β-Galactosidase and luminescence measurements.

Transcription of the QS genes was assayed by using PAO1-derived strains harboring promoter-lacZ fusions (Table ​(Table1).1). PAO1 reporter strains were prepared as described for pyocyanin and elastase quantification (see below). PAO1 strains (50 μl) were grown in 1 ml of LB medium at 37°C with agitation (175 rpm) supplemented with 10 μl of catechin (4 mM final) or dimethyl sulfoxide (DMSO; 1% [vol/vol], final concentration) and incubated for 8 and 18 h. After incubation, cell growth was assessed by spectrophotometry (i.e., the optical density at 600 nm [OD₆₀₀]) using a SpectraMax M2 device (Molecular Devices), and the absorbance of the medium (OD₆₀₀) after centrifugation of the bacteria (16,000 × g) was used as a blank. The sample used for cell growth assessment was used to perform the β-galactosidase assay with o-nitrophenyl-β-d-galactopyranoside as previously described (92). Promoterless lacZ fusion strains were used as controls. E. coli biosensor strains were grown in LB medium for 24 h, and 50-μl portions were subcultured for 8 h at 37°C in 1 ml of LB medium (the starting OD₆₀₀ ranged between 0.02 and 0.025 corresponding to ~5 × 10⁶ CFU) supplemented with 10 μl of DMSO (1% [vol/vol] final), 10 μl of catechin dissolved in DMSO (4 mM, final concentration), 10 μl of the appropriate homoserine lactone (10 μM, final concentration), or 10 μl of catechin (4 mM, final concentration) and the appropriate homoserine lactone (10 μM, final concentration). C4-HSL was added to pAL101 and pAL102, while 3-oxo-C12-HSL was added to pAL105 and pAL106. After incubation for 8 h at 37°C with agitation (175 rpm), 200 μl of culture was transferred to 96-well OptiPlate-96 F plates from Perkin-Elmer, and the luminescence of each sample was measured by using a TopCount NXT device from Perkin-Elmer. The LasR⁻ (pAL106) and RhlR⁻ (pAL102) biosensors were used for background subtraction, and the OD₆₀₀ values were measured to account for the differences in cell density. All experiments were performed in six replicates (n = 6). The statistical significance of each test was evaluated by conducting Student t tests using GraphPad Prism software, and P values of ≤0.05 were considered significant.

C. albiflorum sample preparation, chromatography, and electrospray ionization mass spectrometry (ESI-MS) analysis.

Dried powdered samples (5 g) of leaf and stem bark of C. albiflorum (Tul.) Jongkind (Combretaceae) were macerated overnight at room temperature in MilliQ water (25 ml). After centrifugation and filtration on Whatman paper, followed by lyophilization, the resulting residue was dissolved in 1 ml of MilliQ water and loaded onto a Sephadex LH-20 (GE Healthcare Life Sciences) column (2.5 by 7.0 cm) that was eluted successively with 100 ml of water (fractions 1 to 10), water-ethanol (9:1) (fractions 11 to 14), water-ethanol (8:2) (fractions 15 to 18), water-ethanol (1:1) (fractions 19 to 22), water-ethanol-acetone (1:0.5:0.5) (fractions 23 to 24), and water-acetone (1:1) (fractions 25 to 27). Fractions were evaporated and stored at −20°C until required for further analysis.

One active fraction (fraction 21) was further fractionated by HPLC using a reverse-phase C18 column (Symmetry 300, 2.4 by 150 mm) that was eluted with a gradient of water-methanol containing 1% acetic acid (from 0 to 100% methanol in 30 min). Analytical high-pressure liquid chromatography (HPLC) was performed by using a Waters apparatus equipped with a 626 pump, a 626 controller, and a 996 photodiode array detector. Samples were analyzed on an HPLC apparatus (Waters 600 system) coupled to both UV (Waters 2487 detector) and mass (Micromass Waters VG Quattro II mass spectrometer) detectors. A BIO Wide Pore C-18 column (250 by 4.6 mm; 5 μm) was used, and the solvent elution consisted of a linear gradient of water and acetonitrile (from 5 to 100% acetonitrile in 30 min) at a flow rate of 1 ml/min. After UV detection at 215 and 254 nm, the column eluate was split (LC Packings splitter), and 0.1 ml/min was directed to the mass spectrometer fitted with an ESI interface. For mass detection, analyte ionization was achieved by using the positive electrospray mode. The ESI parameters were as follows: nebulizing gas (N₂, 20 liters/h), drying gas (N₂, 250 liters/h), source temperature (80°C), cone voltage (35 V), and capillary voltage (35 kV). The total ion current scanning was from m/z 115 to 1,000 with 1 s/scan.

Commercially available catechin [(2R,3S)-2-(3,4-dihydroxyphenyl)-3,4-dihydro-1(2H)-benzopyran-3,5,7-triol] and epicatechin standards used for HPLC and ESI-MS analysis or for QS inhibition tests in C. violaceum CV026 or P. aeruginosa PAO1 were purchased from Sigma and dissolved in DMSO. The addition of catechin, epicatechin, or DMSO did not result in an increase of the LB broth pH (i.e., the anti-QS effect observed did not result from an opening of the lactone ring of the QS molecules through alkaline hydrolysis) (1).

Quantitative analysis of violacein production in C. violaceum CV026.

Inhibition of violacein production in C. violaceum CV026 by plant extracts (10 μl), chromatographic fractions (10 μl), or the authentic catechin and epicatechin standards (both at 4 mM final concentrations) was tested by using a liquid assay according to previously reported protocols (11, 51). Violacein production was induced in C. violaceum CV026 by adding N-hexanoyl-l-homoserine lactone (HHL; Sigma) at a final concentration of 3 μM. For violacein quantification, an overnight culture of C. violaceum CV026 was diluted 100 times in 250 μl of LB medium that were subsequently incubated for 18 h at 28°C with agitation (150 rpm) and supplemented with the appropriate sample. After assessment of bacterial growth by measuring the OD₆₀₀ by using a SpectraMax M2 device (Molecular Devices), violacein contents were quantified as described previously (11). Briefly, cells were centrifuged (16,000 × g) and washed twice with fresh LB medium. After centrifugation, cells were suspended in 200 μl of ethanol and sonicated. Cell debris were discarded by centrifugation (16,000 × g), and the absorbance (A₅₇₅) of the solution was measured. The statistical significance of each test (n = 4) was evaluated by conducting Student t tests using the GraphPad Prism software, and a P value of ≤0.01 was considered significant.

Quantitative analysis of pyocyanin and elastase production in P. aeruginosa wild-type and mutant strains.

Inhibition of pyocyanin and elastase production was assessed according to previously described procedures (34, 55). P. aeruginosa PAO1 or mutant cells (single colony from LB agar plates) were grown for 18 h in 5 ml of LB-MOPS medium (37°C and agitation at 175 rpm). The cells were washed twice in fresh LB-MOPS medium to eliminate the homoserine lactones, and the pellets were suspended in LB-MOPS medium. Then, 50-μl portions of the cell suspension were added to 1 ml of LB-MOPS (in order to obtain a starting OD₆₀₀ of the LB-MOPS medium ranging between 0.020 and 0.025 and corresponding to ~10⁷ CFU/ml) supplemented with 10 μl of plant extract or authentic catechin standard (4 mM [final concentration] or as stated in the text) dissolved in DMSO and, when appropriate, with 3-oxo-C12-HSL or C4-HSL. After 8 h of growth, samples were taken to assess the growth (OD₆₀₀). After centrifugation (16,000 × g), the absorbance at 600 nm of the medium was used as blank, and 900 μl was used for pyocyanin determination (A₃₈₀). Elastase production was assessed through the measurement of elastase activity using elastin-Congo red (A₄₉₅). The statistical significance of each test (n = 4) was evaluated by conducting Student t tests using GraphPad Prism software, and a P value of ≤0.01 was considered significant.

Biofilm formation and quantification.

P. aeruginosa PAO1 was grown overnight in LB medium at 37°C with agitation. After growth, the culture was diluted with TB (tryptone broth) medium (OD₆₀₀ of ~0.02), and 50 μl of the diluted culture was added to 940 μl of TB medium supplemented with 10 μl of DMSO or catechin (4 mM, final concentration). PAO1 cells were incubated statically for 18 h at 37°C in 24-well polystyrene plates. After incubation, planktonic bacteria were discarded, and the biofilms were washed three times with phosphate-buffered saline buffer. Washed biofilms were fixed with 1 ml of methanol (99%). After 15 min, the methanol was discarded, and the plates were dried at room temperature. Crystal violet (0.1% in water) was then added to each well (1 ml/well), and the plates were incubated for 15 min at room temperature. Crystal violet was then discarded, and stained biofilms were washed three times with 1 ml of water. Acetic acid (33% in water) was added to the stained biofilms (2 ml) in order to solubilize the crystal violet, and the absorbance of the solution was read at 590 nm with a SpectraMax M2 device (Molecular Devices). The statistical significance of each test (n = 6) was evaluated by conducting Student t tests and using the GraphPad Prism software; a P value of ≤0.01 was considered significant.

Chromatographic fractionation of the C. albiflorum bark extract and identification of flavonoid-containing active fractions.

In order to identify the inhibitory compound(s) responsible for the QS inhibition activity, the C. albiflorum aqueous bark extract was subjected to gel filtration chromatography (Fig. ​(Fig.2A).2A). Twenty-seven fractions were collected, and the last seven eluted fractions (fractions 21 to 27) were found to inhibit violacein production (Fig. ​(Fig.2A).2A). The first active fraction (fraction 21) was further fractionated, using reversed-phase HPLC (RP-HPLC), into 19 subfractions, and subfractions F4 to F16 inhibited violacein production (Fig. ​(Fig.2B).2B). Fraction 21 was chosen because (i) preliminary thin-layer chromatography showed that this fraction contained few different compounds compared to the following fractions, (ii) growth of the bacteria was not affected, and (iii) the amount of residue obtained allowed the further chemical characterization of the active compound(s) (data not shown). The UV spectra of these subfractions showed that most of the peaks presented typical flavonoid-type UV spectra profiles, i.e., two main absorption bands comprised between 240 to 270 nm and 320 to 380 nm, respectively (31; data not shown). These data suggest that the active fractions of the C. albiflorum bark extract contain flavonoid-like structures interfering with violacein production in C. violaceum CV026 without any bactericidal or bacteriostatic activity.

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FIG. 2.

Chromatographic scheme of C. albiflorum bark extract fractionation steps used to isolate and identify the anti-QS compounds and identification of catechin as the main compound present in the active fraction F7. (A) Gel filtration fractionation of the crude C. albiflorum bark extract. The column was successively eluted with water, water-ethanol, and water-acetone, and 27 fractions were collected. These fractions were added to HHL-induced C. violaceum CV026, and inhibition of violacein production was observed starting from fraction 21. (B) RP-HPLC profile of fraction 21 (280 nm). Nineteen subfractions were collected and tested on HHL-induced C. violaceum CV026, and inhibition of violacein production was observed with fractions F4 to F16. AU, arbitrary unit. (C) UV spectrum of fraction F7. (D) UV spectrum of an authentic standard of catechin showing the characteristic absorption maximum of catechin at 278 nm. (E) Chemical structure of catechin with the two benzene cycles connected by the heterocycle typical of flavonoids. (F and G) ESI-MS of fraction F7 (F) and of the authentic standard of catechin (G) with the main peaks (Da/e) at 123, 139, and 291.

Catechin is one of the C. albiflorum flavonoid inhibiting the production of violacein in C. violaceum CV026 and pyocyanin in P. aeruginosa PAO1.

The peak purity of the HPLC subfractions F4 to F16 was assessed using the Waters Millennium chromatography manager. The presence of a single peak in subfractions F7 (with a retention time of 11 min) and F9 (with a retention time of 13 min) prompted us to further investigate the structural characterization of the QS inhibitory compound present in these subfractions. First, as shown in Fig. 2C and D, the UV spectrum of the major peaks in subfractions F7 is similar to that of catechin (of which the structure is given in Fig. ​Fig.2E).2E). Second, analysis of a standard of catechin in the same HPLC conditions showed that catechin has the same retention time as the compound present in subfraction F7 (data not shown). Third, coinjection of the catechin standard with subfraction F7 showed no difference in the retention times, which further supported the presence of catechin in this subfraction (data not shown). Fourth, the ESI-MS analysis indicated that the MS spectrum of subfraction F7 was similar to that of authentic catechin standard showing an identical quasimolecular ion at m/z (Da/electron [e]) 291 with ion at 139 and 123 (Fig. 2F and G, respectively) in agreement with data reported previously (22). Epicatechin, which is the epimer of catechin, has the same UV and mass spectra but is found in fraction F9, as demonstrated by HPLC separation of an epicatechin standard (which had a retention time of 13 min, as did fraction F9) and coinjection of epicatechin and fraction F9 in the same HPLC conditions (data not shown). Finally, the catechin standard (Fig. ​(Fig.2E)2E) inhibited violacein production when added to HHL-induced liquid cultures of C. violaceum CV026, as did subfraction F7, further confirming that catechin is the QS inhibitory compound. Indeed, as shown in Fig. ​Fig.3,3, serial dilutions of the catechin standard were tested on HHL-induced cultures of C. violaceum CV026 in order to determine the range of concentrations inhibiting violacein production with a limited effect on bacterial growth (Fig. 3A and C). When concentrations of catechin higher than 4 mM were used, C. violaceum CV026 growth was strongly reduced and, as a consequence, no violacein production occurred (Fig. 3A and C). From 0.25 to 4 mM, violacein production was significantly affected by catechin (Fig. ​(Fig.3A),3A), an effect that was not associated with a decrease in growth of C. violaceum CV026 (Fig. ​(Fig.3C).3C). Serial dilutions of catechin were also tested on P. aeruginosa PAO1. As shown in Fig. ​Fig.3B,3B, concentrations between 0.125 and 16 mM had a significant negative effect on pyocyanin production without affecting P. aeruginosa PAO1 growth (Fig. ​(Fig.3D).3D). The epicatechin standard, tested at 4 mM, had an inhibitory effect on pyocyanin production in P. aeruginosa PAO1, as did subfraction F9, while this compound had a bactericidal effect on C. violaceum CV026 (data not shown). The effect of catechin and epicatechin (at 4 mM) was also tested on elastase production in P. aeruginosa PAO1 by performing an elastolysis assay as described in Material and Methods. As shown in Fig. ​Fig.4A,4A, after 8 h, the addition of catechin (as well as epicatechin) reduced the production of elastase by ca. 30%. For comparison, the production of pyocyanin by the ΔPA3476 (ΔrhlI) and ΔPA3477 (ΔrhlR) mutants was lower than in the wild-type strain PAO1 by 69% ± 6% and 73% ± 4%, respectively, whereas the production of elastase (as measured by the elastolysis assay) by the ΔPA1432 (ΔlasI) and ΔPA1430 (ΔlasR) mutants was lower than in the wild-type strain PAO1 by 53% ± 5% and 76% ± 6%, respectively.

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FIG. 3.

Effect of serial dilutions of an authentic standard of catechin on violacein (A) and pyocyanin (B) production in C. violaceum CV026 and P. aeruginosa PAO1, respectively. (C and D) Growth of both bacteria assessed at 600 nm. Violacein and pyocyanin were extracted as described in Materials and Methods and quantified by reading the OD values of the solution at 575 nm (A) and 380 nm (B), respectively. DMSO-supplied cultures were used as controls. The statistical significance of each test (n = 4) was evaluated by conducting Student t tests, and a P value of ≤0.01 was considered significant. ***, Data that are statistically different (P ≤ 0.01).

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FIG. 4.

Effect of authentic standards of catechin (4 mM) and epicatechin (4 mM) on elastase production (A), biofilm formation (B), P. aeruginosa PAO1 viability after 8 h (C) and 18 h (D) of culture, and P. aeruginosa PAO1 growth kinetics (E). DMSO-supplied cultures were used as controls. Elastase production by P. aeruginosa PAO1 (A) was quantified through an elastolysis assay as described in Materials and Methods. No difference in cell densities (OD₆₀₀) was observed between the different treatments (data not shown). Elastase production by the ΔlasR mutant strain PA1430 was used as a comparison to assess the potency of catechin. (B) Biofilm formation by P. aeruginosa PAO1 after 18 h of incubation was assessed by using crystal violet. (C and D) The viability of P. aeruginosa PAO1 cells was assessed by using SYTO-9 and PI as described in Materials and Methods after 8 h (C) and 18 h (D) of culture. DMSO-supplied cultures were used as controls, and their viability was used to plot the viability of catechin- and epicatechin-treated cultures. The statistical significance of each test (n = 6) was evaluated by conducting Student t tests, and a P value of ≤0.01 was considered significant. ***, Data that are statistically different (P ≤ 0.01). (E) Growth kinetics of DMSO- and catechin-treated P. aeruginosa PAO1. The growth kinetics were divided into two phases: phase I was between 0 and 6 h (corresponding to the exponential growth phase), and phase II was between 6 and 20 h (corresponding to the stationary phase). Doubling times were calculated for both phases by using GraphPad Prism software (n = 6). Arrows indicate the time points at which samples were taken for pyocyanin, elastase, biofilm, or gene expression analysis.

Effect of catechin on biofilm formation in P. aeruginosa PAO1.

P. aeruginosa PAO1 can switch to a biofilm mode of growth. Biofilms are matrixes of polysaccharides in which bacteria are enmeshed and protected from the environment, increasing their resistance to antibiotics and the immune system, thereby increasing their capacity to remain in the infected host (15, 56). Since biofilm formation is partially controlled by QS mechanisms (16), the effect of catechin and epicatechin on P. aeruginosa PAO1 biofilm formation was assessed after 18 h of growth. As shown in Fig. ​Fig.4B,4B, catechin reduced biofilm formation by ca. 30%, while epicatechin had no effect. To determine whether the effect of both compounds on pyocyanin, elastase, and biofilm was due to a drop in cell viability, P. aeruginosa PAO1 viability was assessed after 8 and 18 h. As shown in Fig. 4C and D, none of the compounds had an effect after 8 h of incubation (Fig. ​(Fig.4C)4C) but, after 18 h, epicatechin-treated P. aeruginosa PAO1 cells had a viability of 66%, whereas catechin-treated cells were not affected in their viability compared to DMSO-treated cells (Fig. ​(Fig.4D).4D). Consistently, catechin had no significant effect on P. aeruginosa PAO1 growth parameters, as shown in Fig. ​Fig.4E.4E. Indeed, the doubling times for control and catechin-treated P. aeruginosa PAO1 cells were, respectively, 0.94 ± 0.13 and 0.92 ± 0.03 h during the first phase of the kinetics (phase I) and 8.02 ± 0.24 and 7.46 ± 0.30 h during the second phase of the kinetics (phase II).

O.M.V. is a Postdoctoral Researcher of the FRS-FNRS (Fonds de la Recherche Scientifique, Belgium). M.B. is a Senior Research Associate of the FRS-FNRS. M.K. is indebted to the Coopération Universitaire pour le Développement-Coopération Universitaire Institutionnelle. S.R. is indebted to the Agence Universitaire de la Francophonie for a predoctoral fellowship.

We thank Barbara Iglewski from the Rochester University School of Medicine and Dentistry (United States) for kindly providing plasmids pLP170, pPCS223, pPCS1001, pLPR1, and pPCS1002; Junichi Kato from the Department of Molecular Biotechnology, Hiroshima University, Hiroshima, Japan, for kindly providing plasmids pQF50, pβ01, pβ02 and pβ03; Brian Ahmer from Ohio State University for providing the E. coli biosensor strains pAL101, pAL102, pAL105, and pAL106; and Frédéric Paulart from the Institute for Medical Immunology from the Université Libre de Bruxelles for helping with the use of the TopCount NXT.