This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Christensen, B. B.
Right arrow Articles by Molin, S.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Christensen, B. B.
Right arrow Articles by Molin, S.
Agricola
Right arrow Articles by Christensen, B. B.
Right arrow Articles by Molin, S.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, May 2002, p. 2495-2502, Vol. 68, No. 5
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.5.2495-2502.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Metabolic Commensalism and Competition in a Two-Species Microbial Consortium

Bjarke B. Christensen,1,2 Janus A. J. Haagensen,1 Arne Heydorn,1 and Søren Molin1*

BioCentrum-DTU, Molecular Microbial Ecology Group, Technical University of Denmark, DK-2800 Lyngby,1 Division of Microbiological Safety, Danish Veterinary and Food Administration, DK-2860 Søborg, Denmark2

Received 23 October 2001/ Accepted 6 February 2002


arrow
ABSTRACT
 
We analyzed metabolic interactions and the importance of specific structural relationships in a benzyl alcohol-degrading microbial consortium comprising two species, Pseudomonas putida strain R1 and Acinetobacter strain C6, both of which are able to utilize benzyl alcohol as their sole carbon and energy source. The organisms were grown either as surface-attached organisms (biofilms) in flow chambers or as suspended cultures in chemostats. The numbers of CFU of P. putida R1 and Acinetobacter strain C6 were determined in chemostats and from the effluents of the flow chambers. When the two species were grown together in chemostats with limiting concentrations of benzyl alcohol, Acinetobacter strain C6 outnumbered P. putida R1 (500:1), whereas under similar growth conditions in biofilms, P. putida R1 was present in higher numbers than Acinetobacter strain C6 (5:1). In order to explain this difference, investigations of microbial activities and structural relationships were carried out in the biofilms. Insertion into P. putida R1 of a fusion between the growth rate-regulated rRNA promoter rrnBP1 and a gfp gene encoding an unstable variant of the green fluorescent protein made it possible to monitor the physiological activity of P. putida R1 cells at different positions in the biofilms. Combining this with fluorescent in situ hybridization and scanning confocal laser microscopy showed that the two organisms compete or display commensal interactions depending on their relative physical positioning in the biofilm. In the initial phase of biofilm development, the growth activity of P. putida R1 was shown to be higher near microcolonies of Acinetobacter strain C6. High-pressure liquid chromatography analysis showed that in the effluent of the Acinetobacter strain C6 monoculture biofilm the metabolic intermediate benzoate accumulated, whereas in the biculture biofilms this was not the case, suggesting that in these biofilms the excess benzoate produced by Acinetobacter strain C6 leaks into the surrounding environment, from where it is metabolized by P. putida R1. After a few days, Acinetobacter strain C6 colonies were overgrown by P. putida R1 cells and new structures developed, in which microcolonies of Acinetobacter strain C6 cells were established in the upper layer of the biofilm. In this way the two organisms developed structural relationships allowing Acinetobacter strain C6 to be close to the bulk liquid with high concentrations of benzyl alcohol and allowing P. putida R1 to benefit from the benzoate leaking from Acinetobacter strain C6. We conclude that in chemostats, where the organisms cannot establish in fixed positions, the two strains will compete for the primary carbon source, benzyl alcohol, which apparently gives Acinetobacter strain C6 a growth advantage, probably because it converts benzyl alcohol to benzoate with a higher yield per time unit than P. putida R1. In biofilms, however, the organisms establish structured, surface-attached consortia, in which heterogeneous ecological niches develop, and under these conditions competition for the primary carbon source is not the only determinant of biomass and population structure.


arrow
INTRODUCTION
 
Bacteria often live in consortia bound to surfaces, such as in biofilms, flocs, or granules (5). Under these conditions the bacteria are positioned in a heterogeneous environment with gradients of nutrients and waste products as a consequence of diffusion and mass transport processes, and it is therefore to be expected that this heterogeneity is reflected in the physiology of the individual cells. In agreement with this, consortia like biofilms often appear as rather complex and heterogeneous assemblies consisting of clusters of bacteria embedded in polymeric substances, which are separated by void regions (cell-free channels) (12, 13, 27, 28). The development of the cell-free regions may support transport of nutrients and waste products to and from the deeper layers of the biofilms (7). For example, DeBeer et al. (8) showed by using microelectrodes that at the same depths in a biofilm, oxygen concentrations in the void regions were much higher than those in adjacent clusters of biomass.

The development of what seem to be structurally organized communities may argue for the presence of overall regulatory elements, which control the formation of the community structures (6). However, changes in structural organization have been shown to be significantly affected by the nutrients supplied to the community (13, 27). Furthermore, mathematical modeling of bacterial growth in biofilms has indicated that simple rules based on nutrient gradients, diffusion rates, and biomass production may determine basic features of biofilm structures (18, 26). Thus, even though there may be regulatory factors that are actively involved in control of biofilm formation, parameters like mass transport, substrate concentrations, diffusion gradients, detachment-attachment mechanisms, and flow rates probably all have significant influence on biofilm structures.

Syntrophic relationships between different organisms have been demonstrated in several microbial ecosystems, such as the interspecies electron transfer from H2 or formate in anaerobic digesters (1, 25) and the relationship between ammonia-oxidizing and nitrite-oxidizing species in nitrifying communities (22). Communities involving xenobiotic degradation (for a review, see reference 21) and oral communities (2) are other examples of tight metabolic associations between community species.

Applications of scanning confocal laser microscopy (SCLM), fluorescence in situ hybridization (FISH), and microelectrodes have led to a rapidly increasing understanding of structure-function relationships in microbial communities. FISH and the use of microelectrodes have shown that the nitrite-oxidizing bacteria are clustered around ammonia-oxidizing bacteria in nitrite-oxidizing zones (inner part of the biofilm) in a nitrifying wastewater treatment biofilm (17). In addition, Ramsing et al. (20) demonstrated a negative correlation between sulfate-reducing bacteria and the oxygen profile in a photosynthetic biofilm, and in anaerobic granular sludge digesters the structural relationship between different species was shown to be highly organized (9, 11, 23).

To obtain a better understanding of the function of channel structures and of structural relationships between species in relation to the overall functionality of the microbial communities, we also need to be able to determine the physiological state of the individual cells in the microbial consortium. Recently, a reporter system based on a fusion between the rRNA promoter from Escherichia coli and a reporter gene encoding an unstable derivative of the green fluorescent protein (GFP) was developed (24). This reporter system has been used to monitor growth activity in the present study of species relationships in benzyl alcohol-degrading communities comprising two species, Acinetobacter sp. strain C6 and Pseudomonas putida strain R1.


arrow
MATERIALS AND METHODS
 
Strain and growth conditions.
Throughout this investigation, derivatives of P. putida R1 and Acinetobacter strain C6 described in Table 1 were used. P. putida R1 (SM1700) with the mini Tn5-Km-rrnBP1-gfp[AGA]-T0-T1 cassette inserted in the chromosome encodes an unstable variant of the GFP (AGA) with a slower degradation half-life, of approximately 6 to 7 h (19), than those previously described (24). The use of a slowly degrading GFP protein reduced the ability to monitor rapid changes in growth activity. However, cells with a much reduced growth rate could be detected when this GFP protein variant was used.


View this table:
[in this window]
[in a new window]
  		TABLE 1. Strains

A streptomycin-resistant mutant of the natural isolate of Acinetobacter strain C6 was isolated as a spontaneous mutant on Luria-Bertani (LB) broth plates containing 200 µg of streptomycin per ml.

Strains were grown in FAB medium [1 mM MgCl2, 0.1 mM CaCl2, 0.01 mM Fe-EDTA (Sigma Chemical Co., St. Louis, Mo.), 0.15 mM (NH4)SO4, 0.33 mM Na2HPO4, 0.2 mM KH2PO4, and 0.5 mM NaCl] supplied, unless otherwise mentioned, with benzyl alcohol (Merck, Darmstadt, Germany) as the sole carbon source. When required, antibiotics were added at final concentrations of 100 µg/ml for streptomycin, 50 µg/ml for nalidixic acid, and 10 µg/ml for kanamycin.

Chemostat experiments.
The chemostats were made from a 50-ml plastic syringe (Terumo Europe N.V., Leuven, Belgium) with a rubber stopper containing a glass tube for outlet of the effluent culture and another for intake of air, which was passed through a 0.2-µm-pore-size filter. In addition, a thin hypodermic needle was inserted for injection of medium and another was inserted for withdrawal of samples from the chemostat. All chemostats were coated with a dimethyl-dichlorosilane solution (Sigma) to avoid attachment of cells to the chemostat walls. In addition, occurrence of bacterial growth on the chemostat wall was tested after each experiment by scraping of samples from the wall followed by plating. In all experiments the total CFU of suspended cells were at least 100 times higher than the number of surface-attached cells, indicating that a potential bias of the chemostat data by development of biofilms on the chemostat walls was insignificant in the present experiments. Six chemostats were run in parallel at the same time. The chemostats were supplied with FAB medium containing 5 mM benzyl alcohol (Merck), and the dilution rate was kept at 6 ml/h with a 205S peristaltic pump (Watson Marlow Inc., Wilmington, Mass.). The chemostats were inoculated with 25 ml of an overnight culture of either P. putida R1 or Acinetobacter strain C6 grown in FAB minimal medium supplemented with 2 mM benzoate and 5 mM benzyl alcohol. In chemostats where P. putida R1 and Acinetobacter strain C6 were mixed, 25 ml of overnight cultures of both strains were mixed before inoculation. The chemostats were sampled at 1-day intervals. For each sample the optical density at 450 nm was measured and the CFU per milliliter for each strain was enumerated by plating on LB broth plates containing antibiotics for selection of P. putida R1 or Acinetobacter strain C6. The chemostats were checked for contaminating organisms by plating on LB broth plates without addition of antibiotics (data not shown).

Flow chamber experiments.
Biofilms were grown at room temperature in three-channel flow chambers with individual channel dimensions of 1 by 4 by 40 mm. The flow system was assembled and prepared as described previously (3). The substratum was a microscope glass coverslip (st1; Knittel Gläser, Braunschweig, Germany). Each channel was supplied with a flow of 3 ml/h (flow rate of 0.2 mm/s) of FAB medium containing 0.5 mM benzyl alcohol. Flow cells were inoculated with mixtures of overnight cultures of P. putida R1 and Acinetobacter strain C6 grown in LB medium. P. putida R1 was diluted 20 times, and Acinetobacter strain C6 was diluted 4 times, in 0.9% NaCl. After the medium flow was stopped, the flow channels were turned upside down and 250 µl of the diluted mixture was carefully injected into each flow channel with a small syringe. After 1 h, the flow channels were turned around and the flow was resumed using a 205S peristaltic pump (Watson Marlow). Enumeration of detaching biofilm cells was performed by collecting 0.5 to 1 ml of cells from the flow chamber effluent in an Eppendorf tube kept on ice. The cells were vortexed for at least 20 s, which was shown by microscopic inspection to be enough to ensure dispersion of cell clumps. The effluent cells were enumerated for each strain by plating on LB broth plates containing antibiotics for selection of P. putida R1 or Acinetobacter strain C6. The flow chambers were checked for contaminating organisms by plating on LB broth plates without antibiotics (data not shown).

Oligonucleotide probes.
For in situ 16S rRNA hybridizations two different oligonucleotide probes were used. A probe specific for P. putida subgroup A, PP986 (14), was labeled with the indocarbocyanine dye CY5, and the probe specific for Acinetobacter sp. strain C6, ACN449 (15), was labeled with the indocarbocyanine dye CY3. Hybridization was performed in 30% formamide at 37°C, as this stringency proved to be sufficient for distinguishing the species in the model consortium. The probes were purchased from Hobolth DNA Syntese (Hillerød, Denmark).

Embedding and 16S rRNA hybridization of hydrated biofilm samples.
In order to avoid further degradation of the unstable GFP, the embedding and hybridization procedure, as previously described (3, 4, 15), was slightly modified. The biofilms were fixed by carefully inoculating 500 µl of ice-cold 4% paraformaldehyde solution directly into the flow channel, which was then kept on ice for at least 1 h to ensure complete fixation of all cells in the biofilm. The fixed biofilms were washed with 1x phosphate-buffered saline by pumping the solution through the channels for 20 min (flow rate, 0.25 mm s-1), and finally the biofilm was embedded in 1 ml of 20% acrylamide solution containing 200:1 acrylamide-bisacrylamide (Sequagel; National Diagnostics, Atlanta, Ga.), 8 µl of N,N,N',N'-tetramethylethylendiamine (Kodak International Biotechnologies Inc., New Haven, Conn.), and 20 µl of 1% ammonium persulfate (International Biotechnologies Inc.). After the polyacrylamide was allowed to solidify for at least 1 h, the glass coverslip was carefully loosened from the flow cell, and the biofilm containing the polyacrylamide block was lifted out of the flow channel. The block was subsequently cut into slices of approximately 5 mm, placed on a six-well hybridization slide (Novakemi ab, Enskede, Sweden), and prehybridized at 37°C in 45 µl of the hybridization buffer (washing solution I [0.9 M NaCl, 100 mM Tris, pH 7.2] containing 30% formamide at 37°C). After 30 min, the prehybridization buffer was removed and 30 µl of the hybridization buffer containing 75 ng of each probe was added to the hybridization well. The slide was incubated at 37°C for at least 3 h in a moisturized chamber. The polyacrylamide blocks were washed in 45 µl of washing solution I for 30 min at 37°C, followed by washing in 45 µl of washing solution II (0.9 M NaCl, 100 mM Tris, pH 7.2) for another 30 min at 37°C. Finally, the acrylamide blocks were rinsed in 45 µl of Milli-Q water and then immediately mounted on an object glass with a drop of SlowFade phosphate-buffered saline-based antifade solution (Molecular Probes) and a coverslip on top.

Microscopy and image analysis.
All microscopic observations and image acquisitions were performed with a TCD4D SCLM (Leica Lasertechnik GmbH, Heidelberg, Germany) equipped with an argon-krypton laser and three detectors and filter sets for simultaneous monitoring of fluorescein isothiocyanate-GFP and the indocarbocyanine dyes CY3 and CY5.

The x-y images were presented as extended-focus images, which are produced by taking the confocal images from the different depths of the biofilm and projecting them into a single image. The extended-focus images and vertical cross sections through the biofilm were generated by using the IMARIS software package (Bitplane AG, Zurich, Switzerland) running on an Indigo2 workstation (Silicon Graphics, Mountain View, Calif.). Images were further processed for display by using Photoshop software (Adobe, Mountain View, Calif.)

HPLC analysis.
Samples subjected to high-performance liquid chromatography (HPLC) analysis were taken from the effluent of the flow channels. The samples were filtered through a 0.2-µm-pore-size filter, and the contents of benzyl alcohol and benzoate were measured with a Shimadzu (Tokyo, Japan) HPLC equipped with a Supelcosil C18 reverse-phase column (Supelco Park, Bellefonte, Pa.) and a UV-visible detector set at 206 nm. The mobile phase was a solution of 40% acetonitrile (HPLC grade; Sigma Aldrich) and 60% NaH2PO4 (50 mM, pH 3,0) supplied at a flow rate of 1 ml min-1.

Analysis of biofilm thickness.
The thicknesses of the biofilms were measured using a specific function on the digitally controlled microscope (DMRXA microscope; Leica Mikroskopie und Systeme GmbH, Wetzlar, Germany) which makes it possible to measure the distance between two focused planes.


arrow
RESULTS
 
Quantitative analysis of mixed-species and monospecies consortia of P. putida R1 and Acinetobacter strain C6 in chemostats and in flow chambers.
P. putida R1 and Acinetobacter strain C6 were grown as suspended cultures in chemostats, in which heterogeneous structural relationships between the organisms are not established. Acinetobacter strain C6 and P. putida R1 were established either as monocultures (Fig. 1A) or together in a mixed culture (Fig. 1B). As shown in Fig. 1, constant cell densities were reached in the monoculture and mixed-culture chemostats after approximately 3 and 5 days, respectively. With respect to Acinetobacter strain C6, the cell density remained constant in both chemostats over the course of the next 5 days until the experiment was terminated, whereas a slight increase in P. putida R1 cell numbers was observed during the last 2 days. In both the mixed and monospecies cultures Acinetobacter strain C6 reached the same cell density of approximately 109 CFU/ml. In contrast, the cell densities of P. putida R1 were lower than the corresponding levels of Acinetobacter strain C6 in both the mixed-species and the monospecies chemostats. When growing alone in the chemostat, the cell density of P. putida R1 was between 10 and 100 times lower than that of Acinetobacter strain C6, whereas in the chemostats in which Acinetobacter strain C6 and P. putida R1 were mixed, the density of P. putida R1 was reduced to a 500- to 1,000-fold-lower level relative to that of Acinetobacter strain C6. These results show that Acinetobacter strain C6 produces more biomass than P. putida R1.



View larger version (8K):
[in this window]
[in a new window]
  		FIG. 1. Time course analysis of the numbers of P. putida R1 ({blacksquare}) and Acinetobacter strain C6 (•) cells collected from chemostats where the strains were established either as monospecies cultures (A) or as mixed cultures (B). CFU were enumerated on LB plates containing the appropriate antibiotics for selection of P. putida R1 and Acinetobacter strain C6, respectively. Error bars indicate standard deviations.

To investigate how the observed differences in the metabolic efficiency affected the composition in surface-associated cultures over time, nondestructive in situ methods were employed. Two parameters were measured: (i) biofilm thickness and (ii) CFU of cells collected from the flow chamber effluents per milliliter. Although the ratios between biomass, voids, and channels may vary in internal parts of the biofilm (which was not taken into account when measuring biofilm thickness), the thickness of the biofilm may, as shown for other biofilm consortia (10), be used as a relative measure of the total biomass accumulated in the flow channels. As shown in Fig. 2, both Acinetobacter strain C6 and P. putida R1 monocultures reached a thickness of about 7 to 10 µm, whereas the binary biofilm reached a thickness of approximately 12 to 14 µm.



View larger version (13K):
[in this window]
[in a new window]
  		FIG. 2. Time course analysis of biofilm thickness when P. putida R1 ({blacksquare}) and Acinetobacter strain C6 (•) were grown as monoculture biofilms or when the two strains were grown as mixed biofilms ({blacktriangleup}). Each point represents the mean of the mean thickness in at least three independent flow channels run in parallel. Error bars indicate standard deviations. The mean thickness in each flow channel was taken as the mean from four random microscope viewing fields in which the thickness was measured at five different positions (a total of 20 positions in each flow channel).

The cells collected from the effluent are those cells that detach from the flow chamber biofilm. Only in cases where the biofilm cell density has reached a steady state for each species will the effluent cells become an exact representation of the biofilm population. On the other hand, previous studies have shown (3) that in control experiments in which Acinetobacter strain C6 and P. putida R1 were the only species or the predominant species, the effluent data do represent reliable estimates of the entire biofilm population. Determinations of the biofilm effluent cell numbers showed that a ratio of approximately 1:5 of Acinetobacter strain C6 to P. putida R1 in the mixed biofilm was reached after a few days (Fig. 3B). Varying the ratio of Acinetobacter strain C6 and P. putida R1 in the inoculum did not change this ratio significantly (data not shown). After 2 to 3 days the effluent cell numbers of Acinetobacter strain C6 in the monoculture (Fig. 3A) and mixed-culture (Fig. 3B) biofilms had reached a level which remained nearly constant throughout the rest of the experiment. In the monoculture biofilm of P. putida R1, the cell number increased at a lower rate but reached higher numbers than Acinetobacter strain C6 (Fig. 3A), whereas a constant high level of P. putida R1 in the mixed biofilm was reached within a few days (Fig. 3B). Thus, the presence of Acinetobacter strain C6 apparently results in a faster establishment of P. putida R1 in the mixed consortium.



View larger version (9K):
[in this window]
[in a new window]
  		FIG. 3. Time course analysis of the numbers of P. putida R1 ({blacksquare}) and Acinetobacter strain C6 (•) cells collected from flow channel effluents where the strains were established either as monoculture biofilms (A) or as mixed biofilms (B). CFU were enumerated on LB plates containing the appropriate antibiotics for selection of P. putida R1 and Acinetobacter strain C6. Error bars indicate standard deviations.

It was previously observed that in mixed-species biofilms growing with benzyl alcohol as the sole carbon and energy source, the expression of the benzoate-inducible promoter, Pm, from the TOL pathway inserted into P. putida R1 was induced in regions near microcolonies of Acinetobacter strain C6 (15), and it was speculated that benzoate leaking from Acinetobacter strain C6 caused the induction of Pm in P. putida R1. In order to investigate this explanation, we performed HPLC analysis of benzyl alcohol and benzoate in flow channel effluents, and at the same time this analysis was used to further assess the metabolic efficiencies of the two organisms.

Samples were taken at different time points after initial colonization of mono- and mixed-culture flow chamber biofilms of P. putida R1 and Acinetobacter strain C6. In the P. putida R1 monoculture biofilm (Fig. 4A), the amount of degraded benzyl alcohol increased slowly over the entire sampling period, reaching 75% conversion after 10 days. In the Acinetobacter strain C6 monoculture biofilm, more than 90% of the benzyl alcohol was catabolized 4 days after the initial colonization, but in contrast to the case for the P. putida R1 biofilm, a large amount of benzoate was accumulated (Fig. 4B). Thus, Acinetobacter strain C6 does in fact leak benzoate from cells when grown as a biofilm. The effluent from a mixed culture of the two species showed removal of benzyl alcohol as well as efficient degradation of benzoate (Fig. 4).



View larger version (10K):
[in this window]
[in a new window]
  		FIG. 4. HPLC analysis of the flow channel effluents. The contents of benzyl alcohol (A) and benzoate (B) were measured for monoculture biofilms of P. putida R1 ({blacksquare}) and Acinetobacter strain C6 (•), and for the binary Acinetobacter strain C6-P. putida R1 biofilm ({blacktriangleup}). In all flow channel biofilms the inlet concentration of benzyl alcohol was 0.5 mM. Error bars indicate standard deviations.

In situ analysis of the mixed biofilm consortium.
To further analyze the metabolic interactions between Acinetobacter strain C6 and P. putida R1 cells and their consequences for the structural development of the consortium, mixed-species biofilms were fixed and embedded 1, 2, 3, and 6 days after the initial colonization. For specific identification of the two organisms, we used FISH with the fluorescence-labeled probes PP986 and ACN449, targeting P. putida R1 and Acinetobacter strain C6, respectively. By employing SCLM in combination with FISH, the exact positions of Acinetobacter strain C6 and P. putida R1 cells could be visualized. The changes and distributions of microbial growth activity of P. putida R1 cells were analyzed by insertion of a transposon with the growth phase-regulated promoter rrnBP1 fused to a gfp gene encoding an unstable variant of GFP. Applications of this type of monitor system inserted into P. putida R1 for determinations of growth activity in biofilms have previously been demonstrated (24). This reporter system can be used to distinguish between fast-growing (bright green) and slow-growing or nongrowing (weak or no signal) cells.

The images presented in Fig. 5 represent examples of the most frequently occurring structural relationships between the two species observed at days 1, 2, 3, and 6 after the initial colonization. In order to ensure that the changing structures observed were a consequence of a systematic process and not just some random process occurring in some cases and not in others, the experiments was run in five independent rounds each time with three independent flow channels running in parallel. In all experiments the same developmental pattern was observed, indicating that the dynamic changes in the structural relationships between the two species are caused by nonrandom processes.



View larger version (93K):
[in this window]
[in a new window]
  		FIG. 5. SCLM micrographs showing the structural relationships between Acinetobacter strain C6 and P. putida R1 cells with low and high activity, respectively, in a mixed biofilm consortium which was supplied with 0.5 mM benzyl alcohol as the sole carbon source. At days 1 (A), 2 (B), 3 (C and E), and 6 (D and F) after inoculation, biofilms were embedded and hybridized. P. putida R1 and Acinetobacter strain C6 were hybridized with PP986 labeled with CY5 (blue) and ACN449 labeled with CY3 (red), respectively. The active P. putida R1 cells were monitored as cells emitting green fluorescence due to the rrnBP1-gfp[AGA] fusion inserted in the chromosome of P. putida R1. These cells appear as cyan due tothe combination of green (GFP) and blue (hybridization). For each panel similar images were collected from at least five independent biofilm experiments. Panels A and B are representative of the biofilm structures observed at day 1 and 2. Panels C and D are examples of the large Acinetobacter strain C6 microcolonies that developed after 3 days (C) and which later were overgrown by P. putida R1 (D). Panels E and F are examples of Acinetobacter strain C6 microcolonies (arrow) that were established in the upper part of P. putida R1 cell clusters after 2 to 3 days (E), which resulted in production of large macrostructures of associated P. putida R1 and Acinetobacter strain C6 cells (F). All x-y plots are presented as extended-focus images. Shown to the right and above the x-y plots are vertical sections through the biofilm collected at the positions indicated by the white triangles. The arrow indicates the direction of flow. Bars, 20 µm. Note that the green fluorescent intensities on the images have been amplified to give the best differentiation between P. putida R1 cells with low and high activity in each image. Thus, the intensity levels in the different images cannot be directly compared.

One day after inoculation of the strains, the P. putida R1 cells with the highest apparent growth activity (highest GFP signal) were observed in the regions near small microcolonies of Acinetobacter strain C6 cells (Fig. 5A). As the Acinetobacter strain C6 microcolonies grew in size, the P. putida R1 cells with highest growth activity were located in the center of the colonies (Fig. 5B). At day 3, two distinct types of associations had developed. In one of these, P. putida R1 was excluded from the center of the rather large Acinetobacter strain C6 colonies (Fig. 5C), and the cells with the highest growth activity were now observed at the periphery of the colonies. After 6 days, many of these Acinetobacter strain C6 microcolonies were overgrown by P. putida R1 cells and the fluorescence intensity of the P. putida R1 cells near the Acinetobacter strain C6 colonies was reduced to low levels (Fig. 5D). In the other type of association, small Acinetobacter strain C6 microcolonies started to build up in the upper layers of the P. putida R1 biofilm (Fig. 5E). The result was production of large structures (at least 100 to 200 µm thick) in which Acinetobacter strain C6 microcolonies became integrated in the P. putida R1 cell structures (Fig. 5F).


arrow
DISCUSSION
 
Competition for substrate is considered to be one of the major evolutionary driving forces in the bacterial world, and numerous experimental data obtained in the laboratory under well-controlled conditions show how different organisms, or variants of one organism, may effectively outcompete others because of better utilization of a given energy source. Such competition experiments may be performed very convincingly in chemostats, where nutrient limitation allows for metabolic competition among the cells present in the reactor. One important factor for optimal performance of a chemostat is the homogeneous distribution of the suspended cells, and it is considered important to prevent surface attachment of the cells to the reactor walls. Under such optimal conditions, in which only one nutrient is limiting for growth, it may be expected that an organism will totally outcompete all others if it has an improved efficiency of substrate utilization relative to the others. The concentration of the particular nutrient will always be below the threshold level for the less efficient cells, and the rate of washout is therefore higher than the rate of cell proliferation for these cells.

In the present study the two organisms were grown as monospecies and as mixed-species cultures with benzyl alcohol as the sole carbon source, both as surface-attached organisms (biofilms) in flow chambers and as suspended cultures in chemostats. In the chemostats, differences in the respective cell numbers at steady state showed that Acinetobacter strain C6 produced the highest yield on the limiting concentration of benzyl alcohol. When Acinetobacter strain C6 and P. putida R1 were mixed in the chemostat, both organisms rapidly reached relatively stable cell densities in which the cell number ratio was approximately 1 to 500 in favor of Acinetobacter strain C6. This again showed the competitive advantage of Acinetobacter strain C6, but it also showed an unexpected stable presence of P. putida R1 at a low but significant level. In the flow chamber-based biofilms, effluent cell counts showed that within a few days the two organisms reached nearly stable cell densities, in which P. putida R1 was present in higher numbers than Acinetobacter strain C6 (approximately 5 to 1). Furthermore, measurements of the biofilm thickness suggested that the simultaneous presence of both strains resulted in a significant increase of the overall biomass in the biofilm. Thus, despite the better utilization of benzyl alcohol by Acinetobacter strain C6, P. putida R1 was maintained in both growth systems. This finding may be explained by the previous suggestion that growing with benzyl alcohol as the carbon source makes Acinetobacter strain C6 cells excrete benzoate, a carbon source readily utilized by P. putida R1, into the reactor environment (15). This hypothesis has been confirmed in the present study. The HPLC analysis of the effluent from flow chambers shows that benzoate accumulates in Acinetobacter strain C6 monospecies biofilms, whereas in the presence of P. putida R1 almost all the excreted benzoate is degraded. In the P. putida R1 monospecies biofilm, degradation of benzyl alcohol was rather inefficient, and only a low concentration of benzoate was detected in the effluent. This suggests that Acinetobacter strain C6 relatively easily converts benzyl alcohol to benzoate, whereas the degradation of benzoate is slow, resulting in a metabolic bottleneck that leads to accumulation of benzoate. In contrast, P. putida R1 encodes a better pathway for degradation of benzoate, which has the capacity to mineralize both the benzoate derived from the conversion of benzyl alcohol to benzoate via its own degradation pathway and the external supply from Acinetobacter strain C6.

To further explain the differences in ratios between P. putida R1 and Acinetobacter strain C6 in the flow chambers and chemostats, a detailed analysis of the distribution and activity of the P. putida R1 cells in the flow chamber biofilms was carried out. In situ rRNA hybridization for identification of the individual organisms was used to analyze the spatial distribution of the two strains in the flow chamber biofilms. After 2 to 3 days of growth, large numbers of P. putida R1 cells were found to cluster around large surface-associated microcolonies of Acinetobacter strain C6. By employing the rrnBP1::gfp[AGA] monitor cassette inserted into P. putida R1, it was shown that the P. putida R1 cells in these regions had a higher growth activity than those further away from the Acinetobacter strain C6 microcolonies. Similar distribution patterns have also been observed in nitrifying biofilms (17) and in a two-species biofilm growing with chlorobiphenyl as the sole carbon and energy source (16). However, in neither of these studies was it possible for the individual strains to be established as monospecies biofilms under the conditions described, and thus, in contrast to the present study, there was no competition for the primary carbon source in the mixed biofilms.

For the Acinetobacter strain C6 microcolonies that were initially established at the flow chamber glass surface, a stable structural relationship between P. putida R1 and Acinetobacter strain C6 cells was not maintained, because P. putida R1 started to overgrow the Acinetobacter strain C6 microcolonies (Fig. 5C and D). The result was a significant reduction in the growth activity of P. putida R1 (reduced GFP signal) in these regions. We suggest that the establishment of P. putida R1 cells on the outside of the surface-associated microcolonies reduced the supply of benzyl alcohol to the Acinetobacter strain C6 cells and thereby also the production of benzoate from these cells. Instead, small microcolonies of Acinetobacter strain C6 cells became established in the upper layer of the P. putida R1 biofilm where they were near the medium flow and thereby exposed to a constant supply of benzyl alcohol. This allowed the Acinetobacter strain C6 cells to accumulate benzoate in these regions, which was metabolized by the associated P. putida R1 cells, which obtained a growth advantage over other, nonassociated P. putida R1 cells (Fig. 5E). We suggest that due to the metabolic interactions, these mixed Acinetobacter strain C6-P. putida R1 structures grew faster than the rest of the biofilm and resulted in the production of large structures of biofilm. Thus, taking all these observations together, the two organisms in the biofilm competed as well as exhibited commensal interactions, depending on the physical positioning of the organism. Similar interacting mechanisms, i.e., different species being able to degrade the primary substrate but having different capacities for degradation of the metabolic intermediates, may be expected to prevail in natural environments. This may explain why some natural communities grown on a single carbon source, such as in the studies presented by Wolfaardt et al. (27) and Møller et al. (13), often develop quite complex biofilm compositions and organizations.

The present investigation of the structure-function relationships in a binary biofilm growing with benzyl alcohol as the only added carbon and energy source thus offers an explanation of the contradictory population data from suspended cultures in chemostats and in flow chamber biofilms, respectively. In the chemostat the individual cells are all surrounded by the same environmental conditions, and they cannot stay in the reactor if their growth rate is lower than the dilution rate (no adherence). In the biofilm there is a range of conditions surrounding the cells due to the heterogeneity of the consortium structure; i.e., microniches develop in which the supplies of primary and secondary nutrients differ significantly. There will therefore be locations in the biofilm where the conditions favor one or the other or both of the organisms, depending on the local structure. In addition, the cells may adhere to the surface or to each other, thus reducing washout. One important consequence of the biofilm configuration, therefore, is that substrates may be more optimally utilized by the consortium (increased mineralization of metabolic intermediates), resulting in faster degradation of the primary nutrient and a faster buildup of biomass. The reorganization of mixed-species consortia in response to the nutrient conditions in order to achieve optimal conditions for the present organisms may require active motility coupled with chemotaxis, cell-to-cell communication signals for coordinated organizational development, or a liquid flow moving cells around for detachment and reattachment. In the present consortium neither chemotaxis nor cell-to-cell communication is the obvious mechanism behind the presented development of the consortium, because our unpublished investigations have not so far shown evidence of any of these properties in P. putida R1. It is therefore concluded that passive transport of cells by the flow through the biofilm may be solely responsible for the continuous structural development taking place in the consortium.


arrow
ACKNOWLEDGMENTS
 
This work was supported by grants from the Danish Biotechnology Program.

We thank Anne Nielsen and Tove Johansen for expert technical assistance.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: BioCentrum-DTU, Molecular Microbial Ecology Group, Building 301, The Technical University of Denmark, DK-2800 Lyngby, Denmark. Phone: 45 45 25 25 13. Fax: 45 45 88 73 28. E-mail: imsm{at}pop.dtu.dk. Back


arrow
REFERENCES
 
    1
  1. Boone, D. R., R. L. Johnson, and Y. Liu. 1989. Diffusion of the interspecies electron carriers H2 and formate in methanogenic ecosystems and its implications in the measurement of Km for H2 or formate uptake. Appl. Environ. Microbiol. 55:1735-1741.[Abstract/Free Full Text]
    2. 2
  2. Bradshaw, D. J., K. A. Homer, P. D. Marsh, and D. Beighton. 1994. Metabolic cooperation in oral microbial communities during growth on mucin. Microbiology 140:3407-3412.[Abstract/Free Full Text]
    3. 3
  3. Christensen, B., C. Sternberg, J. B. Andersen, L. Eberl, S. Møller, M. Givskov, and S. Molin. 1998. Establishment of new genetic traits in a microbial biofilm community. Appl. Environ. Microbiol. 64:2247-2255.[Abstract/Free Full Text]
    4. 4
  4. Christensen, B. B., C. Sternberg, J. B. Andersen, R. J. Palmer, A. T. Nielsen, M. Givskov, and S. Molin. 1999. Molecular tools for study of biofilm physiology. Methods Enzymol. 310:20-42.[Medline]
    5. 5
  5. Costerton, J. W., Z. Lewandowski, D. E. Caldwell, D. R. Korber, and H. M. Lappin-Scott. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49:711-745.[CrossRef][Medline]
    6. 6
  6. Davies, D. G., M. R. Parsek, J. P. Pearson, B. H. Iglewski, J. W. Costerton, and E. P. Greenberg. 1998. The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280:295-298.[Abstract/Free Full Text]
    7. 7
  7. DeBeer, D., P. Stoodley, and Z. Lewandowski. 1994. Liquid flow in heterogeneous biofilms. Biotechnol. Bioeng. 44:636-641.[CrossRef]
    8. 8
  8. DeBeer, D., P. Stoodley, F. Roe, and Z. Lewandowski. 1994. Effects of biofilm structures on oxygen distribution and mass-transport. Biotechnol. Bioeng. 43:1131-1138.[CrossRef]
    9. 9
  9. Harmsen, H. J., H. M. Kengen, A. D. Akkermans, A. J. Stams, and W. M. de Vos. 1996. Detection and localization of syntrophic propionate-oxidizing bacteria in granular sludge by in situ hybridization using 16S rRNA-based oligonucleotide probes. Appl. Environ. Microbiol. 62:1656-1663.[Abstract]
    10. 10
  10. Heydorn, A., A. T. Nielsen, M. Hentzer, C. Sternberg, M. Givskov, B. K. Ersbøll, and S. Molin. 2000. Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology 146:2395-2407.[Abstract/Free Full Text]
    11. 11
  11. MacLeod, F. A., S. R. Guiot, and J. W. Costerton. 1990. Layered structure of bacterial aggregates produced in an upflow anaerobic sludge bed and filter reactor. Appl. Environ. Microbiol. 56:1598-1607.[Abstract/Free Full Text]
    12. 12
  12. Massol-Deya, A. A., J. Whallon, R. F. Hickey, and J. M. Tiedje. 1995. Channel structures in aerobic biofilms of fixed-film reactors treating contaminated groundwater. Appl. Environ. Microbiol. 61:769-777.[Abstract]
    13. 13
  13. Møller, S., D. R. Korber, G. M. Wolfaardt, S. Molin, and D. E. Caldwell. 1997. Impact of nutrient composition on a degradative biofilm community. Appl. Environ. Microbiol. 63:2432-2438.[Abstract]
    14. 14
  14. Møller, S., A. R. Pedersen, L. K. Poulsen, E. Arvin, and S. Molin. 1996. Activity and three-dimensional distribution of toluene-degrading Pseudomonas putida in a multispecies biofilm assessed by quantitative in situ hybridization and scanning confocal laser microscopy. Appl. Environ. Microbiol. 62:4632-4640.[Abstract]
    15. 15
  15. Møller, S., C. Sternberg, J. B. Andersen, B. B. Christensen, J. L. Ramos, M. Givskov, and S. Molin. 1998. In situ gene expression in mixed-culture biofilms: evidence of metabolic interactions between community members. Appl. Environ. Microbiol. 64:721-732.[Abstract/Free Full Text]
    16. 16
  16. Nielsen, A. T., T. Tolker-Nielsen, K. B. Barken, and S. Molin. 2000. Role of commensal relationships on the spatial structure of a surface-attached microbial consortium. Environ. Microbiol. 2:59-68.[CrossRef][Medline]
    17. 17
  17. Okabe, S., H. Satoh, and Y. Watanabe. 1999. In situ analysis of nitrifying biofilms as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 65:3182-3191.[Abstract/Free Full Text]
    18. 18
  18. Picioreanu, C., M. C. M. Van Loosdrecht, and J. J. Heijnen. 1998. Mathematical modeling of biofilm structure with a hybrid differential-discrete cellular automaton approach. Biotechnol. Bioeng. 58:101-116.[CrossRef][Medline]
    19. 19
  19. Ramos, C., L. Molbak, and S. Molin. 2000. Bacterial activity in the rhizosphere analyzed at the single-cell level by monitoring ribosome contents and synthesis rates. Appl. Environ. Microbiol. 66:801-809.[Abstract/Free Full Text]
    20. 20
  20. Ramsing, N. B., M. Kuhl, and B. B. Jørgensen. 1993. Distribution of sulfate-reducing bacteria, O2, and H2S in photosynthetic biofilms determined by oligonucleotide probes and microelectrodes. Appl. Environ. Microbiol. 59:3840-3849.[Abstract/Free Full Text]
    21. 21
  21. Reanney, D., P. C. Gowland, and J. H. Slater. 1983. Genetic interactions among microbial communities. Symp. Soc. Gen. Microbiol. 34:379-422.
    22. 22
  22. Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K. H. Schleifer. 1996. Structure and function of a nitrifying biofilm as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 62:4641-4647.[Abstract]
    23. 23
  23. Sekiguchi, Y., Y. Kamagata, K. Nakamura, A. Ohashi, and H. Harada. 1999. Fluorescence in situ hybridization using 16S rRNA-targeted oligonucleotides reveals localization of methanogens and selected uncultured bacteria in mesophilic and thermophilic sludge granules. Appl. Environ. Microbiol. 65:1280-1288.[Abstract/Free Full Text]
    24. 24
  24. Sternberg, C., B. B. Christensen, T. Johansen, A. T. Nielsen, J. B. Andersen, M. Givskov, and S. Molin. 1999. Distribution of bacterial growth activity in flow-chamber biofilms. Appl. Environ. Microbiol. 65:4108-4117.[Abstract/Free Full Text]
    25. 25
  25. Thiele, J. H., M. Chartrain, and J. G. Zeikus. 1988. Control of interspecies electron flow during anaerobic digestion: role of floc formation in syntrophic methanogenesis. Appl. Environ. Microbiol. 54:10-19.[Abstract/Free Full Text]
    26. 26
  26. Wimpenny, J. W. T., and R. Colasanti. 1997. A unifying hypothesis for the structure of microbial biofilms based on cellular automaton models. FEMS Microbiol. Ecol. 22:1-16.
    27. 27
  27. Wolfaardt, G. M., J. R. Lawrence, R. D. Robarts, S. J. Caldwell, and D. E. Caldwell. 1994. Multicellular organization in a degradative biofilm community. Appl. Environ. Microbiol. 60:434-446.[Abstract/Free Full Text]
    28. 28
  28. Zahid, W. M., and J. J. Ganzarczyk. 1994. Fractal properties of the RBC biofilm structure. Wat. Sci. Technol. 29:271-279.

Applied and Environmental Microbiology, May 2002, p. 2495-2502, Vol. 68, No. 5
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.5.2495-2502.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Min, K. R., Rickard, A. H. (2009). Coaggregation by the Freshwater Bacterium Sphingomonas natatoria Alters Dual-Species Biofilm Formation. Appl. Environ. Microbiol. 75: 3987-3997 [Abstract] [Full Text]  
  • Slater, F. R., Bruce, K. D., Ellis, R. J., Lilley, A. K., Turner, S. L. (2008). Heterogeneous Selection in a Spatially Structured Environment Affects Fitness Tradeoffs of Plasmid Carriage in Pseudomonads. Appl. Environ. Microbiol. 74: 3189-3197 [Abstract] [Full Text]  
  • Barreiros, L., Fernandes, A., Ferreira, A. C. S., Pereira, H., Bastos, M. M. S. M., Manaia, C. M., Nunes, O. C. (2008). New insights into a bacterial metabolic and detoxifying association responsible for the mineralization of the thiocarbamate herbicide molinate. Microbiology 154: 1038-1046 [Abstract] [Full Text]  
  • Brenner, K., Karig, D. K., Weiss, R., Arnold, F. H. (2007). Engineered bidirectional communication mediates a consensus in a microbial biofilm consortium. Proc. Natl. Acad. Sci. USA 104: 17300-17304 [Abstract] [Full Text]  
  • Simoes, L. C., Simoes, M., Vieira, M. J. (2007). Biofilm Interactions between Distinct Bacterial Genera Isolated from Drinking Water. Appl. Environ. Microbiol. 73: 6192-6200 [Abstract] [Full Text]  
  • Merod, R. T., Warren, J. E., McCaslin, H., Wuertz, S. (2007). Toward Automated Analysis of Biofilm Architecture: Bias Caused by Extraneous Confocal Laser Scanning Microscopy Images. Appl. Environ. Microbiol. 73: 4922-4930 [Abstract] [Full Text]  
  • Hansen, S. K., Haagensen, J. A. J., Gjermansen, M., Jorgensen, T. M., Tolker-Nielsen, T., Molin, S. (2007). Characterization of a Pseudomonas putida Rough Variant Evolved in a Mixed-Species Biofilm with Acinetobacter sp. Strain C6. J. Bacteriol. 189: 4932-4943 [Abstract] [Full Text]  
  • Moons, P., Van Houdt, R., Aertsen, A., Vanoirbeek, K., Engelborghs, Y., Michiels, C. W. (2006). Role of Quorum Sensing and Antimicrobial Component Production by Serratia plymuthica in Formation of Biofilms, Including Mixed Biofilms with Escherichia coli. Appl. Environ. Microbiol. 72: 7294-7300 [Abstract] [Full Text]  
  • Laue, H., Schenk, A., Li, H., Lambertsen, L., Neu, T. R., Molin, S., Ullrich, M. S. (2006). Contribution of alginate and levan production to biofilm formation by Pseudomonas syringae.. Microbiology 152: 2909-2918 [Abstract] [Full Text]  
  • Peterson, S. B., Dunn, A. K., Klimowicz, A. K., Handelsman, J. (2006). Peptidoglycan from Bacillus cereus Mediates Commensalism with Rhizosphere Bacteria from the Cytophaga-Flavobacterium Group. Appl. Environ. Microbiol. 72: 5421-5427 [Abstract] [Full Text]  
  • Reisner, A., Holler, B. M., Molin, S., Zechner, E. L. (2006). Synergistic Effects in Mixed Escherichia coli Biofilms: Conjugative Plasmid Transfer Drives Biofilm Expansion.. J. Bacteriol. 188: 3582-3588 [Abstract] [Full Text]  
  • Bruhn, J. B., Haagensen, J. A. J., Bagge-Ravn, D., Gram, L. (2006). Culture Conditions of Roseobacter Strain 27-4 Affect Its Attachment and Biofilm Formation as Quantified by Real-Time PCR. Appl. Environ. Microbiol. 72: 3011-3015 [Abstract] [Full Text]  
  • Kives, J., Guadarrama, D., Orgaz, B., Rivera-Sen, A., Vazquez, J., SanJose, C. (2005). Interactions in Biofilms of Lactococcus lactis ssp. cremoris and Pseudomonas fluorescens Cultured in Cold UHT Milk. J DAIRY SCI 88: 4165-4171 [Abstract] [Full Text]  
  • Rice, S. A., Koh, K. S., Queck, S. Y., Labbate, M., Lam, K. W., Kjelleberg, S. (2005). Biofilm Formation and Sloughing in Serratia marcescens Are Controlled by Quorum Sensing and Nutrient Cues. J. Bacteriol. 187: 3477-3485 [Abstract] [Full Text]  
  • Egland, P. G., Palmer, R. J. Jr., Kolenbrander, P. E. (2004). Interspecies communication in Streptococcus gordonii-Veillonella atypica biofilms: Signaling in flow conditions requires juxtaposition. Proc. Natl. Acad. Sci. USA 101: 16917-16922 [Abstract] [Full Text]  
  • Yi, K., Rasmussen, A. W., Gudlavalleti, S. K., Stephens, D. S., Stojiljkovic, I. (2004). Biofilm Formation by Neisseria meningitidis. Infect. Immun. 72: 6132-6138 [Abstract] [Full Text]  
  • Kreft, J.-U. (2004). Biofilms promote altruism. Microbiology 150: 2751-2760 [Abstract] [Full Text]  

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Christensen, B. B.
Right arrow Articles by Molin, S.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Christensen, B. B.
Right arrow Articles by Molin, S.
Agricola
Right arrow Articles by Christensen, B. B.
Right arrow Articles by Molin, S.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»