

D. Visualization of bacterial susceptibility to octanol by plating serially diluted bacterial cultures on LB plates. N = 3.

E. Time-dependent activation of trans-UFA production in P. putida F1 treated with 2.5 mM octanol. N = 3.

F. Time-dependent β-galactosidase activity of pFCtiPP1 in P. putida F1 treated with 2.5 mM octanol, showing that octanol activates cti expression within 20 min after octanol addition. N = 3. For all panels, mean ± SEM is shown. N indicates biological replicates corresponding to independent experiments. Statistical significance was determined by using one-way ANOVA (Turkey multiple comparison). ns, not significant.

Previously, P. putida P8 cells were found to reach their highest trans to cis ratio 30 min after addition of the membrane-toxic agent 4-chlorophenol (Heipieper et al., 1992). To ask if octanol treatment produced the similar effect, we assayed the time course of formation of trans-UFA in P. putida F1 treated with 2.5 mM octanol. In agreement with (Heipieper et al., 1992), the highest trans-UFA levels were found 30 min after solvent addition (Fig. 2E). The cti expression was activated 20 min after addition of 2.5 mM octanol to bacterial cultures (Fig. 2F), indicating the possible link between octanol-dependent activation of cti expression and increase in trans-UFA production.

Altogether, these results show that the treatment with at least 2.5 mM octanol activates cti expression. Even if the trans-UFA production increased when cultures were treated with lower than 2.5 mM octanol concentrations, the highest amount of trans-UFA was detected in 2.5 mM octanol treated cells. This indicates a possible correlation between solvent-dependent activation of cti expression and trans-UFA production in P. putida F1. The activation of cti expression 20 min after octanol addition followed by the formation of highest trans-UFA levels 30 min after solvent addition supports the hypothesis of correlation between cti expression and trans-UFA production during the solvent addition stress.

Temperature increase and stationary growth phase active the cti expression

Previous studies of the effects of temperature on trans-UFA production in P. putida P8 showed that trans-UFA plays an important role in bacterial adaptation to temperature increase when de novo FA synthesis is blocked. However, when do novo FA synthesis was active, no significant changes in trans-UFA production were detected in response to temperature raise (Diefenbach et al., 1992). We used the P. putida F1 pFCtiPP1 and pFCtiPP2 fusions to assess the effect of growth temperature on cti expression. Given that the production of trans-UFA is considered a fast-adaptive response, we first tested the temperature shift effect. As reported by Diefenbach and colleagues (1992), cultures were grown at optimal growth temperature 30°C until the mid-exponential growth phase and then the growth temperature was shifted to 20 or 3°C for 3 h. The cti-279 and cti-77 expression levels increased with the temperature increase to 37°C and decreased with temperature decrease to 20°C (Fig. 3A). However, the temperature shift produced the opposite effect on the trans-UFA synthesis; the amount of trans-UFA in cultures growing at 20°C was slightly higher that seen in cultures growing at 37°C (Fig. 3B). The same trend was seen in cultures growing with or without tetracycline addition, allowing us to use tetracycline to maintain the reporter plasmids. To ask if this temperature-dependent cti activation is maintained at different growth stages, cultures were grown at 20, 30 and 37°C and the samples were taken at different growth stages for β-galactosidase and FA analyses (Fig. 3C). In agreement with previously reported results, the highest cti-279 and cti-77 expression levels were found in cultures growing at 37°C, whereas at 20°C, cti-279 and cti-77 expression levels were significantly lower than seen in cultures grown at the higher temperatures (30 and 37°C) (Fig. 4D). This temperature-dependent induction was particularly high in stationary growth phase. At all tested temperatures, cti expression was significantly higher in stationary growth phase comparing to exponential growth phase. This agrees with results reported for P. putida DOT-T1E (Bernal et al., 2007), showing an increase in cti expression in stationary growth phase, and parallel others reporting constitutive cti expression in Pseudomonas spp. (Junker and Ramos, 1999; Kiran et al., 2005).

Fig. 3.

Fig. 3.

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Effect of growth temperature and phase on cti expression and production of trans-UFA in P. putida F1.

A. Effect of temperature shift on β-galactosidase activity of pFCtiPP1 and pFCtiPP2 showing the increasing cti expression when the temperature increases. Bacterial cultures grown in PMM at 30°C with vigorous shaking (250 r.p.m.) and constant aeration for OD600 of 0.6 were incubated for 3 h at 20, 30 and 37°C. N = 3.

B. Percentages of trans-UFA from total bacterial FA. P. putida F1 WT + pFCtiPP1 cultures were grown in the presence of 20 μg ml−1 Tet. N = 3.

C. Growth curves of P. putida F1 at 20°C (left), 30°C (centre) and 37°C (right) in PMM. The arrows show the growth stages at which samples of cultures were taken for assays. N = 3.

D. β-Galactosidase activity of pFCtiPP1 (top) and pFCtiPP2 (bottom). N = 4. P-values are calculated by comparing with 30°C.

E. Percentages of total bacterial FA for trans-UFA and cyclo-FA. N = 3. For all panels, mean ± SEM is shown. N indicates biological replicates corresponding to independent experiments. Statistical significance was determined by using one-way ANOVA (Turkey multiple comparison). ns, not significant.

Fig. 4.

Fig. 4.

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Cti does not compete with Cfa.

A. Left panel: growth curves of P. putida F1 strains in PMM at 30°C. The arrows show the growth stages at which samples of cultures were taken for assays. N = 3. Right panel: percentages of trans-UFA and cyclo-FA in total bacterial FA. The numbers on x-axis show the growth stages at which samples of cultures were taken for GC–MS analyses. N = 3. Strain names: P. putida F1 Δcti, PPut 3319; P. putida F1 Δcti compl. cti, PPut 3319C; P. putida F1 ΔcfaB, PPut 5273; P. putida F1 ΔcfaB compl. cfaB, PPut 5273C.

B. Left panel: fractions of cis-UFA and SFA from total bacterial FA in E. coli strains growing with the addition of cis-UFA. Right panel: fractions of cyclo-FA and SFA from total bacterial FA in E. coli strains growing with the addition of cyclo-FA. The numbers on the x-axis show the times after cti induction at which samples of cultures were taken for GC–MS analyses (see Fig. 3C). N = 3. Strain names: ΔfabA Δcfa, TK411; ΔfabA Δcfa cti, TK422; ΔfabA Δcfa ctiΔG23, TK436; ΔfabA Δcfa ctiΔSP, TK455. Sp, signal peptide.

C. Expression of cti leads to E coli growth inhibition. Left panel: growth curves in M9 supplemented with cis-C18:1 or cyclo-C19:0. The black arrows show the time (h) after induction of cti expression at which samples of cultures were taken for GC–MS analyses (N = 3). Right panel: visualization of bacterial susceptibility to Cti production by plating serially diluted bacterial cultures on LB + cis-C18:1 plates. N = 3. For all panels, mean ± SEM is shown. N indicates biological replicates corresponding to independent experiments. Statistical significance was determined by using one-way ANOVA (Turkey multiple comparison).

In agreement with temperature shift results (which corresponded to point 2 in Fig. 3C–E), the trans-UFA content was slightly lower in cultures growing at 37°C relative to cultures growing at 20°C in all growth phase, except exponentially growth phase. Interestingly, at all tested temperatures, the trans-UFA content of the cells significantly decreased after 24 h of bacterial growth (Fig. 3E). This decrease was followed by a significant increase of cyclo-FA. It is, consistent with previous reports, found the cyclo-FA to be predominantly produced in stationary phase of growth (Wang and Cronan, 1994; Grogan and Cronan, 1997). Taken together these data indicated that in P. putida F1, cti expression is activated by temperature rise and, in stationary phase of growth, conditions in adaptation to which trans-UFA were reported to play an important role. The mismatch between the fraction of trans-UFA and cti expressions suggested the possible post-transcriptional control of trans-UFA production in P. putida F1. In addition, the decrease in trans-UFA production after 24 h of bacterial growth in favour of cyclo-FA overproduction suggested the possibility of competition between Cti and Cfa.

The absence of competition between Cti and Cfa

Previous studies tested the competition between Cti and Cfa in early stationary growth phase (corresponding to point 3 in Fig. 3C–E) and suggested the absence of competition between those two enzymes in P. putida KT2440 (Pini et al., 2011, 20,100). We observed decreased trans-UFA and increased cyclo-FA production after 24 h of growth (starting at point 4 in Fig. 3C–E), which raised a question about the possibility of competition between Cti and Cfa after 24 h of growth. To test this possibility, we first made the knockout mutants of the corresponding genes in P. putida F1 and compared the FA composition of the mutant strains with that of the WT strain at different stages of growth. The P. putida F1 Δcti (PPut3319) strain produced a level of cyclo-FA indistinguishable from that of the WT strain (Fig. 4A). However, after 24 h of growth the P. putida F1, ΔcfaB (PPut5273) strain produced twice the level of trans-UFA seen in the WT strain. Complementation with cfaB gene in trans (PPut5273C) was effective and the complemented strain produced less trans-UFA than the mutant. These results supplement those obtained in a previous report (Pini et al., 2011) and have multiple interpretations. The competition between Cti and Cfa aside, the inability of the ΔcfaB strain to synthetize cyclo-FA can alter membrane fluidity. Given that both cyclo-FA and trans-UFA are able to order the membrane and decrease membrane fluidity (Loffhagen et al., 2007), P. putida F1 ΔcfaB may overproduce trans-UFA to compensate for the absence of cyclo-FA.

To dissect this possibility, we made an E. coli ΔfabA Δcfa mutant strain (TK411) that is unable to synthetize cis-UFA (and hence cyclo-FA) and expressed P. putida F1 cti in this strain (ΔfabA Δcfa cti, TK422). As a control, we used the ΔfabA Δcfa ctiΔG23 strain (TK436), which was unable to form trans-UFA due to a frame-shift (deletion of base G23) in the cti sequence. The culture medium was supplemented with either a cis-UFA or a cyclo-FA to allow growth of the strain. When fed with cis-UFA, ΔfabA Δcfa cti strain formed trans-UFA. However, the fraction of trans-UFA did not exceed 9.6% of the total FA and remained quite stable for 13 h of cti induction (Fig. 4B). Contrary to our expectations, the fraction of cis-UFA increased with cti induction, suggesting the absence of competition between Cti and Cfa, as well as an important role of cis-UFA in trans-UFA-forming bacteria. At 6 h of cti induction, strain ΔfabA Δcfa cti contained a significantly lower SFA content compared to the cti frameshift strain ΔfabA Δcfa ctiΔG23. This difference was most striking at 13 h of cti induction, when the strain ΔfabA Δcfa cti contained 2.5-fold more cis-UFA (thus 2.5-fold less SFA) than ΔfabA Δcfa ctiΔG23 strain. When fed with the cyclo-FA, no trans-UFA was formed and no large change in membrane FA composition was found. However, at 13 h of cti induction ΔfabA Δcfa cti had 10% more cyclo-FA and 10% less SFA than ΔfabA Δcfa ctiΔG23, whereas the FA compositions of strains ΔfabA Δcfa ctiΔG23 and ΔfabA Δcfa were similar. These data indicated the absence of competition between Cti and Cfa within the bacterial membrane and suggested that E. coli can adapt the membrane FA composition to trans-UFA production by increasing the fraction of cis-UFA into the membrane.

Cti toxicity

Interestingly, the growth of cti expressing strain ΔfabA Δcfa cti was completely inhibited after induction of the arabinose pBAD promoter driving its expression whereas ΔfabA Δcfa ctiΔG23 grew as well as ΔfabA Δcfa (Fig. 4C). This growth defect was independent of trans-UFA production, as shown when ΔfabA Δcfa cti was fed with cyclo-FA. Hence, toxicity seemed due to the Cti protein per se rather than its action. Given that Cti is a periplasmic protein (Pedrotta and Witholt, 1999), its overexpression could result in a lethal jamming of the E. coli secretion machinery (Shuman and Silhavy, 2003). To test this possibility, we deleted the cti signal peptide sequence (ctiΔSP) and assayed the growth of ΔfabA Δcfa ctiΔSP strain (TK455). This strain failed to produce any detectable amount of trans-UFA and had a membrane FA composition identical to that of strains ΔfabA Δcfa and ΔfabA Δcfa ctiΔG23 (Fig. 4B), indicating that the periplasmic location of Cti is necessary to produce trans-UFA. Moreover, this bacterium grew as well as the ΔfabA Δcfa and ΔfabA Δcfa ctiΔG23 strains, arguing that Cti toxicity in E. coli is very likely due to the lethal jamming of secretion system.

Crp represses cti transcription in P. putida F1

Upon binding cAMP, Crp is one of the most important transcription factors in E. coli (Deutscher, 2008; Fic et al., 2009). When the cAMP–CRP complex acts on a binding site for gene regulation, it has one of the two opposing regulatory roles: activation or repression of gene transcription (Mori and Aiba, 1985; Yang et al., 2014). We analysed the P. putida F1 cti promoter region and identified two putative Crp-binding sequences (Crp1 and Crp2, Fig. 5A). The Crp1-binding region was located downstream of the cti-77 TSS, close to the RBS sequence. The Crp2-binding region was located just downstream of the RNA polymerase −35 binding site of cti-279 promoter. Similar Crp-binding sites were predicted by PredCRP-model, which assumes that when Crp binds to these regions, it blocks the transcription process (Tsai et al., 2018).

Fig. 5.

Fig. 5.

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Crp binds a region downstream of the P. putida F1 cti P1 promoter and represses cti expression.

A. DNA sequence of cti promoter region. Transcriptional start sites cti-77 and cti-279 are shown as black arrows. The deduced −10 and −35 sequences are underlined. Two putative Crp-binding sequences identified using the PRODORIC Database (Münch et al., 2003) are in grey boxes.

B. EMSA showing cAMP–Crp binding to 5′ cti fragment. Left top: cAMP–Crp binding to 5′ cti fragment (Pcti). Note the shifts seen for cAMP–Crp complex versus the absence of shift seen for the Crp along or cAMP along samples. Right top: The absence of cAMP–Crp binding to 5′ cfaB fragment (Pcfa). Left bottom: cAMP–Crp binding to Pcti containing the deletion of Crp 2 binding sequence (Pcti Δcrp2). Right bottom: The absence of cAMP–Crp binding to Pcti containing the deletion of Crp 1 binding sequence (Pcti Δcrp1).

C. β-Galactosidase activity of pFCtiPP1 showing than Crp represses cti expression in P. putida F1. Bacteria were grown in PMM at 30°C. The numbers on x-axis show the growth stages at which samples of cultures were taken for β-galactosidase analyses (see Fig. 2B). N = 3. Strain names: Δcrp, PPut0457; Δcrp compl. crp, PPut0457C.

D. β-Galactosidase activity showing the expression from Pcti Δcrp1, Pcti Δcrp2 and Pcti. N = 3.

E. Proposed mechanism of Crp–cAMP-dependent repression of cti transcription in P. putida F1. Crp is shown in grey as a dimer and cAMP is shown as a rhomb. Crp–cAMP complex binds Crp1 site (black rectangle) located near ribosome-binding site (RBS) and represses cti transcription. The schema is drawn in scale. For all panels, mean ± SEM is shown. N indicates biological replicates corresponding to independent experiments. Statistical significance was determined by using one-way ANOVA (Turkey multiple comparison). ns, not significant.

Previous studies showed that the properties of E. coli Crp regarding its interaction with target DNA sequences have been preserved in the P. putida counterpart. Moreover, threading of the P. putida Crp sequence on the crystal structure of E. coli Crp strengthens the notion that they are not just two related proteins but are essentially the same protein with minor changes, which do not affect its key functions (Milanesio et al., 2011). Given the difficulties reported in the production of pure and high-activity P. putida Crp protein (Arce-Rodríguez et al., 2012), we purified E. coli Crp and performed electrophoretic mobility shift assays (EMSA) to test if Crp binds the cti promoter sequence in vitro (Fig. 5B). The cAMP-CRP complex bound the P. putida F1 cti promoter region (Pcti), whereas binding was not observed in the absence of cAMP or to the P. putida F1 cfa promoter region (Pcfa). We next deleted the Crp-binding sites from Pcti (giving Pcti Δcrp1 and Pcti Δcrp2) and tested Crp binding to obtained DNA fragments. Crp bound Pcti Δcrp2 as well as Pcti, whereas no binding was observed to Pcti Δcrp1, indicating that the Crp1 site is necessary for Crp binding. In addition, both Crp1- and Crp2-binding sites were conserved in P. putida strains (Supporting Information Fig. S2), indicating the possible Crp-dependent cti regulation in several P. putida strains.

To test in vivo if Crp regulates P. putida F1 cti transcription, we made a P. putida F1 Δcrp mutant strain (PPut0457) and assayed the activity of the pFCtiPP1 fusion in this strain. This mutant showed a significant increase in β-galactosidase activity comparing to WT (Fig. 5C) indicating that Crp represses cti expression in P. putida F1. Complementation with crp gene was effective and the complemented strain (PPut0457C) showed lower expression of the pFCtiPP1 fusion than did the mutant strain. We also assayed the activity of the pFCtiPP1crp1KO strain and pFCtiPP1crp2KO fusion strains containing Pcti Δcrp1 and Pcti Δcrp2 DNA fragments, respectively, in P. putida F1 WT (Fig. 5D). The activity of pFCtiPP1crp1 KO was modest and growth stage independent, suggesting that Crp1-binding site may play a dual and important role in cti expression. The activity of pFCtiPP1crp2KO was also growth stage independent but significantly higher than that of pFCtiPP1 in exponential growth phase and similar to it in stationary growth phase. This indicated that Crp2-binding site may play a role in cti regulation in exponential growth phase by a Crp independent mechanism. In agreement with PredCRP-model (Tsai et al., 2018), our data showed that Crp represses cti expression. However, our EMSA results suggested that only the Crp1 site located distal downstream of cti promoter sequences is necessary for Crp binding, highlighting the importance of validation of the in-silico studies by experimental assays.

cti is co-transcribed with downstream genes in Pseudomonas spp.

Previous studies assumed that cti is transcribed as a monocistronc unit (Junker and Ramos, 1999; Heipieper et al., 2003). We analysed the cti genomic environment in Pseudomonas spp. using the MaGe platform. In all available Pseudomonas spp., cti was the first gene downstream of which one or several genes transcribed at the same direction were located (Fig. 6A). To ask if these genes are transcribed in operon with cti, we used RT-PCR in octanol-treated P. putida F1 and P. aeruginosa PAO1 cells. In both strains, cDNA amplifications using primers specific for cti and downstream genes were obtained (Fig. 6B and C), indicating that in tested experimental conditions, cti is co-transcribed with the downstream genes. Gene PA1847 in P. aeruginosa PAO1, as well as Pput_3317 in P. putida F1, is annotated as coding for NfuA, a protein involved in Fe/S biogenesis in E. coli (Angelini et al., 2008) and Acinetobacter baumannii (Zimbler et al., 2012). The gene Pput_3318 in P. putida F1 is annotated as acyltransferase 3 with unknown function. To ask if these genes are involved in trans-UFA synthesis in Pseudomonas spp., we made nonpolar knockout mutants of both genes and assayed FA composition of obtained mutant strains. Neither mutant strain showed a significant difference in trans-UFA content compared to the WT strain (Fig. 6D), indicating that these genes are not involved in trans-UFA synthesis.

Fig. 6.

Fig. 6.

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cti is co-transcribed with downstream genes in P. putida F1 and P. aeruginosa PAO1.

A. Physical map of cti and downstream genes in eight Pseudomonas spp. type strains. The ORFs are based on Pseudomonas spp. genome sequences and annotation available from the MaGe Platform. The Pput_3319 (cti) orthologs are shown in green; Pput_3318 orthologs are shown in orange; Pput_3317 orthologs are shown in blue; Pput_3316 orthologs are shown in purple; and Pput_3315 orthologs are shown in yellow. Black lines show the chromosomal DNA. Double lines show the different chromosome location.

B. Top: physical map of cti and genes located downstream in P. putida F1. Predicted Rho-independent transcriptional terminator is indicated with lollipop shapes. The location of primers along the DNA sequence is indicated by light grey arrows with their names given. Bottom: genomic DNA and cDNA amplicons separated in 1% agarose gel electrophoresis (N = 3). RT– lines show the no-reverse-transcriptase-negative controls. (−) negative PCR control with H2O. (+) Genomic DNA amplicons.

C. Left: physical map of cti and genes located downstream in P. aeruginosa PAO1. Right: genomic DNA and cDNA amplicons separated in 1% agarose gel electrophoresis (N = 3).

D. Deletion of genes located downstream from cti does not affect the production of trans-UFA in P. putida F1 (left) and P. aeruginosa PAO1 (right). Bacteria were grown in PMM until OD600 0.6 for P. putida F1 (N = 6) and OD600 0.4 for P. aeruginosa PAO1 (N = 3). P-values are calculated by comparing with WT. Mean ± SEM is shown. For all panels, N indicates biological replicates corresponding to independent experiments. Statistical significance was determined by using one-way ANOVA (Turkey multiple comparison). ns, not significant. Strain names: Δ3317, PPut3317; Δ3318, PPut3317; ΔPA1847, PAO1847.

Discussion

The Pseudomonas spp. comprise one of the most widely distributed groups of bacteria. Consistent with this wide distribution, these bacteria developed several resistance mechanisms to respond to diverse environmental stresses (Rainey, 1999; Monteil et al., 2016; Moradali et al., 2017). One mechanism is membrane FA modification by the conversion of cis-UFA to their trans isomers (Heipieper and Fischer, 2010). Although the first report of trans-UFA in Pseudomonas was 31 years ago (Guckert et al., 1987), the regulation and mechanism of their formation remain a puzzle. We have examined cti transcriptional regulation in the soil-borne solvent degrader P. putida F1, competition between its Cti and Cfa enzymes, as well as the toxicity of expression of P. putida F1 Cti in E. coli.

First, our data demonstrate that P. putida F1 produces trans-UFA during the growth. This agrees with several previous reports (Diefenbach and Keweloh, 1994; Heipieper et al., 1996) and disagrees with other reports (Härtig et al., 2005; Loffhagen et al., 2007). This could be due to technical issues or possibly differences in the strains studied. This is the first report of cti action and regulation in the otherwise heavily studied P. putida F1 strain.

We identified two P. putida F1 cti TSS. Our data showed that in P. putida F1, cti is mainly expressed from the P2-279 promoter, which is highly conserved in all available P. putida strains. Whereas cti was expressed at basal level without stressors addition, promoter P2 is moderately activated upon addition of organic solvent octanol, as well as in stationary growth phase. These results are similar to those obtained in a previous report (Bernal et al., 2007) and are in contrast to two other studies reporting constitutive cti expression (Junker and Ramos, 1999; Kiran et al., 2005). In the first report, the Ramos research group used a Cti::PhoA fusion. PhoA becomes active only in the periplasmic space (Junker and Ramos, 1999), which raises the possibility that effects on cti expression could have been masked by effects on secretion. In our work as well as in that reported by Bernal and colleagues (2007) expression of a Cti′::′LacZ fusion was monitored. The second report was focused on cti expression in P. syringae Lz4W strain. Given the genetic heterogeneity of the genus Pseudomonas (Bodilis et al., 2012), cti expression in P. putida and in P. syringae species could well differ markedly.

The previously reported constitutive cti expression in solvent-treated bacteria led to the establishment of a Cti regulation model, according to which Cti activity is not controlled at the transcriptional level, but rather by simply allowing or prohibiting the active center of the enzyme to reach the double bond, which would depend only on membrane fluidity (Heipieper et al., 2010; Eberlein et al., 2018). The activation of cti expression in octanol-treated P. putida F1 cultures observed in our studies demonstrates that in response to solvent addition cti is regulated at the transcriptional level. The quick activation of cti expression after solvent addition followed by overproduction of trans-UFA allows us to propose that cti overexpression is activated to respond to high solvent concentrations and leads to the production of highest trans-UFA levels necessary to maintain the optimal membrane fluidity. Thus, our study adds an additional level of Cti regulation to the previous model, showing the complexity of the mechanisms that regulate Cti action. However, Cti overexpression in E coli led to stable but modest production of trans-UFA. Given that P. putida S12 was found to have only 2–3 copies of Cti per cell (Neumann et al., 2003) and Cti expression in E. coli would be expected to result in greater trans-UFA production than we observed. Thus, our data agree with Heipieper et al. model, indicating possible postsynthetic Cti regulation. However, future studies are necessary to test this possibility, as well as the role of membrane fluidity in Cti regulation.

We found that cti expression is induced by increased growth temperature. The effect of growth temperature on trans-UFA production is well documented in P. putida (Diefenbach et al., 1992) and in Vibrio spp. (Okuyama et al., 1990, 1991). However, to the best of our knowledge, this is the first report of modulation of cti expression by growth temperature. However, production of trans-UFA did not increased with increasing temperature in agreement with a previous report (Diefenbach et al., 1992). This mismatch between cti expression and production of trans-UFA suggests posttranscriptional Cti regulation in Pseudomonas spp. and requires future study. Posttranscriptional Cti regulation could also explain the decreased trans-UFA production in stationary growth phase, the stage of bacterial growth at which we found the highest cti expression. Previously, the effect of growth phase on the production of trans-UFA in P. putida was only tested at the transition from exponential to stationary growth phase (Diefenbach et al., 1992).

The decrease in trans-UFA production followed by the increase in cyclo-FA raised the question of possible competition between Cti and Cfa since they modify the same substrate. Previous studies suggested the absence of competition between those two enzymes in early stationary growth phase (Pini et al., 2011, 20,100), whereas we assayed the competition at different stages of the growth of P. putida F1 cultures and also in an E. coli model we constructed. First, the P. putida F1 ΔcfaB strain produced more trans-UFA than the WT strain after 24 h of growth. However, the E. coli model indicated the absence of competition between Cti and Cfa. In contrast to our expectations, the fraction of cis-UFA in E. coli membrane increased with the induction of cti, indicating that cis-UFA are incorporated into the bacterial membrane in higher quantities to offset the trans-UFA production and optimize membrane fluidity. Hence, we propose that the increase in trans-UFA formation in P. putida F1 ΔcfaB is not due to the competition between Cti and Cfa, but to the increased production of this FA perhaps to optimize membrane fluidity in the absence of cyclo-FA. These data complement previous report (Pini et al., 2011) and add additional information about Cti and Cfa competition. In addition, these assays led us to discover a rationale for Cti toxicity in E. coli cells, which is probably caused by lethal jamming of E. coli secretion machinery. However, the observed decrease in SFA content in E. coli expressing Cti could also explain Cti toxicity. Given that E. coli does not encode a Cti enzyme, FabR-dependent membrane regulation (Zhu et al., 2009), as well as the functioning of several membrane proteins (Bogdanov et al., 2008; Dowhan and Bogdanov, 2011), could be impaired when trans-UFA are produced leading to the observed toxic effect.

In E. coli and many other bacteria, Crp plays a role of catabolite repressor (Brückner and Titgemeyer, 2002; Deutscher, 2008), whereas Crp ortholog in P. aeruginosa (named Vfr) is mostly associated with virulence factors’ expression (Fuchs et al., 2010). In P. putida, Crp is not involved in metabolic control (Milanesio et al., 2011) and to date its role in this bacterium remains unelucidated. Here, for the first time, we showed that Crp represses cti expression in P. putida F1, thereby demonstrating one of the first Crp targets in P. putida. Interestingly, most phenotypes of P. putida Δcrp mutant were previously shown to be traced to envelope-associated functions (peptide permeases, membrane-disrupting agents) (Milanesio et al., 2011). Our study adds an additional envelope-associated Crp target in P. putida and indicates that P. putida Crp could be associated with the regulation of cell envelope composition. In addition, our EMSA studies complemented previously reported PredCRP model (Tsai et al., 2018), by showing that even in the presence of two putative Crp-binding sites, only the downstream site is necessary for Crp binding and repression of cti expression (Fig. 5E). Despite the general assumption that repressors of prokaryotic operons act exclusively by preventing the onset of transcription, Crp seems to bind only Crp1 site located distal downstream of cti promoters and not to the Crp2 site located within the DNA sequence covered by promoter-bound RNA polymerase. Similar results were shown for lac operon regulation, where a lac repressor-operator complex located downstream of the promoter sequence can directly interfere with gene expression by efficiently terminating transcription (Deuschle et al., 1986).

The results presented here provide the most detailed assessment of cti transcriptional regulation in any bacteria to date. This study brings a stone in understanding of Pseudomonas spp. adaptation to environmental stimuli and can be useful to develop the strategies for biodegradation of toxic compounds in natural environments.

Experimental procedures

Strains and growth conditions

Strains and bacterial plasmids used in this study are listed in Supporting Information Table S1, and details of their construction are in the Supporting Information. Escherichia coli K-12 derivatives were grown at 37°C in LB, SOC or M9 medium. Diaminopimelic acid (1 mM) was added to the media for the growth of E. coli WM6029. P. aeruginosa PAO1 and P. putida F1 were grown at 37 and 30°C, respectively, in LB or Pseudomonas minimal medium (PMM) with sodium succinate as carbon source (Kirner et al., 1996). Given that the growth conditions are relevant for the cis–trans isomerization, all assays were performed in PMM with vigorous shaking (250 r.p.m.) and the constant aeration (the volume of cultures represented 10% of the total flask volume). For plasmid maintenance, PMM was supplemented with 20 μg ml−1 tetracycline hydrochloride (Tet). Oligonucleotides used in this study are listed in Supporting Information Table S2. For plasmid maintenance in E. coli, the medium was supplemented with (in μg ml−1) kanamycin sulfate (Km) and gentamicin sulfate (Gm), 50 or Tet, 10. The antibiotic concentration (in μg ml−1) used in LB medium for P. putida F1 was Km, Tet and Gm, 50 whereas the concentrations for P. aeruginosa PAO1 were Km, 1000 and Gm, 50. The detailed media composition is in the Supporting Information.

Extraction of total RNA, cDNA synthesis and amplification

Total RNA was purified using RNease Mini Kit (Qiagen). Details of this procedure are provided in SI Appendix, Supplemental Materials and Methods. RNAs were non-specifically converted to single stranded cDNAs using the ProroScript First Stand cDNA Synthesis Kit (NEB Biolabs). The control samples, RT− (lacking reverse transcriptase) were made during cDNA synthesis from the total RNA using the ProroScript First Stand cDNA Synthesis Kit without the addition of reverse transcriptase. All experiments were performed in three biological replicates.

RLM-race

The 5′ ends of cti mRNA in P. putida F1 were mapped using RLM-RACE according to the manufacturer’s instructions. Detailed procedure is reported in the Supporting Information. To identify the 5′ ends of the cti mRNA, the PCR products were cloned into vector pMTL23P (Chambers et al., 1988) and sequenced. All experiments were performed in three biological replicates.

Monitoring of cti transcription

To construct the cti′–′lacZ translational fusions in P. putida F1, 5′ fragments of cti (Pcti′) were fused in the correct translational reading frame to a large 3′ fragment of lacZ (′lacZ) gene in vector pSRKTc (Shuman and Silhavy, 2003; Khan et al., 2008). Detailed procedure is reported in the Supporting Information. Then, the deletions into cti promoter regions were made using Site Directed Mutagenesis Kit (NEB Biolabs).

β-Galactosidase assays

Bacterial cultures (2 ml) were centrifuged, and the pelleted cells were assayed for β-galactosidase activity after chloroform/sodium dodecyl sulfate lysis as described by (Miller, 1972). The cell debris in the assay mix was removed by centrifugation before absorbance was read at 420 and 550 nm. All experiments were performed at least in three biological replicates.

Construction of Pseudomonas spp. knockout mutants

To make P. putida F1 knockout mutants, the suicide plasmids were constructed as described previously (Choi and Schweizer, 2005). Briefly, the sequences’ upstream and downstream of target gene, as well as the Gm cassette flanked by FRT sites, were amplified from pFGm1 vector (Choi et al., 2005) and assembled into the EcoRI BamHI digested pMTL23P vector using the Gibson Assembly Kit (NEB). The obtained insert was ligated into BamHI EcoRI digested pK19mobsacB suicide vector (Schäfer et al., 1994). A rapid electroporation method described elsewhere (Choi et al., 2006) was used to transfer the pK19mobsacB-borne deletion mutations to Pseudomonas spp. strains. When indicated the Gm cassette was removed from the chromosomal DNA with help of flippase encoded by plasmid pFLP3 (Choi et al., 2005) according (Choi and Schweizer, 2005). The presence of the correct mutations was verified by colony PCR and sequencing. To complement the mutations, the corresponding genes were PCR amplified with their native promoters from genomic DNA. The obtained fragments were ligated into vector pUC18R6KT-mini-Tn7T-Km and inserted into P. putida F1 chromosome using pTNS2 transposase expression vector (Choi et al., 2005). The detailed protocol can be found in the Supporting Information.

FA analysis

Phospholipids were extracted as described by Bligh and Dyer (1959), and FA methyl esters were prepared according the lab protocol (Zhu and colleagues (2010)). FA methyl esters and their standards were measured by GC–MS on highly polar chiral (CP-Sil88) column. When indicated, FA methyl esters were prepared according to a prior report (Härtig et al., 2005) and analysed using a standard non-polar (ZB-5MS) column. All experiments were performed in three biological replicates (for more details see Supporting Information).

Crp expression and purification

Hexahistidine-tagged E. coli Crp protein was produced in E. coli BL21 (DE3) carrying the expression plasmid pET28-crp (Supporting Information Table S1) as previously reported by Feng and Cronan (2012). The detailed protocol can be found in the Supporting Information. P. putida F1 Crp (Pput_0457) is not only 63% identical and 80% similar to the E. coli counterpart but also keeps the known motifs for the activation of RNA polymerase through direct protein-protein interactions to maintain the same helix-turn-helix (HTH) motif for DNA binding (identity HTH domains: 72%) and to preserve the basic architecture of its cyclic nucleotide (cNMP)-binding site (identity cNMP domains 55%) (Milanesio et al., 2011; Arce-Rodríguez et al., 2012).

Electrophoretic mobility shift assays

These assays of the interaction between the cAMP-CRP complex and P. putida F1 cti promoter region were done essentially as previously reported (Herrera et al., 2012; Henke and Cronan, 2016). The DNA-binding reaction contained 200 μg ml−1 of bovine serum albumin, 50 mM Tris-HCl (pH 7.5), 100 mM NaCl, 0.1 mM dithiothreitol (DTT), 100 μM cAMP and 1 mM EDTA, 40 nM DNA and the indicated concentrations of Crp. Reactions were incubated at room temperature for 30 min and run in a 6% DNA retardation gel. The gel was stained with SYBR Green I nucleic acid gel stain and visualized using BioRad Chemidoc XRS and Quantity One software.

Heterologous cti expression in E. coli

The essays were carried out in M9 medium supplemented with 0.01% oleic acid (cis-C18:1) or 0.01% cis-9,10-methyleneoctadecanoic acid (cyclo-C19:0). After 11 h of growth (OD600 ≈ 0.5) in M9 supplemented with 0.4% glucose and 0.8 mM d-fucose, the cells were washed and resuspended in M9 medium supplemented with 0.8% glycerol and 0.1% arabinose to induce the PBAD promoter responsible for cti expression. FA compositions were assayed at 0, 3, 6 and 13 h after induction. After 13 h of induction, 10-fold serial dilutions of the bacterial cultures were plated and incubated at 37°C for 16 h. All experiments were performed in three biological replicates.

Bioinformatic analyses

Analysis of the cti operons of Pseudomonas spp. was performed with the ‘Synteny Line Plot’ tool available from the MaGe Platform (http://www.genoscope.cns.fr/agc/mage, last accessed 10 June 2018), which carried out a global comparison of 63 Pseudomonas spp. genomes on the basis of synteny results. Analysis of cti promoter regions was carried out by PRODORIC® Database (Münch et al., 2003) with standard settings and P. aeruginosa PAO1 as a query. To align cti and its 5′UTR sequences of P. putida strains, all available P. putida cti orthologs were extracted from the NCBI Platform (last accessed 27 December 2018), which carries out the Blast analysis using P. putida F1 cti nucleic acid sequence (Pput_3319) and 507 bp of cti 5′UTR as a query in BLAST-N searches. The resulting sequences were aligned using CLC Sequence Viewer 7 (CLC bio).

Supplementary Material

Supplement Complete

Supporting Information

Additional Supporting Information may be found in the online version of this article at the publisher’s web-site:

Fig. S1. Representative GC-MS total ion chromatogram of fatty acid methyl esters showing that P. putida F1 forms trans-UFA during the growth. (A) Workflow. (C) TIC of methyl esters separated by CP-Sill 88 column. P. putida F1 samples are shown in black, E. coli K-12 MG1655 samples are shown in green and standards are shown in red. Experiments were made in three biological replicates corresponding to three independent experiments. RT, room temperature; TCA, trichloroacetic acid; PMM, Pseudomonas minimal medium.

Fig. S2. Alignment of cti promoter region sequences in P. putida strains. Crp 1 and Crp 2 binding sequences are shown as blue squares. Ribosome-binding site (RBS) is shown as a green square, and translational start is shown as a green star. Transcriptional start sites are shown as black arrows. The deduced −10 and −35 sequences are underlined.

Appendix S1: Supporting Information 1.

Appendix S1: Supporting Information 2.

Acknowledgements

We thank Dr. Alex Ulanov of the Metabolomics Centre, University of Illinois for help with GC–MS analyses. This work was supported by National Institutes of Health Grant AI15650 from the National Institute of Allergy and Infectious Diseases.

Footnotes

Conflict of interest

We have no conflicts of interest to declare.

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Fig. S1. Representative GC-MS total ion chromatogram of fatty acid methyl esters showing that P. putida F1 forms trans-UFA during the growth. (A) Workflow. (C) TIC of methyl esters separated by CP-Sill 88 column. P. putida F1 samples are shown in black, E. coli K-12 MG1655 samples are shown in green and standards are shown in red. Experiments were made in three biological replicates corresponding to three independent experiments. RT, room temperature; TCA, trichloroacetic acid; PMM, Pseudomonas minimal medium.

Fig. S2. Alignment of cti promoter region sequences in P. putida strains. Crp 1 and Crp 2 binding sequences are shown as blue squares. Ribosome-binding site (RBS) is shown as a green square, and translational start is shown as a green star. Transcriptional start sites are shown as black arrows. The deduced −10 and −35 sequences are underlined.

Appendix S1: Supporting Information 1.

Appendix S1: Supporting Information 2.

NIHMS1592192-supplement-Supplement_Complete.pdf (8.4MB, pdf)

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