Establishment of New Genetic Traits in a Microbial Biofilm Community

Abstract

Conjugational transfer of the TOL plasmid (pWWO) was analyzed in a flow chamber biofilm community engaged in benzyl alcohol degradation. The community consisted of three species, Pseudomonas putida RI, Acinetobacter sp. strain C6, and an unidentified isolate, D8. Only P. putida RI could act as a recipient for the TOL plasmid. Cells carrying a chromosomally integrated lacI^q gene and a lacp-gfp-tagged version of the TOL plasmid were introduced as donor strains in the biofilm community after its formation. The occurrence of plasmid-carrying cells was analyzed by viable-count-based enumeration of donors and transconjugants. Upon transfer of the plasmids to the recipient cells, expression of green fluorescence was activated as a result of zygotic induction of the gfp gene. This allowed a direct in situ identification of cells receiving the gfp-tagged version of the TOL plasmid. Our data suggest that the frequency of horizontal plasmid transfer was low, and growth (vertical transfer) of the recipient strain was the major cause of plasmid establishment in the biofilm community. Employment of scanning confocal laser microscopy on fixed biofilms, combined with simultaneous identification of P. putida cells and transconjugants by 16S rRNA hybridization and expression of green fluorescence, showed that transconjugants were always associated with noninfected P. putida RI recipient microcolonies. Pure colonies of transconjugants were never observed, indicating that proliferation of transconjugant cells preferentially took place on preexisting P. putida RI microcolonies in the biofilm.

Biological wastewater treatment, removal of organic compounds from contaminated soil, biogas reactors, etc., all involve processes based on the action of microbial communities. Increasing demands for improving the efficiencies of degradation in these systems have resulted in the need for a better understanding of the function and significance of the individual organisms in the community. One way to stimulate degradation of certain pollutants would be to introduce new genetic information into the microbial community. This process, termed bioaugmentation, involves the addition of new bacteria to the community, thus introducing new biodegradation pathways for metabolic conversion of the pollutants. However, enhancement of biodegradation often fails due to poor establishment and/or survival of the new strain in the environment (20, 25, 44). An alternative approach is to transfer the relevant genes on conjugative plasmids to indigenous organisms, from which the genes may spread further in the community (13, 15, 17, 18, 28, 32).

If plasmid genes (e.g., for resistance to heavy metals or antibiotics, for metabolic pathways, etc.) are acquired from foreign organisms and exchanged between members of the community, the capacity of a community to develop new microbial traits may be increased, making the community more responsive to environmental changes (19, 24, 40). However, little is known about the features which are important for the integration of new genetic traits by microbial communities.

Methods for determination of the organism composition and for tracing specific genes in microbial communities include the use of nucleic acid probes targeting specific DNA regions (for a review, see Sayler and Layton [34]), frequently by including a PCR step to amplify the region of interest (31, 39); in situ rRNA hybridization employing 16S or 23S rRNA probes targeting specific organisms (for a review, see Amann et al. [2]); or simple plating on selective plates.

During the last 5 to 10 years, several new techniques have been developed for more detailed analysis of complex microbial communities. The application of scanning confocal laser microscopy (SCLM) in combination with a number of fluorescent biomarkers has been used for analysis of structural features, such as the arrangement of cells, polymers, and channels in microbial environments (27, 46, 47). In situ hybridization with fluorescence-labeled 16S or 23S rRNA probes in combination with SCLM is a powerful approach for visualization of the spatial distribution of important group organisms in bacterial communities (29, 41).

Fluorescent markers may also be introduced by genetic engineering. The gfp gene encoding the green-fluorescent protein (Gfp) (7) and enhanced by Cormack et al. (10) has been very useful as a reporter for studies of gene expression in single cells. For example, fusions between the relevant promoter and the gfp gene have been used to monitor subcellular protein localization during sporulation in Bacillus subtilis (42) and to analyze mycobacterial interactions with macrophages (14). Gfp has also been used as a reporter to monitor the kinetics of TOL plasmid transfer between bacteria growing on agar surfaces (8) as well as to track individual cells in an activated-sludge community (16).

In the present work, we have investigated the establishment of the TOL plasmid in a biofilm community growing on benzyl alcohol as the sole carbon and energy source. A consortium of bacteria, originally isolated from a creosote-polluted aquifer, was previously used in a waste gas biofilter for the degradation of toluene (29). From this complex consortium of bacteria, a model community consisting of three species (Pseudomonas putida RI; a strain of Acinetobacter sp.; and a strain, D8, tentatively associated with the β subgroup of Proteobacteria strains), isolated as individual clones from the original waste gas biofilter, was defined for our biofilm investigations. In situ rRNA hybridization was used to identify the three species present in the community, and an approach based on zygotic induction of the Gfp protein developed to study plasmid transfer directly in the community (8) was employed in order to localize conjugational activities.

MATERIALS AND METHODS

Strains, plasmids, and growth conditions.

The bacterial strains and plasmids and their relevant characteristics are listed in Table 1. The strains were grown in Luria-Bertani (LB) broth (containing 10 g of tryptone, 5 g of yeast extract, and 4 g of NaCl). When required, antibiotics were added at final concentrations of 50 μg/ml for rifampin and nalidixic acid and 10 μg/ml for kanamycin.

TABLE 1.

Strains and plasmids

Strain or plasmid	Relevant characteristics	Source or reference
E. coli K-12
CC118λpir	Δ(ara-leu) araD ΔlacX74 galE galK phoA thi-1 rpsE rpoB argE(Am) recA λpir lysogen	21
HB101	E. coli K-12/B hybrid; SmrrecA thi pro leu hsdRM+	4
P. putida
RI (SMO116)	Natural isolate; γ subgroup of class Proteobacteriaa	29
JB154	SMO116 Rifr	This work
JB156	SMO116 Nalr	This work
KT2440	hsdR/TOL	K. Timmis
KT2442	hsdR Rifr	K. Timmis
SM1443	KT2442 lacIq; npt removed by site-specific recombination	This work
BBC443	SM1443/TOLgfpmut3b	This work
BBC458	JB156/TOLgfpmut3b	This work
BBC516	JB154/TOLgfpmut3b	This work
Acinetobacter sp. strain C6 (SMO112)	Natural isolate; γ subclass of class Proteobacteriab	29
Unknown genus D8 (SMO125)	Natural isolate; β subclass of class Proteobacteriac	29
Plasmids
RK600	Cmr ColE1 oriV RP4 oriT; helper plasmid in triparental matings	22
pUC18Not	Same as pUC18 (49) but with polylinker flanked with NotI sites	21
pUTKm	Apr Kmr; transposon delivery vector for mini-Tn5 Km	11
pCK242	mini-Tn5 transposon delivery vector; npt gene flanked by res sites	23
pJBA27	SphI-HindIII gfpmut3b fragment cloned downstream of PA1/O4/O3	3
pJBA28	NotI fragment PA1/O4/O3::gfpmut3b from pJBA27 cloned into pUT mini-Tn5 Km	3
pSM1431	lacIq fragment (38) cloned into pUC18Not (Sca-SalI)	This work
pSM1435	NotI fragment (lacIq) of pSM1431 inserted into pUTKm	This work
TOL	Self-transmissible approximately 117-kb plasmid; from P. putida mt-2	43
TOLgfpmut3b	Integration of PA1/O4/O3::gfpmut3b from pJBA28 into TOL	This work

^aFrom reference 29. 

^bBased on 16S rRNA sequence. 

^cBased on hybridization with group-specific probes. 

Mutants of the natural isolate P. putida RI, resistant to either nalidixic acid or rifampin, were isolated as spontaneous mutants on LB broth plates containing concentrations of 100 μg of nalidixic acid or rifampin per ml, respectively.

A modified version of the pUT vector (12), comprising a mini-Tn5 transposon with RP4 resolvase sites flanking the npt gene responsible for kanamycin resistance (pCK242) (23), was used for insertion of the lacI^q gene (38) into the chromosome of KT2442. Triparental mating among a donor strain (carrying the lacI^q delivery plasmid pSM1435), the helper strain HB101(RK600), and the recipient P. putida KT2442 resulted in KT2442 derivatives conferring kanamycin resistance with the lacI^q gene inserted in the chromosome. Subsequently, the npt gene was deleted as described by Christensen et al. (8). One such kanamycin-sensitive clone was picked and designated SM1443.

The gene encoding the green fluorescent protein Gfpmut3b was obtained from Cormack et al. (10). The gfpmut3b gene was PCR amplified as a 0.7-kb SphI-HindIII fragment. When introducing a SphI site in the start codon of gfpmut3b the sequence was changed in the PCR such that the Gfpmut3b protein contained an arginine instead of a serine residue at position 2.

The gfpmut3b fragment was cloned downstream of the promoter P_A1/O4/O3 (6), at an optimal distance from a synthetic ribosome binding site (RBSII, from plasmid pQE70; Qiagen GmbH, Germany), and upstream of a region with two strong transcriptional terminators, T₀ (from phage lambda) and T₁ (from the rrnB operon of Escherichia coli), and with translational stop codons in all three reading frames. The NotI fragment from the resulting plasmid (pJBA27), containing RBSII, gfpmut3b, the translational stop codons, and the transcriptional terminators, was inserted into the NotI site of pUTKm, resulting in the transposon delivery vector pJBA28, containing a cassette with P_A1/O4/O3::gfpmut3b and the npt gene.

Insertion of the P_A1/O4/O3::gfpmut3b cassette into the TOL plasmid was performed in two steps. First, a triparental mating was performed, in which the helper plasmid RK600 was used to mobilize the delivery plasmid pJBA28 from the donor strain, CC118λpir, into the recipient strain, P. putida KT2440. Selection on AB-minimal plates (9) containing 10 mM citrate and 50 μg of kanamycin/ml resulted in KT2440 derivatives carrying the P_A1/O4/O3::gfpmut3b cassette inserted either in the chromosome or in the TOL plasmid. Clones with the cassette integrated in the TOL plasmid were selected from a second round of conjugation. Colonies from the first-round selective plates (>1,000 on one plate) were suspended in 1 ml of 0.9% NaCl. The suspended cells were then mated with the kanamycin-sensitive recipient, SM1443. Selection on plates containing 50 μg of kanamycin/ml and 50 μg of rifampin/ml resulted in SM1443 Km^r derivatives carrying the TOL plasmid with the P_A1/O4/O3::gfpmut3b cassette inserted at random positions. One clone (designated BBC443) was chosen for subsequent experiments based on the following criteria: it was able to grow on AB-minimal plates supplemented with either 5 mM benzyl alcohol or 5 mM sodium benzoate as the sole carbon and energy source; the colonies were green fluorescent upon addition of 1 mM IPTG (isopropyl-β-d-thiogalactopyranoside); and finally, the conjugation frequency on plates of the gfp-tagged plasmid was similar to that of the wild-type TOL plasmid.

Cultivation of biofilms.

The biofilm model community comprised the following organisms: P. putida RI JB156, which is resistant to nalidixic acid, or JB154, which is resistant to rifampin; Acinetobacter sp. strain C6; and an unidentified strain, isolate D8, of the β subgroup of the class Proteobacteria (Table 1). All of the strains are able to mineralize toluene and benzyl alcohol (30).

Biofilms were cultivated as mixtures of the three species in rectangular two- or four-channel flow cells (47) with individual channel dimensions of 1 by 4 by 40 mm supplied with a flow of FAB substrate [1 mM MgCl₂, 0.1 mM CaCl₂, 0.01 mM Fe-EDTA [catalog no. E6760; Sigma, St. Louis, Mo.), 0.15 mM (NH₄)SO₄, 0.33 mM Na₂HPO₄, 0.2 mM KH₂PO₄, 0.5 mM NaCl]. Benzyl alcohol (Merck KGaA, Darmstadt, Germany) at a concentration of 0.5 mM was used as the sole carbon source.

Flow chamber experiments.

The flow system was assembled with autoclaved silicone tubing. Complete sterilization was performed by pumping a solution of 0.5% sodium hypochlorite into the system and leaving it overnight. The following day approximately 0.2 liters of sterile water per flow chamber was flushed through the system before the medium was pumped in.

During biofilm growth, the medium was pumped through the flow cells at a rate of 0.2 mm/s with a peristaltic pump (model 205S; Watson-Marlow Inc., Wilmington, Mass.). The flow cells were inoculated with mixtures of cultures of the three strains pregrown for 2 days in LB medium in the ratio 1:10:5 for P. putida, Acinetobacter, and isolate D8, respectively. The mixtures were sonicated with a Branson sonifier (Branson Ultrasonics Corp., Danbury, Conn.) for 1 min at output control 3 and duty cycle 40%, and 0.25 ml of the mixed culture was injected into each channel.

Sequencing of 16S rRNA.

Sequencing of 16S rRNA was performed with an automatic 373A DNA sequencer (Applied Biosystems, Foster City, Calif.) directly on PCR products generated from chromosomal DNA extracts according to the manufacturer’s recommendations. The following primers were used (in the 5′→3′ direction): 11F, GTTTGATC(A/C)TGGCTCAGATTG; 344R, CCCCACTGCTGCCTCCCGT; 515R, GTATTACCGCGGC(G/T)GCTGGCAC; 922R, GCTTGTGCGGGCCCCCGTC; 1101R, GACAAGGGTTGCGCTCGTT; 1389R, GTGACGGGCGGTGTGTACAAG; and 1465R, CCCCAGTCATGAATCATAAAGTGGT. Initial phylogenetic analysis of the sequenced rRNAs was obtained from on-line services of the Ribosomal Database Project (SIMILARITY_RANK [26]). In addition, potential signature sequences were identified as described by Woese (45), allowing differentiation between the major groups of the class Proteobacteria.

Oligonucleotide probes.

For 16S rRNA hybridization, the probes EUB338 (specific for the domain Bacteria [1]) and PP986 (specific for the P. putida subgroup A [29]) were used. Based on the sequence information obtained, specific rRNA probes for Acinetobacter sp. strain C6 and isolate D8 were designed. The specificities of Acn449 and D8_647 were tested against published sequences with the CHECK_PROBE program from the Ribosomal Database Project (26). All probes were tested against the organisms of the community and found to be specific for their respective target organisms (data not shown). Oligonucleotide probes labeled with fluorescein isothiocyanate or the indocarbocyanine fluorescent dyes CY3 and CY5 were purchased from Hobolth DNA Syntese (Hillerød, Denmark).

Embedding of hydrated biofilm samples.

Embedding was performed as a nondestructive method to maintain a fixed biofilm in its 3-dimensional native hydrated state and at the same time to allow easy handling of the fixed biofilm. Throughout the embedding procedure, pumping of solutions through the flow cells was performed at 0.8 mm/s. Biofilms were fixed in freshly made 3% paraformaldehyde solution (33) by pumping the solution through the flow channels with attached biofilms. To ensure the complete fixation of all cells in the biofilms, the solution was retained in the channels for 1 h at room temperature before the biofilms were washed by flowthrough of phosphate-buffered saline for 5 min. Embedding was performed by introducing a solution of 1 ml of 20% acrylamide (200:1 acrylamide-bisacrylamide) mixed with 8 μl of N,N,N′,N′-tetramethylethylenediamine (Kodak International Biotechnologies Inc., New Haven, Conn.), and immediately prior to inoculation, 20 μl of ammonium persulfate (Kodak International Biotechnologies Inc.) was also added. Approximately 0.5 ml of the solution was pumped into the channel, where it solidified within 2 min after addition of the ammonium persulfate.

Hybridization of embedded biofilm cells.

After fixation and embedding, the polyacrylamide block with biofilm was placed on a 6-well hybridization slide (Novakemi AB, Enskede, Sweden) and equilibrated for 15 min with hybridization buffer containing 30% formamide at 37°C. Then 30 μl of hybridization mixture (30% formamide, 0.9 M NaCl, 100 mM Tris [pH 7.2], 0.1% sodium dodecyl sulfate) containing 75 ng of probe was added to each hybridization well. Cells were incubated with hybridization solution for 3 h at 37°C in a moisture chamber. For washing, 50 μl of each washing solution was added to each well as follows. First, the acrylamide blocks were washed in solution I (30% formamide, 0.9 M NaCl, 100 mM Tris [pH 7.2], 0.1% sodium dodecyl sulfate) for 40 min at 37°C, then they were washed for 40 min in washing solution II (0.1 M Tris [pH 7.2], 0.9 M NaCl) at 37°C, and finally, they were rinsed two times in 50 μl of distilled water. The acrylamide blocks were mounted in 2× SlowFade phosphate-buffered saline-based antifade solution (Molecular Probes, Eugene, Oreg.).

Microscopy and image analysis.

All microscopic observation and image acquisition was performed on a TCS4D confocal microscope (Leica Lasertechnik GmbH, Heidelberg, Germany) equipped with three detectors and filter sets for simultaneous monitoring of fluorescein isothiocyanate/green-fluorescent protein and the indocarbocyanine dyes CY3 and CY5.

Multichannel simulated fluorescence projection (SFP; a shadow projection) images and vertical cross sections through the biofilms were generated with the IMARIS (Bitplane AG, Zürich, Switzerland) software package running on an Indigo 2 (Silicon Graphics, Mountain View, Calif.) workstation. The images were further processed for display with Photoshop software (Adobe, Mountain View, Calif.).

The spatial distribution of organisms was estimated by measuring the area covered by hybridization signal (represented by different colors) in series of two-channel optical sections by using simple thresholding. One channel represented the cells targeted with the general eubacterial probe EUB338 (1), and the other channel represented the cells hybridized with a species-specific probe. Before thresholding was done, the images were preprocessed with IMARIS: background was removed by a lowpass filtering and background subtraction, and a final correction for loss of intensity from deeper layers was performed with the emission attenuation algorithm of IMARIS. Thresholding was performed on the processed images with the HIPS image analysis package.

Nucleotide sequence accession number.

The sequence reported here will appear in the EMBL, GenBank, and DDBJ nucleotide sequence databases under accession no. Y11464 (Acinetobacter sp. strain C6) and Y11465 (isolate D8).

RESULTS AND DISCUSSION

The model bacterial community.

Flow chamber biofilms were established by mixing three strains which had been precultured separately for 2 days at 30°C in LB media prior to inoculation. Benzyl alcohol was added as the sole carbon and energy source for growth of the community. A quantitative analysis of the distribution of the different community members at different depths of a 7-day-old biofilm was done by probing with fluorescent 16S ribosomal DNA probes. Probes targeting the three selected species, P. putida, Acinetobacter sp. strain C6, and isolate D8, were used for identification. The analysis (Fig. 1A) revealed that P. putida RI was the most common species in the upper layers, whereas Acinetobacter dominated the layers near the substratum, where most of the overall biomass was observed (Fig. 1B). Isolate D8 was not a dominant community member at any depth of the biofilm.

FIG. 1.

Quantitative analysis showing the spatial distribution of the different organisms of the mixed-culture biofilm sampled at day 7. (A) Vertical profile through the biofilm showing percentage of Acinetobacter sp. strain C6 (▵), P. putida RI (◊), and isolate D8 (□) relative to the total number of cells targeted with the general eubacterial 16S rRNA probe (EUB338 [1]). Each profile is an average of three images (each consisting of 31 optical sections) captured at three random locations in the biofilm. (B) Relative area covered by cells in each section, taken as an average over six images. Standard deviations are indicated by error bars.

Introduction of genes encoding the TOL biodegradation pathway into the flow chamber community.

The three species comprising the microbial community are able to utilize benzyl alcohol as their sole carbon and energy source. Based on DNA hybridization and investigations of substrate profiles and metabolite formation, we have previously concluded that all the strains seem to harbor degradation pathways similar to that of the P. putida plasmid TOL (pWWO) (48). However, despite the similarities, the three pathways are probably not identical (30).

To analyze the establishment of a new metabolic pathway in the microbial community, the TOL plasmid was chosen. The TOL plasmid derivative TOLgfpmut3b (Table 1), which carries a fluorescent reporter tag and a kanamycin resistance marker gene, was used for this investigation. When grown in batch culture with benzyl alcohol as the sole carbon source the doubling time of P. putida RI was approximately 110 min (not shown). Introduction of the TOL plasmid into this strain by conjugation decreased the doubling time to approximately 80 min. The TOL plasmid had no significant effect on host cell generation times in media supplemented with carbon sources unrelated to the TOL pathway.

Establishment of the TOL plasmid degradation pathway in the biofilm community could occur through colonization of the biofilm by the plasmid-carrying donor strain, through plasmid transfer to the indigenous bacteria, or both. In the experiment, these possibilities could be monitored independently, and the experimental results are presented accordingly.

The population size of the incoming donor strain—a rifampin-resistant derivative of P. putida RI carrying the plasmid TOLgfpmut3b (BBC516)—was determined as rifampin- and kanamycin-resistant viable counts present in the effluent from the flow chamber. These determinations represent good estimates of the total biofilm populations, as indicated by control experiments in which entire biofilm communities have been analyzed and compared with effluents (not shown). Transfer of the TOL plasmid to the indigenous nalidixic acid-resistant strain of P. putida RI was estimated from cell counts on selective plates supplemented with nalidixic acid and kanamycin. Reversal of the antibiotic marker phenotypes in the incoming and indigenous strains of P. putida did not influence the outcome of the experiment (not shown).

The donor strain was introduced 2 days after the initial colonization of the model community. Three different concentrations of an overnight culture (5 · 10⁴, 5 · 10⁵, or 5 · 10⁶ CFU/ml) were injected into separate flow channels. Total cell counts in the effluents, as well as relative numbers of the indigenous P. putida RI cells, were marginally affected by the introduction of the P. putida RI cells carrying the TOL plasmid (Fig. 2A and B). However, as shown in Fig. 2C, the relative proportion of incoming P. putida RI cells started to increase exponentially immediately after their introduction. Independent of their abundance at the start of the experiment, they reached a steady-state level of approximately 10% of the total count 6 to 7 days after their introduction. Despite their apparently more effective degradation pathway (faster growth) and their rapid rate of establishment during the first few days, P. putida RI cells carrying the TOL plasmid did not outcompete the indigenous P. putida RI cells (Fig. 2C), even over a period of 40 days (data not shown).

FIG. 2.

Time course analysis of the distribution of donor cells, P. putida RI, and transconjugants relative to the total number of cells collected from flow channel effluents. The donor (P. putida RI/TOLgfpmut3b) was introduced at day 2 in three different densities: 5 · 10⁴ (▴), 5 · 10⁵ (▪), and 5 · 10⁶ (⧫) CFU/ml. Total counts (A) were enumerated on pure LB broth plates. The P. putida RI cells (Nal^r) (B), donor cells (Rif^r) (C), and transconjugants (Nal^r Km^r) (D) were enumerated on LB broth plates containing the appropriate antibiotics, and the numbers were taken relative to the total counts. Each data point is the average of two independent experiments.

Mating experiments performed on an agar surface showed that the TOLgfpmut3b plasmid could only be transferred to P. putida RI. When the donor cells (P. putida RI/TOLgfpmut3b) and the isogenic recipient cells (P. putida RI) were mixed in equal numbers on an agar surface, approximately 20% of the recipients hosted the plasmid after 24 h. Based on these results, it was surprising to see that plasmid transfer occurring in the biofilm community was almost negligible during the first several days and remained low throughout the experiment (Fig. 2D).

In conclusion, the data presented in Fig. 2 show (i) that establishment of the incoming donor strain in the biofilm was possible; (ii) that the TOL plasmid was established mainly by growth of the incoming donor cells (vertical transfer), possibly facilitated by the growth advantage mediated by the TOL degradation pathway; and (iii) that the TOL plasmid was transferred at a low but measurable rate to recipient cells in the community.

In a separate experiment, the influence of the strain background of the incoming organism was investigated. The strain P. putida KT2442 (Rif^r) carrying the TOLgfpmut3b plasmid is not isogenic with P. putida RI, although the 16S rRNA sequences of the two strains only differ by a few bases (unpublished data). In addition, strain KT2442 grows in batch culture with benzyl alcohol as the sole carbon source with a doubling time of approximately 80 min, which is significantly faster than that measured for cultures of P. putida RI. Furthermore, when P. putida KT2442 (Rif^r) carrying the TOLgfpmut3b plasmid was cocolonized with the other three species, the strain constituted more than 90% of the total cell counts only a few days after colonization (data not shown), indicating that the strain is a good colonizer and competitor when introduced together with the three community strains.

In the following experiment, presented in Fig. 3, the impact of the donor cell background on the efficiency of establishment in the biofilm was investigated. The donor strain (KT2442/TOLgfpmut3b) was introduced 2 days after the initial colonization of the microbial flow chamber community. The incoming donor was introduced into separate flow chambers in three different concentrations (5 · 10⁶, 5 · 10⁷, or 5 · 10⁸ CFU/ml) of an overnight culture. Time course analysis of the population profile of cells collected from the effluents showed that neither the total cell count nor the relative proportion of P. putida RI cells was significantly affected by the introduction of the new strain (Fig. 3A and B).

FIG. 3.

Time course analysis of the distribution of donor cells, P. putida RI, and transconjugants relative to the total number of cells collected from flow channel effluents. The donor (P. putida KT2442/TOLgfpmut3b) was introduced at day 2 in three different concentrations: 5 · 10⁶ (▴), 5 · 10⁷ (▪), and 5 · 10⁸ (⧫) CFU/ml. Total counts (A) were enumerated on pure LB broth plates. The P. putida RI cells (Nal^r) (B), donor cells (Rif^r) (C), and transconjugants (Nal^r Km^r) (D) were enumerated on LB broth plates containing the appropriate antibiotics, and the numbers were taken relative to the total counts. Each data point is the average of two independent experiments.

Figure 3C shows that the relative number of donor cells collected from the flow channel effluents during the first few days after their introduction reflected the differences in the inoculation concentrations. Moreover, there was an initial phase of washing out of this strain, in contrast to what was observed for incoming P. putida RI cells (Fig. 2C). It should also be noted that KT2442, when introduced at a concentration of 5 · 10⁶ CFU/ml, was established in the community at a 1,000-fold-lower level than that observed for the RI strain. Thus, when KT2442 harboring the TOL plasmid was introduced after initial colonization of the three community species it did not establish itself very effectively, and therefore the apparent growth advantage over the indigenous RI strain, as observed in batch cultures of suspended cells, was not sufficient to compete effectively in the biofilm community.

The relative proportion of transconjugants produced upon transfer of the TOLgfpmut3b plasmid to the indigenous P. putida RI cells increased rapidly until it reached a steady-state level of approximately 10% of the total population (Fig. 3D). The accumulation of transconjugants (P. putida RI/TOLgfpmut3b Nal^r) in this experiment and the accumulation of the introduced donor cells (P. putida RI/TOLgfpmut3b Nal^r) in the previous experiment show similar kinetics (compare Fig. 2C and 3D). This suggests that once P. putida RI cells carrying the TOL plasmid are formed or introduced in the biofilm, they grow and become established independently of the way in which they arise in the population. The results also suggest that the dominant mode of establishment of the TOL plasmid in the present community is through the rapid growth of cells representing an optimal host-plasmid combination; actual plasmid transfer plays a quantitatively minor role, which only becomes significant in situations where the donor strain cannot establish itself directly. This suggests that despite the difficulty with which new organisms may become established in already existing microbial communities, new genetic information can be introduced quite effectively when located on mobile elements, even if the actual transfer rates are low.

Similar observations have been made in more complex ecosystems: in soil microcosms (5, 13) and in freshwater flowthrough systems (17, 18), strains carrying catabolic plasmids were found to be outcompeted shortly after their introduction, whereas the plasmids were transferred and established in a number of different indigenous community strains.

Vertical TOL plasmid transfer is dependent on the existing pool of P. putida RI cells in the biofilm.

In the experiments described so far the biofilm community has been kept constant, i.e., the relative proportion of recipient P. putida RI cells has been high, constituting approximately half of the total population (Fig. 2 and 3). The influence of recipient concentration in the flow chambers on the rate of plasmid transfer and replication was investigated subsequently. Three channels were cultivated with the standard mixed-culture inoculum in the first channel (as in the experiments described above), and in the other two channels we reduced the number of inoculated P. putida RI cells 10 and 1,000 times, respectively, relative to the number of P. putida RI cells introduced in the first channel.

Figure 4B shows that, when introduced in low numbers, P. putida RI cells grew rapidly during the first 2 days after colonization, followed by a slow decline. In all three channels, donor cells of the P. putida strain KT2442 harboring the TOL plasmid described above were introduced at day 2 at a concentration of 10⁸ CFU/ml. Figure 4C shows that the relative proportion of donor cells remaining in the biofilm decreased approximately 1 order of magnitude over the next 3 to 4 days in all channels, in agreement with the results shown in Fig. 3C, confirming that this strain of P. putida does not become established easily in the community. Accumulation of transconjugants occurred during the first 2 days (Fig. 4D), but only in the channel with the highest concentration of indigenous P. putida RI cells did accumulation continue. It is striking that even small decreases in the relative numbers of P. putida RI cells in the community eventually led to a cessation of transconjugant accumulation. Thus, the size of the preexisting pool of P. putida RI cells in the biofilm community significantly influenced transconjugant establishment and the introduction of new genetic traits.

FIG. 4.

Time course analysis of the distribution of donor cells, P. putida RI, and transconjugants relative to the total number of cells collected from flow channel effluents. Without changing the inoculation concentration of the two other isolates in the model community, P. putida RI was introduced in three different concentrations of 10⁵ (▴), 10⁷ (▪), and 10⁸ (⧫) CFU/ml. Donor cells (P. putida KT2442/TOLgfpmut3b) (5 · 10⁸ CFU/ml) were introduced at day 2. Total counts (A) were enumerated on pure LB broth plates. The Nal^r P. putida RI cells (B), Rif^r donor cells (C), and Nal^r Km^r transconjugants (D) were enumerated on LB broth plates containing the appropriate antibiotics, and the numbers were taken relative to the total counts. Each point is the average of two individual experiments.

In situ visualization of transconjugants in the biofilm community.

Tagging of the TOL plasmid with a gfp gene (encoding the green fluorescent protein Gfpmut3b) fused to the strong lac promoter P_A1/O4/O3 (see Materials and Methods) allowed us to monitor the in situ establishment of the plasmid within the biofilm. To specifically follow the occurrence and growth of transconjugants, we further inserted the lacI gene in the chromosome of the donor strain, P. putida KT2442, resulting in repression of gfp expression from the plasmid. Thus, expression of the gfp gene will be induced upon transfer of the TOL plasmid to a recipient in the community (P. putida RI) not harboring the lacI gene (zygotic induction of fluorescence).

In an experiment where donor cells were introduced at a concentration of 10⁶ CFU/ml, regions with high densities of transconjugants were observed. One such “high-density zone” was observed for 2 days. Five days after introduction of the donor a strongly green-fluorescent microcolony was observed (Fig. 5A), and other fluorescent microcolonies were found downstream but not upstream of this microcolony. In the following days a progressively increasing number of fluorescent microcolonies were detected in this hot-spot region (Fig. 5B).

FIG. 5.

On-line monitoring of transconjugant proliferation on microcolonies in the direction of flow at days 5 and 6 after donor introduction. The white patches are regions with strong green-fluorescent signal (microcolonies with transconjugants), and the gray regions are weak autofluorescent signals emitted from P. putida cells. This signal is easy to distinguish from the strong green-fluorescent signal emitted from cells expressing Gfp and could be used to visualize the location of noninfected microcolonies. On day 5 (A) a strongly green-fluorescent microcolony (solid arrow) was observed and other green-fluorescent microcolonies were located in a region straight downstream from this colony, but not upstream. On day 6 (B) more green-fluorescent colonies were observed. The scale bar also indicates the direction of flow. Open arrows indicate examples of microcolonies which had been infected with transconjugants from day 5 to day 6.

Based on the results obtained here and described above, we suggest that in the first microcolony shown in Fig. 5A the TOL plasmid initially transferred from a donor cell (P. putida KT2442) to a recipient cell (P. putida RI). The growth advantage of the resulting transconjugant cell allowed the transconjugant cell and its progeny to grow quickly and consequently to produce a cluster of fluorescent cells. Subsequently, some of the cells detached and were carried by the substrate flow further downstream, where they again became established, forming new clusters of transconjugants. According to this interpretation, each “colony” of transconjugants in a flow channel arose from a single horizontal plasmid transfer event followed by effective proliferation of the progeny cells.

The more precise organization of the transconjugants relative to the indigenous recipient cells of P. putida RI and the other members of the community was investigated 8 days after introduction of 10⁸ CFU of donor cells of P. putida KT2442/ml, carrying the gfp-tagged TOL plasmid. The biofilm was fixed and embedded, and in situ hybridization was performed in combination with SCLM. By using the probe PP986 targeting P. putida cells, the spatial distribution of transconjugant (fluorescent) cells and noninfected recipient cells could be visualized. In addition, the probe ACN449 was used to target Acinetobacter, the other dominant member of the community.

Obviously, the donor and recipient P. putida strains cannot be distinguished with the specific 16S ribosomal DNA hybridization probe used here. However, due to the poor establishment of the P. putida KT2442 donor cells relative to the P. putida RI recipient cells (compare Fig. 3C with B), most P. putida cells identified by the PP986 probe were assumed to be P. putida RI cells. Figure 6A shows a representative region of the biofilm.

FIG. 6.

(A) Spatial distribution of green-fluorescent transconjugants (green or yellow) relative to noninfected P. putida RI cells and Acinetobacter sp. strain C6 in a biofilm analyzed 8 days after introduction of donor cells. The organisms P. putida (red) and Acinetobacter sp. strain C6 (purple) were identified by hybridization. After hybridization, green-fluorescent transconjugants appear as either yellow or green, depending on the ratio between the green Gfp signal and the red hybridization signal. The x-y plot is presented as a SFP, where long shadows indicate a large and/or high microcolony. Shown to the right and below are vertical sections through the biofilm collected at the positions indicated by the white triangles. (B) Magnification of a P. putida colony with green-fluorescent cells covering the surface. Vertical sections through the colony are shown to the right and below. The microcolony is a SFP of a region 10 to 19 μm from the glass surface.

Similar to the data presented in Fig. 1A, Acinetobacter was preferentially located in the substratum and P. putida RI cells were located in the upper layers, organized in clusters (microcolonies) sticking up from the surface. With few exceptions, the P. putida RI microcolonies were covered with green-fluorescent transconjugant cells.

A magnification of a microcolony is shown in Fig. 6B. A vertical cross section of the colony further shows how the transconjugants appear in layers of cells covering the colony. Inspection of a number of images like the one presented in Fig. 6 showed that transconjugant cells only very rarely became established as new microcolonies. Instead, they were observed to be preferentially on top of already established microcolonies of recipient cells. This supports the results shown in Fig. 5, where it was observed that the green-fluorescent clusters always showed up on existing microcolonies. Thus, the dominant mode of establishment of cells harboring new genetic information seems to be colonization of existing recipient microcolonies in the flow chambers. This would explain why accumulation of transconjugants in the biofilm is strongly dependent on the relative concentration of recipient P. putida RI cells (as indicated in Fig. 4).

Although the transconjugant and recipient cells seemed to be extremely tightly associated in the microcolonies, horizontal transfer of the TOL plasmid through the entire recipient colony was never observed. This observation is analogous to our previous finding that TOL plasmid transfer between isogenic P. putida KT2442 donor and recipient colonies growing on an agar surface only occurred in a narrow border zone between the two colonies (8).

A number of explanations may account for these observations. Previous studies have shown that TOL plasmid transfer is strongly dependent on the physiological state of the bacteria (36, 37). If the P. putida RI microcolonies contained a steep substrate gradient from the surface to the inner parts, only cells at the surface of the microcolony would be exposed to nutrient concentrations sufficiently high to allow plasmid transfer.

Another explanation could be that although the TOL plasmid has been reported to be naturally derepressed for pilus synthesis (5), this might only be true for cells grown under optimal conditions in well-defined environments. In more complex environments, where cells are organized in dense microbial communities, newly formed transconjugants may be exposed to a number of environmental signals, which could cause repression of pilus synthesis and consequently suppress further conjugation.

It is also possible that the recipient cells in a colony and the newly formed transconjugant cells on the surface of the colony will grow and develop as separate cell lines due to small differences in their phenotypes. Studies of colonies growing on agar surfaces have shown that even very small changes (a single mutation) in a cell may give rise to the formation of colony sectors caused by individual cell lines growing out from the center in separate zones (8, 35).

Finally, it is possible that the Gfp protein does not fold correctly due to a low oxygen tension in the middle of a colony—a possibility that was tested as follows. The biofilm in one flow chamber was disintegrated, and it was found that the relative proportion of green-fluorescent cells constituted approximately 10% of the total population when counted directly under the microscope, a number which correlated well with the proportion of kanamycin- and nalidixic-resistant cells (i.e., transconjugants) determined by plating on the selective plates, strongly indicating that all the transconjugants carrying the TOL plasmid are also green fluorescent.

In conclusion, we have shown that a new biochemical pathway offering a growth-selective advantage can be introduced effectively in a mixed microbial surface community when carried on a conjugative plasmid. Successful establishment of the new genetic information was independent of the efficiency with which the donor strain itself could be established; however, proliferation of the transconjugant cells occurred primarily on the surfaces of preexisting colonies of isogenic recipient cells in the community.