INTRODUCTION
Quorum sensing (QS) is a bacterial cell-cell communication process that allows bacteria to synchronize particular behaviors on a population-wide scale. Within current knowledge, QS in
Stenotrophomonas maltophilia depends on the diffusible signal factor QS (DSF-QS) system, which is based mainly on the fatty acid DSF (
cis-11-methyl-2-dodecenoic acid) (
1,
2). DSF synthesis is fully dependent on RpfF, an enoyl coenzyme A hydratase encoded by the
rpf (regulation of pathogenicity factors) cluster, a set of genes that includes all of the components necessary for the synthesis and detection of DSF molecules. In addition to RpfF,
rpf encodes the aconitase RpfA, the fatty acid ligase RpfB, the two-component sensor-effector hybrid system RpfC, and the cytoplasmic regulator element RpfG (
1,
2). The DSF-QS system was first described in the phytopathogen
Xanthomonas campestris pv.
campestris, where it plays an important role in virulence regulation (
3). Since then, this system has been described in several members of the order
Xanthomonadales, including the genera
Xanthomonas,
Xylella, and
Stenotrophomonas, as well as in members of the order
Burkholderiales (
1,
3–5). The specific functions regulated by the DSF-QS system are dependent on the species, but it has been suggested that it controls several virulence-related phenotypes (
6). In the case of
S. maltophilia, little is known about the mechanisms implicated in DSF-QS regulation. It has been demonstrated that disruption of DSF signaling has a drastic effect on
S. maltophilia K279a, since the
rpfF mutant shows reduced swimming motility, reduced exoprotease production, altered lipopolysaccharide, reduced tolerance to a range of antibiotics and to heavy metals, and reduced virulence in a
Caenorhabditis elegans infection model (
1). In addition, FecA, a ferric citrate receptor, has been shown to be positively regulated by the DSF-QS system. This receptor contributes to the internalization of iron, an essential element for the expression of virulence-related genes (
7). In the
S. maltophilia WR-C wild-type (WT) strain and a flagellum-defective
xanB mutant, flagellum-independent translocation was stimulated not only by the main DSF but also by its derivative 11-methyl-dodecanoic acid (
2). Regarding the interaction of
S. maltophilia with plants, DSF seems to be involved in oilseed germination, plant colonization, and biofilm architecture in the environmental strain R551-3 (
8). Recently, the BDSF system (a DSF variant in
Burkholderia species) has also been shown to contribute to the swarming motility phenotype of
Burkholderia cenocepacia (
9).
In a recent
S. maltophilia population study, the authors detected
rpfF+ genotypes in 61% of the 89 strains tested, suggesting that an important population of
S. maltophilia lacks the
rpfF gene (
10). With the rapid increase in the number of
S. maltophilia sequenced genomes, it is now possible to compare the
rpf clusters of different strains. A preliminary analysis showed that all of the genomes sequenced contain the
rpfF gene. In addition, at least two
rpf cluster variants can be detected on the basis of sequence and genomic organization, with main differences found in the
rpfF and
rpfC genes. The genetic variation observed in the
rpfF gene translates into two distinct protein variants, here named RpfF-1 and RpfF-2. Furthermore, we can associate each of these RpfF variants with a corresponding RpfC variant, i.e., RpfC-1 and RpfC-2, respectively. We have also investigated the DSF production of representative strains from each variant group, revealing that only the strains carrying the RpfF–RpfC-1 variants show detectable DSF production under the conditions assayed. Moreover, characterization of the Δ
rpfF mutant of a strain from each RpfF variant group indicates that the virulence-related phenotypes are differently regulated in the two populations.
MATERIALS AND METHODS
Strains and growth conditions.
A panel of 78
S. maltophilia clinical isolates were collected from point prevalence studies in the intensive care units of different European hospitals. For the name, geographic origin, hospital, and isolation source of each strain, see Table S1 in the supplemental material. From this collection, E77 (RpfF-1 variant group) and M30 (RpfF-2 variant group) (
11) were used as model strains to characterize Δ
rpfF mutants (see Table S2).
Escherichia coli OP50 was provided by the Caenorhabditis Genetics Center (CGC).
X. campestris pv.
campestris 8523/pL6engGUS was obtained from the authors of reference
12.
Bacteria were routinely grown at 37°C in Luria-Bertani (LB) medium on a rotary shaker. When needed, LB was supplemented with tetracycline (Tc) at 17 μg/ml, chloramphenicol (Cm) at 3.2 μg/ml, erythromycin (Erm) at 500 μg/ml, and ampicillin (Ap) at 20 μg/ml. For phenotypic analysis in minimal medium, strains were grown in BM2 medium (62 mM potassium phosphate buffer, pH 7, 2 mM MgSO4, 10 μM FeSO4, supplemented with glucose 0.4%) or a modified M9-salts medium without NH4Cl (0.5% Casamino Acids, 2 mM MgSO4, 0.1 mM CaCl2) and supplemented with 0.2% glucose.
Sequence determination and analysis.
PCR products of 682 to 721 bp containing the
rpfF promoter and the region encoding the N-terminal fragment were amplified from all 78
S. maltophilia strains with primers PrpfFtypeUp and PrpfFTypeDw (see Table S3 in the supplemental material) and directly sequenced (Macrogen Inc.). Translation of partial open reading frames (ORFs) to amino acids and sequence alignments were done with MEGA V5.2 (
13) and BioEdit, respectively. A phylogenetic tree was constructed with MEGA V5.2 on the basis of a trimmed alignment with the 108 N-terminal residues of RpfF from strain K279a. In parallel, the genomes of strains E77, M30, and UV74 were sequenced and a first draft was constructed (to be reported upon completion). RpfC variant determination was then based on the RpfC sequences from the publicly available sequenced genomes (strains K279a, R551-3, D457, and JV3, with GenBank accession numbers AM743169.1, CP001111.1, HE798556.1, and CP002986.1, respectively) and our draft genome sequences (strains E77, M30, and UV74), by using SMART (
14) for the identification and annotation of protein domains.
Generation and complementation of ΔrpfF and ΔrpfC mutants.
For the primers and plasmids used for cloning, see Tables S3 and S4 in the supplemental material, respectively.
S. maltophilia E77 Δ
rpfF and M30 Δ
rpfF and Δ
rpfC mutants were obtained by allelic-exchange recombination with an Erm resistance cassette. Briefly,
rpfF upstream and downstream flanking regions were amplified by PCR (see Table S3 in the supplemental material) and inserted, flanking the Erm resistance cassette, into the pEX18Tc vector (
15), thus generating plasmids pEXE77
rpfF and pEXM30
rpfF for E77 and M30, respectively. Both strains were electroporated (
16) with the respective suicide vectors, and transformants were selected on LB plates containing 500 μg/ml Erm and subsequently streaked onto LB plates containing 17 μg/ml Tc to discard single-crossover events.
rpfF deletion was also verified by PCR and DNA sequencing. To generate a Δ
rpfC mutant of the M30 strain, the same strategy was used. For the primers used to amplify upstream and downstream regions of
rpfC from M30, see Table S3 in the supplemental material. Both fragments were inserted, flanking the Erm resistance cassette, into pEX18Tc, generating pEXM30
rpfC. Strain M30 was electroporated, and the mutant candidates were screened and verified with the corresponding primers (see Table S3) as described above.
A fragment of ca. 1,100 bp containing either the E77 or the M30
rpfF ORF and the predicted promoter was amplified by PCR, ligated to pBBR1MCS-Cm (
17), and introduced into E77 and/or M30 for either homologous or heterologous
trans complementation of Δ
rpfF. On the other hand, a fragment of ca. 3,000 bp was amplified from M30 and E77 to generate complementation vectors prpfGCM30 and prpfGCE77 (see Table S4), respectively. These fragments contained the
rpfG and
rpfC operon with its own promoters. Both fragments were digested with the respective restriction enzymes and ligated into pBBR1MCS1-Cm. Finally, prpfGCM30 and prpfGCE77 were introduced into E77, M30, and the M30 Δ
rpfC mutant for either homologous or heterologous
trans complementation.
Supernatant DSF extraction.
DSF extraction from culture supernatants was carried out by the ethyl acetate method (
3). Briefly, overnight bacterial cultures grown on LB medium were harvested by centrifugation and the supernatant was extracted with the same volume of ethyl acetate. The organic phase was evaporated to dryness with a rotary evaporator, and the residues were dissolved in an appropriate volume of methanol (for supernatant DSF bioassay and analysis by thin-layer chromatography [TLC]) or dichloromethane (for analysis by gas chromatography-mass spectrometry [GC-MS]).
DSF bioassay and TLC analysis.
DSF determination was performed with
X. campestris pv.
campestris 8523/pL6engGUS (DSF reporter strain) as previously described (
12), with a few modifications. Briefly, the DSF reporter strain was grown in 10 ml of NYG medium (0.3% yeast extract, 0.5% peptone, 2% glycerol) supplemented with Tc (10 μg/ml) to an optical density at 600 nm (OD
600) of 0.7. Cells were harvested, reconstituted with 1 ml of fresh NYG, added to 100 ml of cold NYG medium containing 1% BD Difco Noble agar (NYGA) supplemented with 80 μg/ml X-Glu (5-bromo-4-chloro-3-indolyl β-
d-glucuronide sodium salt; Sigma), and plated into petri plates upon solidification.
For colony-based DSF bioassays, candidate strains were pin inoculated onto plates of NYGA containing X-Glu (80 μg/ml) seeded with the DSF reporter strain and incubated for 24 h at 28°C. The presence of a blue halo around the colony indicates DSF activity.
For supernatant-based DSF bioassays, bacterial cultures were grown in 250 ml of LB for 48 h at 30°C (OD600 of about 4). Supernatants were extracted by the ethyl acetate method, and residues were dissolved in 200 μl of methanol. A 3-μl volume of each sample was deposited into a hand-generated well in a 5.5-cm plate containing NYGA supplemented with 80 μg/ml X-Glu and seeded with the DSF reporter strain to a final OD600 of 0.07. Plates were incubated for 24 h at 30°C. DSF activity was determined by the presence of a blue halo around the well.
For supernatant TLC analysis, 3-μl aliquots of dissolved methanol residues were spotted onto a silica gel 60 TLC plate (20 by 20 cm; Merck) and separated with ethyl acetate-hexane (20:80, vol/vol) as running solvents. TLC plates were subsequently air dried for at least 1 h and overlaid with 100 ml of unsolidified NYGA containing 80 μg/ml X-Glu and the DSF reporter strain at an OD600 of 0.07. TLC plates were incubated overnight at 28°C, and DSF activity was identified by the presence of blue spots.
Identification of DSF molecules from culture supernatants by GC-MS.
Bacterial cultures were grown in 2 liters of LB for 48 h at 30°C with vigorous shaking (250 rpm). Cultures were centrifuged, and supernatants were extracted by the ethyl acetate method. Dry residues were dissolved in 3 ml of dichloromethane. DSF molecules were identified by GC (Agilent Technologies 6890) with an Agilent 19091S-433 column coupled to an MS detector (Hewlett-Packard 5973).
Determination of virulence in a C. elegans model.
C. elegans CF512 [fer-15(b26)II; fem-1(hc17)IV], a strain showing temperature-dependent sterility, was provided by CGC. Nematodes were routinely maintained on NGM plates (1.7% agar, 50 mM NaCl, 0.25% peptone, 1 mM CaCl2, 5 μg/ml cholesterol, 25 mM KH2PO4, 1 mM MgSO4) seeded with E. coli OP50 at 16°C.
Determination of the virulence of
S. maltophilia strains in the
C. elegans CF512 infection model was based on the “slow killing” method (
18). Strains were grown in brain heart infusion broth overnight at 30°C, and 100 μl of each strain culture was spread onto a 5.5-cm-diameter NGM agar plate and incubated at 30°C for 24 h. Each plate was then seeded with 15 to 20 adult hermaphrodite CF512 worms, incubated at 25°C (sterility conditions), and scored for live worms every 24 h.
E. coli OP50 was used as a negative control. A worm was considered dead when it no longer responded to touch. Three replicates per strain were prepared.
Determination of virulence in a zebrafish model.
Adult (9- to 12-month-old) WT zebrafish (
Danio rerio) were subjected to a 12-h light-dark cycle at 28°C and fed twice daily with dry food. All of the fish used in infection experiments were transferred to an isolated system and acclimated for 3 days before infection. Adult zebrafish (
n = 12 per condition) were infected by intraperitoneal injection (
19) with 20 μl of a 5 × 10
8-CFU/ml suspension of each
S. maltophilia strain. The strains were previously grown at 28°C on blood agar plates (bioMérieux) for 20 h and collected directly from the plates with sterile phosphate-buffered saline (PBS). Two control groups were injected with PBS, and there were no deaths. Fish were observed daily for signs of disease and death.
One fish from each tank was sacrificed at 72 h postinfection and divided into three sections (anterior, abdominal, and posterior regions) with a sterile surgical blade. All weights were annotated, and every section was homogenized in 3 ml of PBS. After serial dilution, bacteria were plated onto LB medium containing 20 μg/ml Ap (for WT E77), LB containing 500 μg/ml Erm (for the E77 ΔrpfF mutant), or LB supplemented with Cm (for the complemented E77 ΔrpfF mutant). Finally, CFU were counted and divided per gram of tissue. All of the isolates obtained postmortem from infected zebrafish were identified as S. maltophilia on the basis of cell and colony morphology, the analytical profile index, and the 16S rRNA gene sequence (data not shown).
Biofilm formation.
To analyze biofilm formation on a polystyrene surface, 200-μl volumes of bacterial cultures grown to an OD600 of 0.1 in modified M9 or BM2 medium were inoculated into the wells of untreated 96-well microtiter plates (BrandTech 781662) and incubated for 24 h at 30°C. The plates were then washed three times with water, fixed at 60°C for 1 h, and stained for 15 min with 200 μl of 0.1% crystal violet. The dye was discarded, and the plates were rinsed in standing water and allowed to dry for 30 min at 37°C. Crystal violet was dissolved in 250 μl of 95% ethanol for 15 min, and the OD550 of the extracted dye was measured.
Biofilm formation on a glass surface was assayed by inoculating 2 ml of the same medium and adjusted OD as described above into glass tubes and incubating them for 24 h at 30°C with agitation (250 rpm). Biofilm formation was measured by crystal violet staining as described above.
Swarming assay.
Swarm agar was made on the basis of modified M9 salts medium without NH4Cl (0.5% Casamino Acids, 2 mM MgSO4, 0.1 mM CaCl2) supplemented with 0.4% glucose and solidified with 0.5% BD Difco Noble agar. Plates containing 20 ml of fresh swarm medium were dried under a laminar-flow hood for 20 min before inoculation. Inoculation was performed with a sterile Drigalski spatula containing biomass from a fresh LB plate by softly depositing it on top of a semisolid modified M9 plate. Inoculated swarm plates were sealed to maintain the humidity and incubated at 28°C for 3 to 5 days.
Quantitative reverse transcription (qRT)-PCR.
Gene expression analysis was performed to determine the ratios of
rpfF to
rpfC mRNAs in
S. maltophilia E77 and M30. Total RNA was isolated from cultures grown under the same conditions as for DSF extraction with a GeneJet RNA purification kit (Thermo Scientific), and DNA was eliminated with TURBO DNase (Ambion, Life Technologies). One microgram of RNA was used to synthesize cDNA with an iScript cDNA synthesis kit (Bio-Rad). Quantitative real-time PCR was performed with the CFX96 real-time PCR system (Bio-Rad), and PCR amplification was detected with SsoAdvanced SYBR green Supermix (Bio-Rad). PCR products of 80 to 110 bp were amplified for
rpfC,
rpfF, and
gyrA; the latter was used as an endogenous gene to normalize gene expression (
20). For the primers used, see Table S3 in the supplemental material. Differences in the relative amounts of mRNA for the
rpfF-1,
rpfC-1,
rpfF-2, and
rpfC-2 genes were determined by the 2
−ΔΔCT method (
21). RNA samples were extracted in three different experiments, and results are given as mean values.
Ethics statement.
Zebrafish were handled in compliance with Directive 2010/63/EU of the European Parliament and of the Council on the Protection of Animals Used for Scientific Purposes and with decree 214/1997 of the Government of Catalonia, which regulates the use of animals for experimental and other scientific purposes. Experimental protocols have been reviewed and approved by the Animal and Human Experimentation Ethics Committee of the Universitat Autònoma de Barcelona, Spain (reference number CEEAH-1968).
Nucleotide sequence accession numbers.
All of the amplified rpfF sequences from this S. maltophilia strain collection have been deposited in the GenBank database and assigned accession numbers KJ149475 to KJ149552.
DISCUSSION
We have characterized 78
S. maltophilia clinical strains isolated from diverse sources in different European hospitals for the
rpfF gene. We have first demonstrated that the 78 strains contain the
rpfF gene but the RpfF product is distributed into two different variants that we have named RpfF-1 and RpfF-2 (
Fig. 1 and
2A). We also show that the isolates produce two RpfC variants, each associated with one of the RpfF variants (
Fig. 2A). The two RpfC variants are different in the N-terminal region, which corresponds to a TM domain (
Fig. 2C) thought to participate in DSF sensing in several
Xanthomonas species (
12). In
S. maltophilia, the RpfC-1 variant contains 10 TM regions that display high similarity to the putative
X. campestris pv.
campestris RpfH-RpfC TM complex (
Fig. 2A and
C). On the other hand, the RpfC-2 variant has only five TM regions, which appear to be related to the
X. campestris pv.
campestris RpfH TM domain rather than that of
X. campestris pv.
campestris RpfC (
Fig. 2A). This phenomenon is also observed in
Xylella fastidiosa,
Xanthomonas oryzae, and
Pseudoxanthomonas species, suggesting that the RpfC-2 variant is widely distributed among the members of the order
Xanthomonadales that share the DSF-QS system. Nevertheless, protein sequence comparison shows a high similarity between the RpfC and RpfH TM domains, suggesting that a duplication event (for
X. campestris pv.
campestris rpfC to
rpfH and
S. maltophilia rpfC-1) or a deletion (for
S. maltophilia rpfC-2) may have occurred.
A previous
S. maltophilia population study suggested that an important group of
S. maltophilia isolates lack
rpfF (
10). PCR-based typing of 89 strains showed an
rpfF+ prevalence of 61.8%, while the remaining 38.2% were considered to be
rpfF mutants. On the basis of our sequence analysis, we can conclude that the work of Pompilio and collaborators (
10) failed to detect
rpfF because the primers they used were designed to hybridize within the most variable region of this gene; more specifically, those primers do not amplify
rpfF in strains carrying what we have defined as variant 2. Accordingly, we hypothesize that all of the
S. maltophilia strains analyzed in the study by Pompilio et al. and showing an
rpfF+ genotype belong to the RpfF-1 variant group, whereas the
rpfF mutant strains would belong to the RpfF-2 variant group. Interestingly, our analysis of
rpfF from a collection of 82
S. maltophilia strains shows similar RpfF variant frequencies in the population. RpfF-1 is present in 59.75% of the strains (including K279a and R551), whereas RpfF-2 is present in 40.25% (including D457 and JV3). Taking the two studies together (171 strains), strains carrying the RpfF-1 variant appear to be more commonly isolated than those carrying the RpfF-2 variant, with relative prevalences of ca. 60 and 40%, respectively.
Surprisingly, we have observed that only strains carrying the RpfC–RpfF-1 pair produce DSF under WT conditions, while strains belonging to the RpfC–RpfF-2 variant group require extra copies of their own
rpfF gene (
Fig. 3 and
4) or the absence of the repressor component RpfC-2 (
Fig. 5) to achieve detectable DSF production levels. These results indicate that RpfF-2 is able to synthesize DSF but the production of this signaling molecule is permanently repressed by RpfC-2 under the conditions assayed. It has been shown that the stoichiometric balance between RpfF and RpfC is crucial for DSF production in many members of the order
Xanthomonadales. In
X. campestris pv.
campestris, RpfC physically interacts with the RpfF active site, inhibiting DSF synthesis activity (
12,
22,
24). RpfC has also been shown to repress the RpfF activity of
X. fastidiosa (
25). Analysis of mRNA levels in E77 and M30 by qRT-PCR shows that the
rpfF/
rpfC expression ratio in the DSF producer strain (variant 1) is double that found in the nonproducer one (variant 2), suggesting, together with the observation that variant 2 strains complemented with extra
rpfF copies produce DSF, that the different phenotypes of the two variants may be partly due to the different regulation of the stoichiometry of these two components. It has also been suggested that in
X. campestris pv.
campestris, RpfC could play a positive-feedback role in DSF synthesis, liberating active RpfF upon the detection of DSF molecules (
22). Assuming similar mechanisms in
S. maltophilia, we hypothesize that DSF production in RpfC–RpfF-1 strains is due to the presence of a competent sensor input domain, i.e., composed of 10 TM regions, in RpfC-1, which would enable the liberation of active RpfF-1 upon DSF detection and the subsequent synthesis of DSF. On the other hand, the missing TM regions in RpfC-2 would render this factor incompetent for DSF sensing, leading to permanent inhibition of RpfF-2 by RpfC-2 in a situation of equal numbers of copies. Demonstrating that RpfC-1 liberates free active RpfF after DSF detection and understanding its mechanism or unveiling why the
S. maltophilia population produces two RpfC variants and what implications it may have for DSF-mediated regulation are questions that require further studies. The possibility that RpfC–RpfF-2 variant strains may produce DSF under specific environmental conditions or that RpfF-2 may produce a different yet undetected DSF derivative cannot be ruled out. Comparison of GC-MS spectra from M30, its Δ
rpfF mutant, and the complemented strain did not, however, reveal any peak compatible with the mass of a DSF derivative.
It is well known that the DSF-QS system regulates certain virulence traits in many bacteria (
1,
3,
9,
12,
26–29). To determine the possible implication of each RpfF variant for virulence regulation, we generated an
rpfF deletion mutant for a strain representative of each variant group, i.e., E77 for the RpfF-1 variant group and M30 for the RpfF-2 group. All of the phenotypes evaluated in M30 were unaltered in the Δ
rpfF mutant and in the corresponding complemented strain, suggesting that RpfC–RpfF-2 variant strains may not use the DSF-QS system to regulate these virulence factors, likely because of their inability to produce and sense DSF molecules under the conditions assayed. On the contrary, the E77 Δ
rpfF mutant showed attenuation in both the
C. elegans (
Fig. 8A) and zebrafish (
Fig. 9A) infection models, proving that DSF-mediated regulation affects the virulence of RpfC–RpfF-1 strains. Moreover, the recovery of bacteria from sacrificed fishes at 72 h postinjection showed that E77 is able to disseminate to the anterior and posterior regions through the fish body, while the E77 Δ
rpfF mutant had serious problems in crossing intraperitoneal barriers (
Fig. 9B). This is in concordance with the results showing a loss of swarming motility (
Fig. 6) and a drastic increase in biofilm formation capacity (
Fig. 7A and
B) by the E77 Δ
rpfF mutant, two important virulence-related traits that would explain attenuation in the animal models and especially in the zebrafish experiments. Much evidence of the implication of RpfF and DSF-like fatty acids in bacterial motility has indeed been reported (
1,
2,
9,
30). Our results thus reinforce previous evidence that one of the main functions of DSF-QS is to regulate bacterial motility. Many studies have also demonstrated the implication of DSF-like molecules in biofilm regulation. There is, however, some controversy about whether DSF-like molecules may act by stimulating or inhibiting the sessile or motile bacterial lifestyle. Thus, DSF molecules have been shown to positively regulate biofilm formation in
X. oryzae pv.
oryzae (
31),
B. cenocepacia (
9,
28), and
X. fastidiosa (
4,
26). On the contrary, in
X. campestris pv.
campestris, the DSF-mediated QS acts as a negative regulator of biofilm development (
32–34). Additionally, fatty acid-mediated biofilm dispersion is not restricted to species with the DSF-QS system. For example, the fatty acid
cis-2-decenoic acid produced by
Pseudomonas aeruginosa PAO1 stimulates biofilm dispersion in several Gram-positive and Gram-negative bacteria (
35,
36). Our findings indicate that the DSF-QS system in
S. maltophilia E77 has a regulatory function similar to that described for
X. campestris pv.
campestris, where DSF also plays an important role in preventing biofilm formation and stimulating bacterial motility.