Research Article
24 July 2013

Induction of Biofilm Formation in the Betaproteobacterium Burkholderia unamae CK43B Exposed to Exogenous Indole and Gallic Acid

ABSTRACT

Burkholderia unamae CK43B, a member of the Betaproteobacteria that was isolated from the rhizosphere of a Shorea balangeran sapling in a tropical peat swamp forest, produces neither indole nor extracellular polymeric substances associated with biofilm formation. When cultured in a modified Winogradsky's medium supplemented with up to 1.7 mM indole, B. unamae CK43B maintains its planktonic state by cell swelling and effectively degrades exogenous indole. However, in medium supplemented with 1.7 mM exogenous indole and 1.0 mM gallic acid, B. unamae CK43B produced extracellular polymeric substances and formed a biofilm. The concentration indicated above of gallic acid alone had no effect on either the growth or the differentiation of B. unamae CK43B cells above a certain concentration threshold, whereas it inhibited indole degradation by B. unamae CK43B to 3-hydroxyindoxyl. In addition, coculture of B. unamae CK43B with indole-producing Escherichia coli in nutrient-rich Luria-Bertani medium supplemented with 1.0 mM gallic acid led to the formation of mixed cell aggregates. The viability and active growth of B. unamae CK43B cells in a coculture system with Escherichia coli were evidenced by fluorescence in situ hybridization. Our data thus suggest that indole facilitates intergenus communication between indole-producing gammaproteobacteria and some indole-degrading bacteria, particularly in gallic acid-rich environments.

INTRODUCTION

Although most microorganisms are traditionally characterized and identified as planktonic, in certain environments they may also form sessile communities known as biofilms. The term biofilm refers to an aggregation of bacteria, algae, fungi, and protozoa enclosed in an envelope composed of a mixture of extracellular polysaccharides (EPS) that promote adherence to surfaces or interfaces (13). Bacteria can transit from a planktonic existence and develop biofilms on both abiotic and biotic surfaces, such as natural aquatic and soil environments, living tissues, and medical or industrial devices (2). Biofilms serve to protect the microbial community from a diverse array of stressors. In particular, biofilms formed by human pathogens and bacterial communities in the rhizosphere enable the bacteria to resist bacteriophages, chemically diverse biocides, host immune responses, and antibiotics (4, 5). Biofilm formation is regulated by a variety of environmental factors, such as pH, the availability of nutrients and oxygen, and the concentration of bacterial metabolites (6, 7).
Bacteria of the subclass Betaproteobacteria produce various N-acyl-l-homoserine lactones (AHLs) as quorum-sensing signals. These compounds stimulate biofilm formation, allowing the bacteria to regulate the expression of specific genes in association with EPS production and functional differentiation in the biofilm (8, 9). In some gammaproteobacteria, AHLs control EPS production and biofilm formation in conjunction with autoinducer-2 (AI-2), whereas AI-2 sometimes shows inconsistency in prevention of biofilm development (10, 11). In Escherichia coli, the key metabolite indole functions as an extracellular signal that regulates the expression of several important genes (12, 13). Specific cellular responses, such as chemical resistance and biofilm formation, are induced when E. coli and other indole-producing gammaproteobacteria are exposed to certain levels of indole (14, 15).
Indole plays a role in AHL-mediated species-specific quorum sensing in Gram-negative bacteria, where it acts as an interspecies communication signal that inhibits biofilm formation of E. coli while stimulating biofilm formation of Pseudomonas spp. (16). Such bacterial relationships via interspecies communication, particularly those between different proteobacterial subclasses, are rarely studied. Accordingly, it is still unclear whether the exogenous indole contributes to such interspecies communication between indole-producible and indole-degradable bacteria, including the induction of EPS production within the dual-species bacterial habitat.
We previously reported the isolation of some l-tryptophan-degrading bacteria from the rhizosphere of wild Shorea saplings growing in peatland in Central Kalimantan, Indonesia (17, 18). Also, we recently reported a cometabolic effect of gallic acid and some other pyrogallol-type polyphenols on aerobic indole degradation by Burkholderia unamae (19). Conversely, gallic acid suppressed indole degradation by static-cultured B. unamae CK43B. In the presence of exogenous indole (0.5 to 1.7 mM), B. unamae CK43B that was exposed to appropriate concentration of gallic acid showed biofilm formation alone with EPS production. In the present report, we show that B. unamae CK43B isolated from Shorea balangeran degrades indole and produces an oxygenated indole (3-hydroxyindoxyl), particularly in nitrogen-poor media, and we also demonstrate that a mixture of gallic acid and indole induces biofilm formation in B. unamae CK43B, suggesting that indole acts as a biologically active compound to some non-indole-producing bacteria.

MATERIALS AND METHODS

General.

Glass plates (Merck Kieselgel 60F254, 0.25-mm thickness; Merck, Darmstadt, Germany) were used for preparative and analytical thin-layer chromatography (TLC). Crystalline indole was obtained from Takeda (Osaka, Japan). Gallic acid, pyrogallol, (+)-catechin, and standard (–)-epigallocatechin were purchased from Wako Pure Chemical Industries (Osaka, Japan). Crystal violet was purchased from Kanto Chemical (Tokyo, Japan). 1H nuclear magnetic resonance (NMR) spectra were recorded on a JEOL EX-270 (JEOL, Tokyo, Japan) operated at 270 MHz. Deuterated NMR solvents (CD3OD and CDCl3) were obtained from Wako Pure Chemical Industries.

Culture media.

As a nitrogen-poor medium, we used modified Winogradsky's (MW) medium. Winogradsky's mineral solution with an optional concentration of sucrose is used for the cultivation of diazotrophic bacteria, so the salt mixture does not contain any nitrogen sources. MW medium is a Winogradsky's mineral solution with 5 g of sucrose liter−1 that is supplemented with 50 mg of yeast extract liter−1 for a minimum nitrogen source (22). With appropriate concentrations of indole (e.g., 1.7 mM) as an additional nitrogen source to the MW medium, B. unamae CK43B cells grew well under static culture conditions. Another medium used is Luria-Bertani (LB) broth. Since both B. unamae CK43B and E. coli K-12 grew well, LB medium was used to coculture B. unamae CK43B with E. coli K-12 IFO 3301. Both bacteria precultured on LB plates were, respectively, suspended into 1 ml of sterile water, of which an aliquot 25 μl was inoculated to 2 ml of LB broth in polystyrene petri dishes (35 × 10 mm; BD Bioscience, San Jose, CA) as a final optical density at 600 nm (OD660) of 0.02.

Bacterial characterization, identification, and metabolic properties.

B. unamae strain CK43B was originally isolated from the rhizoplane of a wild dipterocarpous tree sapling growing in tropical peatland in Central Kalimantan, Indonesia (20, 21). B. unamae CK43B was identified based on its 16S rRNA gene sequence through homology searches using the National Center for Biotechnology Information database, and the sequence was deposited in the DNA database of Japan (DDBJ) under accession number AB714631. In l-Trp containing MW medium, the shake-cultured bacterium metabolized l-Trp to tryptophol or catechol (21). When appropriate, a concentration of indole was added to the medium instead of l-Trp, B. unamae CK43B converted indole to 3-hydroxyindoxyl.
In LB medium, B. unamae CK43B showed remarkable cell growth, whereas an indole-producing Escherichia coli K-12 formed biofilm (23). Therefore, the biofilm-forming E. coli K-12 IFO 3301 was tested for coculturing with B. unamae CK43B to examine whether E. coli K-12 maintains EPS production in the presence of indole-degrading B. unamae CK43B. Coculture was done without shaking at 28°C for 24 h. In addition, the effect of supplemented gallic acid (1 mM) on biofilm formation of the cocultured B. unamae CK43B with E. coli K-12 was examined under the same culture conditions.

Biofilm formation assay.

Modified Winogradsky's mineral solution served as the basal medium (MW) for assessing biofilm formation, but in MW medium B. unamae CK43B did not grow at all. To examine the effect of exogenous indole on biofilm formation of B. unamae CK43B, MW medium was supplemented with 1.7 mM indole (200 mg liter−1) as a nitrogen source or 1.7 mM indole plus 1.0 mM gallic acid as an indole degradation inhibitor. All the medium were adjusted to pH 5.0 using a 4 M aqueous H2SO4 prior to autoclaving. The bacterium precultured on agar was suspended in sterilized water, an aliquot 25 μl of which was inoculated to 2 ml of liquid medium in polystyrene petri dish, and static cultured for 48 h at 28°C in the dark to allow biofilm formation at the flat bottom.
Biofilm formation on the inner surfaces of the petri dishes was observed under a phase-contrast mode on a microscope (Biorevo BZ-9000; Keyence, Osaka, Japan), combined with the crystal violet staining method described by Merritt et al. (22). Bacterial growth was assessed using a microplate reader (Sunrise; Tecan, Männedorf, Switzerland) by monitoring the OD660 of the samples. The biofilms adhering to the bottom of the dish stained with a 0.1% crystal violet solution for 10 min, submerged in tap water to remove excess dye, and air-dried were observed with a light microscope under ×4, ×20, and ×40 objective lenses. Next, 1 or 2 ml of 30% acetic acid was added to the petri dish to solubilize the blue pigment from the stained biofilm. A predetermined volume (200 μl) of the supernatant in a 96-well plate was used to measure the absorbance at 570 nm (A570). Quantification of the biofilm was expressed by dividing the A570 by the OD660 to normalize the value with the cell population. Bacterial cells stained with 2% crystal violet were also observed under a light microscope (Olympus IX70; Olympus Co., Tokyo, Japan) using an oil immersion technique with a ×100 objective lens.

Effect of indole and polyphenols on growth and biofilm formation of B. unamae CK43B.

The effect of indole (0 to 5.0 mM final concentration) on the growth of B. unamae CK43B in MW medium was assessed. The bacterium inoculated into 5 ml of MW medium in 18-cm test tubes was incubated at 28°C for 48 h with shaking at 120 rpm. Growth was monitored by determining the OD660 using a U-3310 UV-Vis spectrophotometer (Hitachi, Tokyo, Japan). By way of comparison to indole, another N-heterocyclic aromatic compound pyridine was examined under the same conditions.
To determine the effect of gallic acid and some other polyphenols on the biofilm formation, 20 μl of 0.2-μm-pore-size-filter-sterilized solution of gallic acid, pyrogallol, (+)-catechin (each 1.0 mM as a final concentration), or (−)-epigallocatechin (0.5 mM) was added to a 2-ml culture of B. unamae CK43B, with or without 1.7 mM indole. The biofilm formation after 2 days of incubation was quantitatively assessed as described above.

Cell viability assay for monocultured B. unamae CK43B in MW medium.

Cell viability assay was done using Live/Dead BacLight bacterial viability kit L13152 (Life Technologies Co., Carlsbad, CA) according to the manufacturer's instructions. To observe effect of exogenous indole and gallic acid on B. unamae CK43B, an aliquot 10 μl of B. unamae CK43B cell suspension (OD660 = 0.1 to 0.2) was inoculated into 2 ml of MW medium with or without 1 mM gallic acid in a polystyrene petri dish (diameter, 35 by 10 mm; Falcon) without shaking at 28°C for 24 to 48 h. Planktonic cells were collected by brief spinning, and the resulting cell pellet resuspended into 100 μl of a 0.85% NaCl solution and spun repeatedly five times was subjected to a viability assay. For the biofilm-forming cells, the planktonic cells were removed by decantation and washing them out with 0.85% NaCl solution several times. The remaining biofilm on the bottom of the petri dish was scraped off using pipette tips, suspended in 0.85% NaCl, and centrifuged at 10,000 × g for 10 min. This process was also repeated several times. Viability was observed under a fluorescence microscope (Biorevo BZ-9000) equipped with filter sets for monitoring living cells stained by SYTO9 (excitation, 470/40 nm; emission, 535/50 nm; with a green fluorescent protein [GFP]-BP filter for green fluorescence) and for monitoring dead cells by using propidium iodide (excitation, 540/25 nm; emission, 605/55 nm; with a tetramethylrhodamine isothiocyanate [TRITC] filter for yellow-orange color and finally displayed it as a red false-color image).

FISH for biofilm formed by coculture of B. unamae CK43B with E. coli.

A fluorescence in situ hybridization (FISH) experiment was performed for cocultured B. unamae CK43B with E. coli in LB broth medium containing 1 mM gallic acid or without any polyphenolic additives in a polystyrene petri dish (35 by 10 mm) at 28°C for 20 or 24 h, according to the procedures described by Hori et al. (23). The cocultured broth medium containing only planktonic cells (1 ml) was transferred to a 1.5-ml Eppendorf tube and spun briefly to collect the planktonic cells, and the pellet was washed twice with 1× phosphate-buffered saline (PBS; NaCl, 50 g liter−1; NaHPO4·2H2O, 25 g liter−1; KCl, 25 g liter−1; KH2PO4, 1 g liter−1 [pH 7.4]). Conversely, slimy colonies that remained on the bottom of the petri dishes of the biofilm-formed culture were scraped off using pipette tips, suspended in 0.85% NaCl, and centrifuged at 10,000 × g for 10 min. To remove the polymeric or capsular material that may prevent hybridization, the biofilm-formed cells were resuspended in 2 M NaCl, gently shaken, and spun briefly. The resulting cell pellets washed twice with 1× PBS were fixed with 4% paraformaldehyde (wt/vol; pH 7.4) for the FISH experiment (23).
The FISH probes used in the present study were purchased from Chromosome Science Labo, Inc. (Sapporo, Japan), as follows: BKH70, 5′-TGC CAT ACT CTA GCG ATG-3′ labeled with Cy3 for B. unamae CK43B; ENT186, 5′-CCC CCT CTT TGG TCT TGC-3′ labeled with Cy5 for E. coli; and EUB338, 5′-GCT GCC TCC CGT AGG AGT-3′ labeled with fluorescein isothiocyanate (FITC) for all eubacteria. For hybridization, a 1-μl aliquot of each probe solution (three probe solutions) was added to 17 μl of hybridization solution. The pellets of the fixed cells resuspended in 20 μl of the hybridization solution were incubated at 46°C for 2 h. After the incubation, excessive probes were remove with wash buffer (30% formamide, 0.9 M NaCl, 0.01% sodium dodecyl sulfate, 20 mM Tris-HCl [pH 7.2]) by further incubation at 46°C for 10 min. The resulting pellet washed twice with 1× PBS was suspended in 10 to 40 μl of 1× PBS. For fluorescence microscopic observation, an aliquot (2 μl) of the suspension was dropped onto a slide glass plate (Micro Slide Glass S1111 for fluorescence microscopic observation, 0.8 to 1.0 mm thick) and sealed under a cover glass.
The in situ-hybridized cells were observed using a Biorevo BZ-9000 fluorescence microscope (objective lens, ×100; field lens, ×15) with oil immersion. The FISH probe EUB338 used for all bacteria labeled with FITC combined with a GFP-BP filter (excitation, 470/40 nm; emission, 535/50 nm) fluoresces green, whereas the probe ENT186 used for E. coli K-12 labeled with Cy5 combined with a Cy5 filter (excitation, 620/60 nm; emission, 700/75 nm) showed a red color. Since color photographs in which EUB338 and ENT186 merged showed a yellowish orange color, BKH70 for B. unamae CK43B labeled with Cy3 originally fluoresces an orange color via a TRITC filter (excitation, 540/25 nm, emission, 605/55 nm) was displayed as blue false-color images (see Fig. S1 in the supplemental material). The samples illuminated with fluorescence were also compared to those observed under visual light.

Quantification of indole and identification of indole metabolites.

A 2-ml aliquot of MW culture medium was centrifuged at 15,650 × g for 10 min. Supernatants that were adjusted to pH 4.0 with a small volume of 1 M HCl were extracted twice with an equivalent volume of ethyl acetate (EtOAc), and the resulting organic layer was dried over anhydrous Na2SO4, concentrated, and redissolved in 2 ml of acetonitrile. Finally, 5 μl of 1.0 mM 2,7-naphthalenediol was added as an internal standard. Indole remaining in cultured MW medium that contained gallic acid was quantified using high-performance liquid chromatography (HPLC). The HPLC system consisted of an L-6320 intelligent pump, an L-4250H UV-Vis detector at 280 nm, and an D-2500 Chromato-Integrator (Hitachi, Tokyo, Japan). The pump was connected to a reversed-phase column (Inertsil PREP-ODS, 10 μm, 6.0 mm [inner diameter] by 250 mm; GL Science, Inc., Torrance, CA), which was eluted with an isocratic mobile phase consisting of acetonitrile-H2O (50:50) containing 0.1% (vol/vol) formic acid, and the eluate was monitored at 280 nm. The injection volume was 5 μl and the flow rate was 1.0 ml min−1. The standard curve constructed using the ratio of the peak intensity for indole to that of the internal standard was highly linear (r2 = 0.99; y = 0.1653x − 0.0006, where y is the indole concentration (in mM) and x is the peak intensity ratio [indole/internal standard]). Using this analytical system, indole production by B. unamae CK43B in the culture fluid in LB medium was measured, together with indole content in plain LB broth (blank).

Identification of indole metabolites.

To identify major indole metabolite converted from supplemented indole, B. unamae CK43B was shake cultured in 1,000 ml of MW medium supplemented with 1.7 mM indole for 48 h at 28°C at 120 rpm, and a mixture of the oxygenated indoles (ca. 150 mg) was extracted with EtOAc from the resulting culture fluid. Using a half of the mixture (74 mg), the target compound, also predominantly yielded in the static cultured medium, was purified by silica gel column chromatography and preparative TLC as we described earlier (19). The chemical structure of the compound was elucidated by 1H-NMR and mass (FD- and FD-HR-MS) spectroscopic analyses to be identical to 3-hydroxyindoxyl (67 mg of colorless powders): 3-Hydroxyindoxyl. UV (CHCl3) λmax (methanol [MeOH]): 210, 254, 295 nm; FD-MS (relative intensity %): m/z 149 ([M]+, 100). HR-FD-MS: found 149.0504 (C8H7NO2, calculated for 149.0477). 1H-NMR (δH, in CDCl3): 7.47 (1H, d, J = 7.6 Hz, 4-H), 7.25 (1H, t-like m, 6-H), 7.09 (1H, t, J = 7.6 Hz, 5-H), 6.86 (1H, d, J = 7.8 Hz, 7-H), 5.08 (1H, d, J = 4.6, 3-H), and 2.94 (1H, br d, 3-OH). 3-Hydroxyindoxyl thus obtained was used as the original solution for a series of diluted solutions (10 to 250 μM).

Assessing the effect of 3-hydroxyindoxyl on planktonic cell and biofilm formation of B. unamae.

Effect of 3-hydroxyindoxyl at different concentrations (0 to 250 μM) was examined on B. unamae CK43B cultured in LB medium at 28°C for 24 h without shaking. Cell growth was assessed by monitoring the OD660 with a CENios microplate reader (Tecan Genios, Männedorf, Switzerland). Biofilm formation was evaluated using the crystal violet staining method according to the same procedure as described above.

Statistical analysis.

Values are expressed as means ± the standard deviations (SD). Multiple comparisons were carried out using the Student t test and one-way analysis of variance (ANOVA). Treatment effects were analyzed using Duncan's multiple-range test and SPSS 18.0 software (IBM, Armonk, NY). Differences were considered to be significant when the P value was <0.05.

RESULTS AND DISCUSSION

Biofilm induction in B. unamae cells treated with indole and gallic acid.

Using HPLC, it was confirmed that B. unamae CK43B does not produce any trace amounts of indole, even in LB broth that is rich in l-Trp. Conversely, B. unamae CK43B static-cultured in indole-supplemented MW medium actively degraded exogenous indole to 3-hydroxyindoxyl (Fig. 1), which inhibited static culture growth at 50 to 100 μM, making it difficult to assess biofilm formation. Along with these traits, B. unamae CK43B did not produce any EPS, irrespective of the presence or absence of an excessive amount of supplemental indole (1.7 mM) in MW liquid medium. The unique morphological differentiation of B. unamae CK43B under exposure to this high concentration of indole was observed as development of vacuole-like large granules inside the cell body (see Fig. S2, the right two panels in the top row, in the supplemental material). Only when an appropriate concentration of gallic acid (1.0 mM) was also added to MW medium supplemented with indole was biofilm formation accelerated in B. unamae CK43B (Fig. 2), along with slight cell swelling (see Fig. S2, right in top panels, in the supplemental material).
Fig 1
Fig 1 Detection of indole metabolite 3-hydroxyindoxyl and unmetabolized indole in the culture of B. unamae CK43B in MW supplemented with 1.7 mM indole in the presence or absence of gallic acid. Static culture of B. unamae CK43B in 1.7 mM indole-containing MW medium at 28°C for 48 h. Control, without polyphenol additive. +1.0 mM gallic acid, gallic acid further supplemented as a polyphenolic additive; +1.0 mM pyrogallol, supplementation of pyrogallol instead of gallic acid as a negative control. HPLC profiles are indole metabolite extracted from each culture fluid with EtOAc. The peaks at 4.6, 6.3, and 14.0 min are of 3-hydroxyindoxyl (open arrows), internal standard (Int. std.; 2,7-naphthalenediol, marked with asterisks) and unchanged indole (filled arrow), respectively. These HPLC profiles clearly show that addition of 1.0 mM gallic acid uniquely inhibited oxidation of indole into 3-hydroxyindoxyl in the static culture.
Fig 2
Fig 2 The effect of supplemented indole and gallic acid on biofilm formation by B. unamae CK43B in MW liquid medium. Turbidity was measured as the OD660 to indicate cell growth (left columns), and biofilm formation was measured as the A570 due to crystal violet staining of the biopolymer (right columns). B. unamae CK43B grew well in MW medium supplemented with indole, and the medium further supplemented with gallic acid showed turbidities (OD660) greater than 0.25 and 0.1, respectively. In MW liquid medium without supplemented indole, B. unamae CK43B allowed neither cell growth nor biofilm formation, irrespective of the presence or absence of 1.0 mM gallic acid. Statistic significance was calculated by using the Student t test. The data represent the mean ± the SD (n = 3). *, P < 0.001 relative to the control (+ indole only). , Plus indole at 1.7 mM; □, plus indole at 1.7 mM and gallic acid at 1.0 mM.
The appearance of the B. unamae CK43B colonies that developed on the bottom of the plastic petri dishes under these conditions was obviously different. In the medium containing both indole and gallic acid, cell aggregates formed flat biofilms, as reported for Pseudomonas aeruginosa (24, 25). Supplementation with 1 mM gallic acid in the absence of indole did not induce biofilm formation by B. unamae CK43B. Furthermore, the addition of 1.0 mM gallic acid to indole-containing MW medium led to the inhibition of indole degradation by B. unamae CK43B (Fig. 1). In the gallic acid-supplemented cultures, up to 20% of the added indole remained unmetabolized compared to the control (0.32 mM relative to the initial 1.7 mM), indicating that gallic acid inhibits the degradation of indole by B. unamae CK43B.
These observations clearly showed that gallic acid itself is not a chemical signal for the induction of biofilm formation by B. unamae CK43B, but indole is the chemical signal. Unlike gallic acid, some other polyphenols, pyrogallol, (+)-catechin, and (−)-epigallocatechin, all of which showed no inhibitory effect on indole degradation by B. unamae CK43B, did not induce biofilm formation of B. unamae CK43B static-cultured in the polyphenol supplemented LB medium (Fig. 3).
Fig 3
Fig 3 Effect of phenolic additives on EPS production by B. unamae CK43B in MW liquid medium supplemented with 1.7 mM indole. (A) Illustrated TLC profiles of indole metabolites produced by B. unamae CK43B. The TLC plate was developed in CHCl3-MeOH-HCOOH (40:10:1). (B) The total biofilm (A570) was normalized to the population of planktonic cells, as reflected by the culture turbidity (OD660). The data represent the means ± the SD (n = 3). The letters a to d denote significant differences (P < 0.001). Columns (left to right): control (no polyphenolic additive), plus 1 mM gallic acid, plus 1 mM pyrogallol, plus 0.5 mM (−)-EGC (epigallocatechin), 1 mM (+)-catechin.
Culturing B. unamae CK43B in MW medium containing both indole and gallic acid resulted in a 12-fold (P < 0.001) increase in biofilm formation compared to control cultures without gallic acid. However, a Live/Dead BacLight kit to assess viability of the biofilm cells indicated that nearly half of the bacterial cells cultured in the indole-supplemented MW medium were viable (Fig. 4). In bacterial biofilms, dead cells often play a role as matrices for the viable cells (26), and the biofilm of B. unamae CK43B that formed in the medium supplemented with indole and gallic acid showed similar cell states (see Fig. S3, bottom panels, in the supplemental material).
Fig 4
Fig 4 Biofilm formation of B. unamae CK43B at the primary stage in MW medium. Biofilm-formed cells of B. unamae CK43B monocultured in MW medium containing 1.7 mM indole plus 1.0 mM gallic acid in a phase-contrast microscopic mode (visual light), and its cell viability assay using a Live/Dead BacLight bacterial viability kit under various fluorescence microscopic modes (Biorevo). Observation took place at ×600 magnification (a ×40 objective lens with a ×15 field lens).

Intergenus communication between B. unamae CK43B and E. coli via indole and exogenous gallic acid.

Several reports showed indole and isatin as quorum-sensing regulators in gammaproteobacteria (13, 27); likewise, our preliminary experiments showed a growth response curve (OD660) of planktonic B. unamae CK43B cells toward concentrations of supplemental indole in MW medium (see Fig. S4 in the supplemental material). Therefore, indole produced by E. coli K-12 is suggested to act as a chemical signal toward indole-degrading B. unamae CK43B. Because E. coli K-12 actively produces indole in LB medium, it is speculated that harmonized cell-to-cell communication between indole-degrading B. unamae CK43B and indole-producing E. coli K-12 in the coculture system is mediated via indole and its metabolites. In the coculture system, we also found a high correlation between the extracellular concentration of indole and flat-type biofilm formation (Fig. 5) normalized by bacterial cell growth (see Fig. S5 in the supplemental material). A high level of unmetabolized indole, uniquely found in gallic acid-containing culture medium, led to the speculation that the unmetabolized indole induces EPS production and morphological differentiation of the cocultured bacterial cells.
Fig 5
Fig 5 Biofilm formation of cocultured B. unamae CK43B with E. coli K-12 IFO 3301 at the primary stage in LB medium. Colonies formed on the bottom of a 3.5-cm-diameter plastic petri dish stained with crystal violet were directly observed in phase-contrast microscopic mode with ×4, ×20, and ×40 objective lenses, respectively. Cocultured cells in plain LB medium without any additives showed repression of biofilm formation on the flat bottom (top panels), whereas those in LB medium containing 1.0 mM gallic acid formed typical flat biofilm (bottom panels). Cells were stained with crystal violet, washed with water, air dried, and then subjected to microscopic observation under visible light. Scale bars located between two panels (top and bottom) are shared by a pair the panels for respective magnification. In the gallic acid-supplemented medium, the colonies attached on the bottom showed a typical flat-biofilm structure.
In the presence of excessive indole (1.7 mM) without gallic acid supplementation, monocultured B. unamae CK43B had drastically decreased culture turbidity compared to that in plain LB, probably due to the production of small amounts of 3-hydroxyindoxyl. Supplementation with 1.0 mM gallic acid was effective in allowing a small recovery of B. unamae CK43B cell growth, but the amount of indole added was probably excessive. An indole metabolite, 3-hydroxyindoxyl, repressed cell growth of B. unamae CK43B at 50 μM in plain LB medium (see Fig. S6 in the supplemental material). In contrast, monocultures of E. coli K-12 were rarely affected by the exogenous indole and gallic acid in terms of cell growth and biofilm formation.
Unlike those monocultures, supplementation with gallic acid accelerated flat biofilm formation in the coculture system with a statistic significance (P < 0.001) (see Fig. S5C in the supplemental material). Both planktonic and biofilm cells thus obtained were assayed for cell viability and assessed by FISH experiments to confirm the identity of the predominant bacterium in the coculture system (see Fig. S7 in the supplemental material). The cell viability of both biofilm and planktonic cells was nearly 80% within 24 h of biofilm formation, irrespective of the presence of 1.0 mM gallic acid (mean viable cell ratios ± standard deviations of 78% ± 1% in the planktonic state and 83% ± 1% in the biofilm state [n = 5]). In a FISH experiment to visualize the predominance of E. coli and B. unamae CK43B, both planktonic and biofilm cells that were hybridized in situ indicated that E. coli is the predominant bacterium in the cell aggregates, rather than B. unamae CK43B, but both bacteria survived and grew in the coculture system (Fig. 6). Thus, a double-labeling FISH experiment clarified that both B. unamae CK43B and E. coli K-12 contributed to biofilm formation in LB medium with appropriate concentrations of indole produced by E. coli and supplemental gallic acid (Table 1).
Table 1
Table 1 Predominance ratio of B. unamae CK43B in a coculture system with E. coli K-12a
Culture condition and cell stateMean ratio of B. unamae CK43B (%) ± SDb
In plain LB medium, planktonic71 ± 1
In plain LB medium, biofilm56 ± 3
In plain LB medium + 1.0 mM gallic acid, planktonic49 ± 6
In plain LB medium + 1.0 mM gallic acid, biofilm33 ± 5
a
The occupation ratio was calculated from merged FISH images of cocultured cells distributed in 145-by-110-μm2 slide glass plates. Cell numbers counted in a view were always >100.
b
n = 5.
Fig 6
Fig 6 FISH of cocultured E. coli K-12 IFO 3301 with B. unamae CK43B, along with EPS production in LB medium. In the FISH experiment, cultured bacterial cells were multiply subjected to in situ hybridization. The cells were observed under visual light using the oil immersion mode (×100 objective lens). (A to C) Cocultured bacterial cells maintained in a planktonic state with no biofilm formation. Images obtained with probe EUB338 for all bacteria (green fluorescence [B]) and a merged image of stained B. umamae CK43B-specific FISH probe BKH70 (false color of blue) and E. coli-specific FISH probe ENT186 (red fluorescence) (C) are also shown. (D to F) Biofilm-formed bacterial cells by coculture of B. unamae CK43B and E. coli K-12 IFO 3301 LB medium with 1.0 mM gallic acid supplementation. (G to I) Cells in the biofilm of cocultured B. unamae CK43B with E. coli K-12 IFO 3301 were vortexed to physically separate them and then subjected to FISH. Before vortexing, the biofilm was washed several times with 1× PBS to remove planktonic cells.
Little is known about intergenus communication among eubacteria, but some examples of communication between subclasses of Proteobacteria have been documented (28) involving signaling via indole and its derivatives, diketopiperazines (DKPs), and 2-alkyl-4(1H)-quinolines (AHQs) (16, 29, 30). Similarly to the intergenus communication mediated by AHL (31), a non-AHL signaling compound, AHQ, also functions as an interspecies communication signal between two subclasses of Proteobacteria (Pseudomonas aeruginosa of the subclass Gammaproteobacteria) and Burkholderia pseudomallei (Betaproteobacteria) (29). In addition, DKPs mediate cell-to-cell communication between two classes of eubacteria (Bacillus spp. of the phylum Firmicutes) and Pseudomonas spp. or Burkholderia spp. (class Proteobacteria) (29, 30).
Indole and some indole metabolites regulate EPS production in E. coli, which is often associated with induction of biofilm formation (see Table S1 in the supplemental material) (12, 16, 27, 3238). Under exposure to certain concentrations of gallic acid that do not adversely affect cell growth, coculturing B. unamae CK43B with indole-producing E. coli K-12 resulted in biofilm formation, partly due to inhibition of indole metabolism (see Fig. S7 in the supplemental material). However, the dynamics of the indole pool, including inhibition of its biosynthesis, in the coculture system with gallic acid are uncertain.
Generally, polyphenols inhibit biofilm formation of eubacteria (3942). For instance, Lee et al. reported that the simple dihydrochalcone phloretin (2′,4,4′,6′-tetrahydroxydihydrochalcone) inhibits biofilm formation of the pathogenic E. coli strain O157:H7. Phloretin, however, does not promote the growth of planktonic E. coli O157:H7 or induce its functional differentiation to produce enterotoxin (42). Conversely, monocultured B. unamae CK43B formed a viable biofilm in MW medium in our experiments only when indole and gallic acid were both present. Thus, unlike phloretin and some other polyphenols, gallic acid displayed the opposite action in biofilm formation of indole-producing bacteria. Our findings are significant because indole is a relatively common metabolite produced from l-Trp in a single-step reaction catalyzed by tryptophanase in some important enterobacteria (43). Whether indole and gallic acid together promote biofilm formation of Burkholderia spp. pathogenic against humans or plants remains to be determined.

ACKNOWLEDGMENTS

We thank Eri Fukushi (GC-MS and NMR Laboratory, Research Faculty of Agriculture, Hokkaido University) for assistance with the MS and NMR analyses.
We are also grateful to the Ministry of Education, Culture, Sports, Science, and Technology of Japan for scholarship assistance to D.K. This research was supported by a Grant-in-Aid for Scientific Research A (no. 20248033 to Y.H.) from the Japan Society for the Promotion of Science.

Supplemental Material

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REFERENCES

1.
Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, and Lappin-Scott HM. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49:711–745.
2.
Donlan RM. 2002. Biofilms: microbial life on surfaces. Emerg. Infect. Dis. 8:881–890.
3.
Vu B, Chen M, Crawford RJ, and Ivanova EP. 2009. Bacterial extracellular polysaccharides involved in biofilm formation. Molecules 14:2535–2554.
4.
Costerton JW, Stewart PS, and Greenberg EP. 1999. Bacterial biofilms: a common cause of persistent infections. Science 284:1318–1322.
5.
Singh R, Paul D, and Jain RK. 2006. Biofilms: implication in bioremediation. Trends Microbiol. 14:389–397.
6.
Ahimou F, Semmens MJ, Haugstad G, and Novak PJ. 2007. Effect of protein, polysaccharide, and oxygen concentration profiles on biofilm cohesiveness. Appl. Environ. Microbiol. 73:2905–2910.
7.
Mayer C, Moritz R, Kirschner C, Borchard W, Maibaum R, Wingender J, and Flemming HC. 1999. The role of intermolecular interactions: studies on model systems for bacterial biofilms. Int. J. Biol. Macromol. 26:3–16.
8.
Davies DG, Parsek MR, Pearson JP, Iglewski BH, Costerton JW, and Greenberg EP. 1998. The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280:295–298.
9.
González Barrios AF, Zuo RY, Hashimoto Y, Yang L, Bentley WE, and Wood TK. 2006. Autoinducer 2 controls biofilm formation in Escherichia coli through a novel motility quorum-sensing regulator (MqsR, B3022). J. Bacteriol. 188:587–598.
10.
Waters CM and Bassler BL. 2005. Quorum sensing: cell-to-cell communication in bacteria. Annu. Rev. Cell Dev. Biol. 21:319–346.
11.
Rivas M, Seeger M, Holmes DS, and Jedlicki E. 2005. A Lux-like quorum sensing system in the extreme acidophile Acidithiobacillus ferrooxidans. Biol. Res. 38:283–297.
12.
Wang D, Ding X, and Rather PN. 2001. Indole can act as an extracellular signal in Escherichia coli. J. Bacteriol. 183:4210–4216.
13.
Snell EE. 1975. Tryptophanase: structure, catalytic activities, and mechanism of action. Adv. Enzymol. Relat. Areas Mol. Biol. 42:287–333.
14.
Vert G and Chory J. 2011. Crosstalk in cellular signaling: background noise or the real thing? Dev. Cell 21:985–991.
15.
March JC and Bentley WE. 2004. Quorum sensing and bacterial cross-talk in biotechnology. Curr. Opin. Biotechnol. 15:495–502.
16.
Lee J, Jayaraman A, and Wood TK. 2007. Indole is an inter-species biofilm signal mediated by SdiA. BMC Microbiol. 7:42.
17.
Sitepu IR, Hashidoko Y, Santoso E, and Tahara S. 2009. Growth-promoting properties of bacteria isolated from dipterocarp plants of acidic lowland tropical peat forest in Central Kalimantan, Indonesia. J. For. Res. Indonesia 6:96–118.
18.
Rahman A, Sitepu IR, Tang SY, and Hashidoko Y. 2010. Salkowski's reagent test as a primary screening index for functionalities of rhizobacteria isolated from wild dipterocarp saplings growing naturally on medium-strongly acidic tropical peat soil. Biosci. Biotechnol. Biochem. 74:2202–2208.
19.
Kim D, Rahman A, Sitepu IR, and Hashidoko Y. Accelerated degradation of exogenous indole by Burkholderia unamae strain CK43B exposed to some pyrogallol-type polyphenols. Biosci. Biotechnol. Biochem., in press.
20.
Hashidoko Y, Tada M, Osaki M, and Tahara S. 2002. Soft gel medium solidified with gellan gum for preliminary screening for root-associating, free-living nitrogen-fixing bacteria inhibiting the rhizoplane of plants. Biosci. Biotechnol. Biochem. 66:2259–2263.
21.
Domka J, Lee J, Bansal T, and Wood TK. 2007. Temporal gene-expression in Escherichia coli K-12 biofilms. Environ. Microbiol. 9:332–346.
22.
Merritt JH, Kadouri DE, and O'Toole GA. 2011. Growing and analyzing static biofilms. Curr. Protoc. Microbiol. 22:1B.1.1–1B.1.18.
23.
Hori T, Haruta S, Ueno Y, Ishii M, and Igarashi Y. 2006. Dynamic transition of a methanogenic population in response to the concentration of volatile fatty acids in a thermophilic anaerobic digester. Appl. Environ. Microbiol. 72:1623–1630.
24.
Klausen M, Heydorn A, Ragas P, Lambertsen L, Aaes-Jørgensen A, Molin S, and Tolker-Nielsen T. 2003. Biofilm formation by Pseudomonas aeruginosa wild type, flagella, and type IV pilus mutants. Mol. Microbiol. 48:1511–1524.
25.
Kirisits MJ and Parsek MR. 2006. Does Pseudomonas aeruginosa use intercellular signalling to build biofilm communities? Cell. Microbiol. 8:1841–1849.
26.
Webb JS, Thompson LS, James S, Charlton T, Tolker-Nielsen T, Koch B, Givskov M, and Kjelleberg S. 2003. Cell death in Pseudomonas aeruginosa biofilm development. J. Bacteriol. 185:4585–4592.
27.
Lee J, Bansal T, Jayaraman A, Bentley WE, and Wood TK. 2007. Enterohemorrhagic Escherichia coli biofilms are inhibited by 7-hydroxyindole and stimulated by isatin by isatin. Appl. Environ. Microbiol. 73:4100–4109.
28.
Ryan RP and Dow JM. 2008. Diffusible signals and interspecies communication in bacteria. Microbiology 154:1845–1858.
29.
Holden MTG, Chhabra SR, de Nys R, Stead P, Bainton NJ, Hill PJ, Manefield M, Kumar N, Labatte M, England D, Rice S, Givskov M, Salmond GPC, Stewart GSAB, Bycroft BW, Kjelleberg S, and Williams P. 1999. Quorum-sensing cross talk: isolation and chemical characterisation of cyclic dipeptides from Pseudomonas aeruginosa and other Gram-negative bacteria. Mol. Microbiol. 33:1254–1266.
30.
Diggle SP, Lumjiaktase P, Dipilato F, Winzer K, Kunakorn M, Barrett DA, Chhabra SR, Cámara M, and Williams P. 2006. Functional genetic analysis reveals a 2-alkyl-4-quinolone signaling system in the human pathogen Burkholderia pseudomallei and related bacteria. Chem. Biol. 13:701–710.
31.
Riedel K, Hentzer M, Geisenberger O, Huber B, Steidle A, Wu H, Høiby N, Giveskov M, Molin S, and Eberl L. 2001. N-Acyl-l-homoserinelactone-mediated communication between Pseudomonas aeruginosa and Burkholderia cepacia in mixed biofilms. Microbiology 147:3249–3262.
32.
Worthington RJ, Richards JJ, and Melander C. 2012. Small molecule control of bacterial biofilms. Org. Biomol. Chem. 10:7457–7474.
33.
Bansal T, Englert D, Lee J, Hegde M, Wood TK, and Jayaraman A. 2007. Differential effects of epinephrine, norepinephrine, and indole on Escherichia coli O157:H7 chemotaxis, colonization, and gene expression. Infect. Immun. 75:4597–4607.
34.
Lee J, Zhang XS, Hegde M, Bentley WE, Jayaraman A, and Wood TK. 2008. Indole cell signaling occurs primarily at low temperatures in Escherichia coli. ISME J. 2:1007–1023.
35.
Domka J, Lee J, and Wood TK. 2006. YliH (BssR) and YceP (BssS) regulate Escherichia coli K-12 biofilm formation by influencing cell signaling. Appl. Environ. Microbiol. 72:2449–2459.
36.
Baca-DeLancey RR, South MM, Ding X, and Rather PN. 1999. Escherichia coli genes regulated by cell-to-cell signaling. Proc. Natl. Acad. Sci. U. S. A. 96:4610–4614.
37.
Garbe TR, Kobayashi M, and Yukawa H. 2000. Indole-inducible proteins in bacteria suggest membrane and oxidant toxicity. Arch. Microbiol. 173:78–82.
38.
Martino PD, Fursy R, Bret L, Sundararaju B, and Phillips RS. 2003. Indole can act as an extracellular signal to regulate biofilm formation of Escherichia coli and other indole-producing bacteria. Can. J. Microbiol. 49:443–449.
39.
Isaacs H Jr, Chao D, Yanofsky C, and Saier MH Jr 1994. Mechanism of catabolite repression of tryptophanase synthesis in Escherichia coli. Microbiology 140:2125–2134.
40.
Duarte S, Gregoire S, Singh AP, Vorsa N, Schaich K, Bowen WH, and Koo H. 2006. Inhibitory effects of cranberry polyphenols on formation and acidogenicity of streptococcus mutans biofilms. FEMS Microbiol. Lett. 257:50–56.
41.
Evensen NA and Braun PC. 2009. The effects of tea polyphenols on Candida albicans: inhibition of biofilm formation and proteasome inactivation. Can. J. Microbiol. 55:1033–1039.
42.
Cho YS, Oh J, and Oh KH. 2010. Antimicrobial activity and biofilm formation inhibition of green tea polyphenols on human teeth. Biotechnol. Bioproc. Eng. 15:359–364.
43.
Lee JH, Regmi SC, Kim JA, Cho MH, Yun H, Lee CS, and Lee J. 2011. Apple flavonoid phloretin inhibits Escherichia coli O157:H7 biofilm formation and ameliorates colon inflammation in rats. Infect. Immun. 79:4819–4827.

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 79Number 1615 August 2013
Pages: 4845 - 4852
PubMed: 23747701

History

Received: 15 April 2013
Accepted: 2 June 2013
Published online: 24 July 2013

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Authors

Dongyeop Kim
Division of Applied Bioscience, Research Faculty of Agriculture, Hokkaido University, Sapporo, Japan
Irnayuli R. Sitepu
Forest Microbiology Laboratory, Forest and Nature Conservation Research and Development Center, Forest Research and Development Agency, Bogor, Indonesia
Yasuyuki Hashidoko
Division of Applied Bioscience, Research Faculty of Agriculture, Hokkaido University, Sapporo, Japan

Notes

Address correspondence to Yasuyuki Hashidoko, [email protected].

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