Abstract
Horizontal gene transfer by natural genetic transformation in Acinetobacter sp. strain BD413 was investigated by using gfp carried by the autonomously replicating plasmid pGAR1 in a model monoculture biofilm. Biofilm age, DNA concentration, and biofilm mode of growth were evaluated to determine their effects on natural genetic transformation. The highest transfer frequencies were obtained in young and actively growing biofilms when high DNA concentrations were used and when the biofilm developed during continuous exposure to fresh medium without the presence of a significant amount of cells in the suspended fraction. Biofilms were highly amenable to natural transformation. They did not need to advance to an optimal growth phase which ensured the presence of optimally competent biofilm cells. An exposure time of only 15 min was adequate for transformation, and the addition of minute amounts of DNA (2.4 fg of pGAR1 per h) was enough to obtain detectable transfer frequencies. The transformability of biofilms lacking competent cells due to growth in the presence of cells in the bulk phase could be reestablished by starving the noncompetent biofilm prior to DNA exposure. Overall, the evidence suggests that biofilms offer no barrier against effective natural genetic transformation of Acinetobacter sp. strain BD413.
A great variety of synthetic chemicals have been released into the environment, thereby causing serious pollution. Biological treatment processes offer a way to reduce the amounts of xenobiotic compounds in the environment. Bioremediation (treatment of waste with microorganisms) (49) and phytoremediation (treatment of polluted soil with plants) (17) are regarded as cost-efficient technologies for the clean-up of polluted waters or soils. Many wastewater treatment plants rely on biofilms. One of the advantages of biofilm reactors is their compactness. The small footprint areas of biofilm reactors are due to two factors, high volumetric biomass concentrations (29) and high retention times (6). The extended retention time of bacteria growing near the substratum is the reason why a greater variety of microorganisms can develop compared to the variety in planktonic cultures (8). Another reason is the formation of microniches due to concentration gradients of nutrients and electron acceptors. Chemical pollutants can adsorb or be degraded further, contributing to the zonation on a microscale. Hence, extended retention times of solids (6), species diversity, higher local nutrient concentrations (14), and high retention times for recalcitrant compounds due to adsorption in the extracellular polymeric matrix (48) in biofilm reactors facilitate remediation of synthetic pollutants. Furthermore, biofilms allow interspecies interactions by signaling and nutrient cycling (20) and offer resistance to toxicant (30), physical (13), or environmental stresses (44).
However, for a system to be able to degrade xenobiotic compounds, the biocatalysts that are needed for degradation of a specific compound need to be present in the system (22). If reactors lack the desired biocatalysts, they need to be bioaugmented to obtain efficient degradation. By addition of exogenous optimally constructed bacteria (11, 43) or functionally adapted bacterial consortia (18) and communities (47), enrichment of the total gene pool can be obtained. Likewise, a reactor can be augmented by in situ gene transfer (42). It is surprising that in spite of several successes, few studies have evaluated bioaugmentation by horizontal gene transfer in activated sludge wastewater treatment reactors (33, 34, 50) or activated sludge microcosms (16, 36, 41). When plasmids carrying catabolic genes were integrated into indigenous bacterial organisms, increased and more rapid degradation of the target compound was observed (33, 34, 36). Likewise, there is little information available regarding bioaugmentation of biofilm reactors by in situ gene transfer by bacterial conjugation (1, 4, 12, 15, 53). Conjugation inside a biofilm matrix offers a great advantage in terms of both gene transfer frequency (1, 15, 19) and subsequent transconjugant stability (15). Still, the effect of in situ natural genetic transformation in order to obtain bioenhancement in activated sludge- or biofilm-based biological process engineering systems has not been studied.
To investigate the feasibility of bioaugmentation by genetic transformation, in situ natural genetic transformation was investigated with a model system consisting of biofilm-cultured Acinetobacter sp. strain BD413 (24) with the autonomously replicating gfp-carrying plasmid pGAR1 (19) as the transforming DNA. Acinetobacter species are ubiquitous, strictly aerobic, nonmotile organisms that can be isolated from soil, water, or wastewater (23). Acinetobacter strains can degrade recalcitrant aromatic and alicyclic compounds, as well as some aromatic amino acids, mineral oils, and synthetic polymers (3, 5, 7, 40). In addition, Acinetobacter strains produce biosurfactants, like emulsan (25, 45) and alasan (2), that enhance the bioavailability of poorly soluble compounds. Furthermore, Acinetobacter sp. strain BD413 (24) is amenable to gene manipulation by conjugation, transformation, and transduction (23), and this property makes the strain particularly interesting as a tool for biologically enhancing the catabolic properties of hazardous waste treatment facilities.
In this study the following conditions were investigated to determine their effects on natural genetic transformation in a model biofilm: biofilm age, free DNA concentration, and growth of the biofilm in the presence or absence of cells in the bulk fluid.
MATERIALS AND METHODS
Bacteria, plasmids, and media.
Acinetobacter sp. strain BD413 (24) was used as the model strain for evaluation of natural genetic transformation in monoculture biofilms. pGAR1 was isolated from Escherichia coli strain GM16 (19). pGAR1 is pRK415 (26), a Mob+ Tra− Tetr Inc P1 plasmid, carrying the wild-type gfp (green fluorescent protein [GFP]) gene (Clontech, Palo Alto, Calif.) under regulation of a Plac promoter. Induction of the promoter by addition of isopropyl-β-d-thiogalactopyranoside was not necessary. The fluorescence intensity of single BD413(pGAR1) cells could be clearly distinguished from the fluorescence intensity of unlabeled single BD413 cells by epifluorescence and confocal laser scanning microscopy (CLSM). Rich Luria-Bertani medium (38) and minimal medium M9 (0.2% gluconate) (46) were used during transformation experiments. Standard microbiological techniques (spectrophotometric DNA concentration measurement, bacterial cultivation techniques, DNA extraction and purification) were performed as described by Sambrook et al. (46).
Transformation in biofilms.
Transformation of biofilms was performed by the methods described by Wuertz et al. (54). Biofilms of Acinetobacter sp. strain BD413 were grown in a flow cell containing four separate flow channels (4 by 4 by 40 mm) for 3 days in rich Luria-Bertani medium by using a flow rate of 2.4 ml/h after DNA was added with minimal medium M9 (0.2% gluconate). Transformation involved incubation of DNA during 1 h of continuous flow with DNA-containing medium, unless indicated otherwise. After overnight incubation in minimal medium M9 (0.2% gluconate) without DNA at a flow rate of 2.4 ml/h, which allowed expression of the received gene, biofilms were prepared for microscopic monitoring.
Microscopic monitoring, image acquisition, and data processing.
Biofilm cells in Acinetobacter sp. strain BD413 monoculture biofilms undergoing transformation were visualized with the general nucleic acid stain Syto 60 (Molecular Probes, Eugene, Oreg.) and were detected with an LSM 410 CLSM (Zeiss, Jena, Germany). The 633-nm laser line and a 665-nm long-pass emission filter were used to detect cells stained with Syto 60. The 488-nm laser line and a 515-nm long-pass emission filter were used for detection of cells expressing gfp (Clontech). Automated image acquisition and data processing were performed by the methods described by Wuertz et al. (54).
Colocalization experiments.
Pure-culture BD413(pGAR1) biofilms grown for 46 h in selective minimal medium (minimal medium M9 containing 0.2% gluconate and 20 μg of tetracycline per ml) were stained with Syto 60 to test colocalization of the two signals. Approximately 18.5% of the signals in the BD413(pGAR1) biofilm were colocalized GFP and Syto 60 signals, 22.6% were single Syto 60 signals, and 58.9% were single GFP signals. Therefore, in transformation experiments the total cell volume was obtained by adding the Syto 60 signals and the GFP signals and subtracting the overlapping signals. GFP signals were considered transformants. Potential underestimation of the 22.6% GFP signals was not considered during calculation of the results because it was not possible to check whether cells still contained the introduced plasmid. When no DNA was added to the medium, no signals were detectable with CLSM settings for detection of GFP signals in two separate tests. In an additional test, signals were obtained by using the 515-nm long-pass emission filter, but no overlap was detectable with the Syto 60 settings. These false-positive signals might have been due to autofluorescent impurities in the inlet medium and could be clearly distinguished from fluorescence-expressing cells on the basis of form and size. Hence, to avoid overestimation of GFP due to autofluorescent impurities, images had to be checked manually to discard possible false-positive signals on the basis of form and size.
Mathematical parameters.
The mathematical equations describing volumes and transformation frequency are given below.
Calculation of the volume of transformants and the volume of recipients was based on the following equation:
![]() |
where VX,e is total volume of transformants or recipients (in cubic micrometers), Xi is the area covered by cells of interest at position i (in square micrometers), zi is the distance from the substratum at position i (in micrometers), i is the scanning position in the z direction starting at the biofilm substratum, and e is the last scanned position in a biofilm in the z direction.
The equation was adapted as follows with the trapezoidal rule to obtain a more correct approximation of the numerical integral as described by Kuehn et al. (28):
![]() |
The transformation frequency (TF) was the fraction of transformant volume per total cell volume: TF = VT,e/VR,e, where VT,e is the volume of transformants obtained with equation 2 (in cubic micrometers) and VR,e is the volume of recipients obtained with equation 2 (in cubic micrometers).
Below, transfer frequencies are expressed as the volume of transgenic cells per volume of recipient cells, unless indicated otherwise.
To obtain a reproducible estimate of the normalized distance from the substratum, we limited the total cell volume and biofilm thickness to contain 98% of the scanned biomass (VR,e) starting from the substratum towards the biofilm-bulk fluid interface within the biofilm volume investigated. The normalized distance from the substratum was calculated by dividing the distance from the biofilm attachment surface by the biofilm thickness: di = zi/zk for VR,k ≥ VR,e × 0.98, where VR,k is the volume of recipients calculated with equation 2 limited to 98% of the total scanned biomass (in cubic micrometers), di is the normalized distance from the substratum at position i (in micrometers), zi is the distance from the substratum at position i (in micrometers), zk is the biofilm thickness or distance from the substratum at position k (in micrometers), and k is the position in a biofilm where VR,k reached 98% of VR,e.
Reproducibility of in situ natural genetic transformation in monoculture Acinetobacter sp. strain BD413 biofilms.
Biofilms should be observed noninvasively and with confidence that reliable and reproducible results will be obtained. Direct in situ detection of large areas is, therefore, desirable. A study to determine statistically representative areas of Pseudomonas fluorescens biofilms showed that a minimal area of 1 × 105 μm2 should be scanned to obtain reproducible results in biofilm investigations (27). In the present study a minimal area of 2.4 × 105 μm2 was scanned to monitor a biofilm volume of at least 1.2 × 107 μm3. The standard transformation frequencies with 0.2 μg of pGAR1/ml, derived from four separate experiments monitoring a minimum volume of 1.2 × 107 μm3, ranged from logarithmic values of −3.5 to −3.1, with a mean value of −3.3 and a standard deviation of 0.15. In contrast to the standard method in which planktonic cells are used (38), the in situ method in which CLSM was used provided reliable and reproducible results.
RESULTS
Effect of biofilm age on natural genetic transformation.
To test the transformability of biofilm cells at different growth stages, experiments were conducted with 0.2 μg of pGAR1 DNA/ml (Table 1). The responses of cells to DNA exposure were measured with two different exposure times. Flow cell experiments were performed with 1- and 3-day-old biofilms. The thickness of Acinetobacter sp. strain BD413 biofilms grown in rich medium reached a maximum after 2 days for four separately investigated biofilms, and after this the biofilm thickness remained at a steady state due to lysis, detachment, and subsequent growth, indicating a mature biofilm stage (results not shown). A 1-day-old biofilm could therefore be considered an actively growing biofilm, and a 3-day-old biofilm could be considered a mature biofilm. Although cells in young and growing biofilms are more readily transformed, the biofilm heterogeneity in 3-day-old biofilms still ensured the presence of a significant fraction of competent cells that were able to take up DNA (Table 1; Fig. 1). Furthermore, cells responded within 15 min to the addition of exogenous DNA.
TABLE 1.
Transformation frequencies obtained for biofilms of different ages exposed to 0.2 μg of plasmid pGAR1 DNA/ml for different periods of timea
Time of exposure to DNA (min) | Transformation frequency
|
|
---|---|---|
1-day-old biofilm | 3-day-old biofilm | |
15 | 1.9 × 10−2 | 7.7 × 10−4 |
45 | 1.3 × 10−2 | 7.4 × 10−4 |
Data were obtained by quantitative microscopy.
FIG. 1.
Microscopic images showing pGAR1 transformants in 3-day-old (A) and 1-day-old (B) Acinetobacter sp. strain BD413 biofilms. The black and white images show GFP signals (panels I) or Syto 60-stained cells (panels II). Superimposed single optical images (panels III) show gfp transformants (yellow, green) against a background of recipient Acinetobacter sp. strain BD413 biofilm cells (red). The edges of each image correspond to a length of 90 μm.
Effect of concentration of added free DNA.
To investigate the influence of DNA concentration on transformation frequency and transformant location in a mature biofilm of the highly competent organism Acinetobacter sp. strain BD413, natural genetic transformation was investigated in biofilms that were exposed for 1 h to a specific pGAR1 DNA load.
The DNA concentration in the inlet medium ranged from 1 × 10−9 to 1.5 μg of pGAR1/ml. The transformation frequency increased as a function of DNA concentration within the range of DNA concentrations investigated (Fig. 2).
FIG. 2.
Transformation in monoculture strain BD413 biofilms with pGAR1 DNA at various concentrations in the inlet medium expressed as a semilog plot. The insert shows the same data points on a log-log scale.
The observed relationship between DNA concentration and transformation frequency corresponds to the results of previous studies involving batch transformation experiments (38). However, a saturation point was not reached in this study. Further examination with DNA concentrations of 100 μg/ml or more would be needed to see if a saturation point is reached or if transformation keeps increasing until 100% of the cells are transformed. Such high concentrations were not investigated in this study as they are unlikely to occur in nature or are not very practical for bioaugmentation of bioreactors.
With low concentrations of pGAR1 in the feed, transformants were formed at the biofilm attachment surface. Exposure to increasing amounts of pGAR1 resulted in gradual accumulation of transformants at the bottom of the biofilm (Fig. 3), where the biofilm density was the greatest, and not in the middle or upper part of the biofilm. If it were true that the fraction of competent cells is homogeneously distributed inside a biofilm, most transformants would be found in the layers with the highest biofilm density and the transformation frequency would be equally high throughout the biofilm. However, this was observed only 5 of 10 times. Hence, other factors may have contributed to the distribution of frequencies detected.
FIG. 3.
Distribution profiles for the volume of gfp transformants as a function of the normalized distance from the substratum when organisms were exposed to 1 × 10−9 μg of DNA/ml (•), 1 × 10−7 μg of DNA/ml (□), 1 × 10−4 μg of DNA/ml (⧫), 1 × 10−1 μg of DNA/ml (▵), and 1.5 μg of DNA/ml (∗).
Effect of biofilm ontogenesis on natural genetic transformation.
Biofilms can grow either in a batch mode (on the surface of a shaken container containing a bacterial suspension) or in a continuous mode (by constant feeding of a biofilm grown on a surface with fresh medium in a flow channel). A biofilm grown in a shaken container emerges with suspended cells present in the bulk fluid, while a biofilm in a flow channel grows in an environment continuously rinsed with suspended cells in the medium. All previous experiments were performed with biofilms grown in continuously fed flow cells. How would a biofilm that had developed in the presence of stationary-phase bacterial cells in the bulk phase respond to the addition of free DNA? To answer this question, flow cells were continuously fed with medium containing stationary-phase Acinetobacter sp. strain BD413 cells to mimic biofilm ontogenesis in the presence of a batch-grown suspended culture. After this, the biofilms were rinsed with a 0.01 M MgSO4 to remove any suspended cells from the flow channel. Cell-free minimal medium containing 0.1 μg of pGAR1 DNA/ml was added to the flow cells for 1 h. The preparations were incubated overnight in cell-free minimal medium, and after this the biofilms were stained, washed, microscopically investigated, and quantified. No transformation was detected in the biofilms, which had grown in the presence of cells in the bulk fluid.
Next, we investigated the effect of prestarvation on transformation of biofilm cells grown in the presence of cells in the bulk phase. Suspended cells were removed as described above by rinsing with 0.01 M MgSO4. The biofilm was then starved by feeding the flow cell either with a continuous supply of minimal medium M9 without a carbon source or with a salt solution (0.01 M MgSO4) for 24 h. Subsequent transformation was done with identical experimental transformation steps, as described above. Transformation was observed at low but measurable frequencies (9.1 × 10−5 and 1.9 × 10−5 for biofilms starved in the presence of minimal medium M9 and in the presence of the salt solution, respectively).
When biofilms that were grown with cells present in the inlet medium were rinsed with 0.01 M MgSO4 and subsequently fed with cell-free rich medium for 24 h before transformation, again no transformation was observed in the biofilms.
DISCUSSION
Standard transformation experiments in which Acinetobacter sp. strain BD413 is used require cells to be transformed in a state of competency (38). Acinetobacter sp. strain BD413 reaches competence at the early log phase. The cells remain competent during the log phase, and competence drops to almost zero when the stationary phase is reached (31). In standard batch culture experiments (31) or soil microcosms (35), additional transformation events were not detected after a prolonged period that was more than 12 h long. Similar to exponentially growing suspended cultures, young and growing biofilms allowed a high frequency of transformation events. In addition, the biofilms still contained significant fractions of competent cells for at least 3 days after inoculation. The biofilm mode of growth could therefore be compared to the continuously exponentially growing steady-state batch or turbidostat cultures of Palmen et al. (37). These authors studied Acinetobacter sp. strain BD413 cultured in a continuous mode in order to prolong the period of competency. After 3 days they were still able to observe a detectable transformation frequency, but it was less than the initial transformation frequencies (37); this was impossible in transformation experiments performed with Acinetobacter sp. strain BD413 cultured in a batch mode (31, 35).
The DNA concentration in the inlet was positively correlated with increasing transformation frequency. Natural genetic transformation occurred readily at high frequencies in monoculture Acinetobacter sp. strain BD413 biofilms. When 1.2 μg of pGAR1/ml was used, transformation frequencies as high as 2.4 × 10−3 were observed. Furthermore, it was possible to obtain detectable transformation frequencies with minute amounts (as little as 1 fg of pGAR1/ml) in biofilms. The lowest tested DNA concentration used for transformation of Acinetobacter sp. strain BD413 reported previously was approximately 1 ng of DNA/ml (38). Transfer frequencies even higher than those reported here may be obtainable if a feed with a DNA concentration greater than 10 μg of pGAR1/ml is added to a biofilm. For bioaugmentation via gene transfer, however, it is not necessary to have maximum transformation frequencies. Even if transfer frequencies are low, transformants may undergo cell division, and the transgenic strain could therefore still establish itself in a reactor system. In such a case, low but significant transfer frequencies should be enough to obtain successful bioaugmentation.
It was interesting that an increased DNA concentration in the influent resulted in accumulation of transformants in the biofilm substratum. However, it seems odd that most transformation events take place at the biofilm base. This observation implies that free DNA first has to diffuse through the biofilm before competent cells take up and subsequently replicate the integrated plasmid. It is possible that immobilization of cells was responsible for the occurrence of most transformation events at the bottom of the biofilm. The location of transformation events seemed to be simply a matter of chance, and the probability increased with increasing cell density at a certain distance from the substratum. In the case of monoculture Acinetobacter sp. strain BD413 biofilms the cell density was greatest near the biofilm attachment surface. Therefore, the chance for transformation was greater near the biofilm substratum. This suggests that the chances for transformation to occur are highest in a tightly packed biofilm (a biofilm with a low porosity value). However, it is not correct that porous biofilms are ill suited for transformation. Young biofilms, for example, are very porous (Fig. 1), and they allowed transformation at increased rates due to enhanced competency levels during exponential growth. Porosity plays an important role only in mature biofilms. Therefore, it is necessary to investigate the true impact of biofilm density and porosity on natural genetic transformation. If density played an important part in natural genetic transformation, anthropogenic manipulation of the factors that decrease biofilm porosity could lead to enhanced occurrence of transformation events. van Loosdrecht et al. (51) included flow rate, nutrient loading rate, and growth rate of the biofilm cells as parameters that influence biofilm thickness and porosity. In addition, cell-to-cell signaling molecules influence biofilm structure (10) and may indirectly influence transformation events.
Another important parameter in this study was the mode of growth in which the biofilm itself emerged. In many biofilm investigations scientists use batch-grown biofilms in 96-well microtiter plates (9, 39). In contrast to batch-grown biofilms, biofilms grown in flow chambers are in continuous contact with fresh medium that does not contain suspended cells. One needs to consider that biofilms grown in different modes could possess different qualities. In the present study, differences in the transformability of Acinetobacter sp. strain BD413 biofilm cells grown in the presence and in the absence of cells in the bulk fluid were observed. The presence of high numbers of cells in the surrounding medium inhibited transformation in biofilms (Hendrickx, results not shown). Likewise, when biofilms developed in the presence of high numbers of cells in the bulk fluid but were rinsed to eliminate cells present in the surrounding medium before they were exposed to naked DNA, transformation was strongly inhibited. Also, differences in the morphotypes of the bacterial cells were observed. Acinetobacter sp. displays two different morphotypes, the bacillar morphotype and the coccoid morphotype (21). In biofilms fed with cell-free medium, both the bacillar and coccoid forms were detected. In contrast, when biofilms were allowed to emerge in the presence of medium containing suspended bacterial cells, the cells exhibited almost exclusively the bacillar morphotype.
As determined by microscopy, batch-cultured stationary-phase Acinetobacter sp. strain BD413 cells also displayed the bacillar morphotype when the organism was grown in rich medium. Hence, it is possible that the lack of transformability in biofilms grown in the presence of cells in the bulk fluid is due to increased attachment of bacterial cells that have entered the stationary phase (and hence have become noncompetent) in the surrounding medium.
Biofilms grown in biofilm reactors may encounter many passing free-floating bacteria. This would discourage bioaugmentation by in situ transformation if the problem of inhibited gene transfer due to the presence of suspended cells during biofilm ontogenesis were unsolvable. Rinsing a biofilm to remove most cells present in the drifting fraction, followed by a starvation period, could reinduce in situ transformation when the cells are exposed to a nutrient-containing substrate with plasmid DNA. James and coworkers (21) discovered that upon starvation, bacillar cells revert to the coccoid form by reduction division, resulting in conservation of biomass but increased cell number. In this study the CLSM investigation revealed that the biofilms treated with starvation-inducing medium contained many cells with the coccoid morphotype, as observed when biofilms were grown in the absence of cells in the bulk fluid. Thus, all signs suggest that coccoid cells are the competent cells and bacillar cells are not competent for DNA uptake. Still, when cell shape was checked as a function of distance from the substratum, it was observed that cells at the biofilm substratum had the bacillar shape, while the prominent morphotype of cells at the biofilm-medium interface was coccoid (Hendrickx, data not shown). However, most transformants were formed at the biofilm substratum, where most bacillus-shaped cells resided. Further experiments are therefore needed to establish if other parameters related to the amount or location of bacillus-shaped or coccoid cells can be correlated with transformation frequency.
Nevertheless, the observed differences in transformation frequency in biofilms grown in the presence and in the absence of planktonic cells show that care should be taken in designing biofilm experiments. In addition to the effects of standard environmental parameters (temperature, pH, nutrient content, substratum, moisture content, biotic and abiotic stresses, flow rate, etc.), the results might differ considerably depending on whether the biofilms are grown under batch or continuous-flow conditions.
It should be noted that the experiments which we performed were not designed to prove the general feasibility of in situ biofilm cell transformation with any highly competent soil bacterium in naturally occurring biofilms. Our results do indicate that transformation has potential as a tool for bioaugmentation of biofilm reactors. It is interesting that transformation with Acinetobacter sp. strain BD413 resulted in transformation frequencies in nonsterile groundwater and wet soil microcosms that were as high as those obtained under sterile conditions (32). Likewise, transformation of Acinetobacter sp. strain BD413 biofilm cells embedded in river epilithon was not inhibited by the presence of indigenous organisms (52). It can be speculated that the presence of an ambient community in wastewater treatment systems should have few negative effects on transformation of Acinetobacter sp. strain BD413 cells. Future research should elucidate the efficiency of transforming cells which reside in natural biofilms inside bioreactors and other systems.
Acknowledgments
This work was supported by the Research Center for Fundamental Studies of Aerobic Biological Wastewater Treatment, Munich, Germany (grant SFB411), and by the University of California, Davis.
REFERENCES
- 1.Angles, M. L., K. C. Marshall, and A. E. Goodman. 1993. Plasmid transfer between marine bacteria in the aqueous phase and biofilms in reactor microcosms. Appl. Environ. Microbiol. 59:843-850. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2.Barkay, T., S. Navon-Venezia, E. Z. Ron, and E. Rosenberg. 1999. Enhancement of solubilization and biodegradation of polyaromatic hydrocarbons by the bioemulsifier alasan. Appl. Environ. Microbiol. 65:2697-2702. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 3.Bauman, P., M. Doudoroff, and R. Y. Stanier. 1968. A study of the Moraxella group. II. Oxidative-negative species (genus Acinetobacter). J. Bacteriol. 95:1520-1541. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4.Beaudoin, D. L., J. D. Bryers, A. B. Cunningham, and S. W. Peretti. 1998. Mobilization of broad host range plasmid from Pseudomonas putida to established biofilm of Bacillus azotoformans. I. Experiments. Biotechnol. Bioeng. 57:272-279. [PubMed] [Google Scholar]
- 5.Benndorf, D., N. Loffhagen, and W. Babel. 2001. Protein synthesis patterns in Acinetobacter calcoaceticus induced by phenol and catechol shows specificities of responses to chemostress. FEMS Microbiol. Lett. 200:247-252. [DOI] [PubMed] [Google Scholar]
- 6.Bishop, P. L. 1997. Biofilm structure and kinetics. Water Sci. Technol. 36:287-294. [Google Scholar]
- 7.Bode, H. B., K. Kerkhoff, and D. Jendrossek. 2001. Bacterial degradation of natural and synthetic rubber. Biomacromolecules 2:295-303. [DOI] [PubMed] [Google Scholar]
- 8.Christensen, B. B., C. Sternberg, J. B. Andersen, L. Eberl, S. Moller, M. Givskov, and S. Molin. 1998. Establishment of new genetic traits in a microbial biofilm community. Appl. Environ. Microbiol. 64:2247-2255. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9.Danese, P. N., L. A. Pratt, and R. Kolter. 2001. Biofilm formation as a developmental process. Methods Enzymol. 336:19-26. [DOI] [PubMed] [Google Scholar]
- 10.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. [DOI] [PubMed] [Google Scholar]
- 11.Duba, A. G., K. J. Jackson, M. C. Jovanovich, R. B. Knapp, and R. T. Taylor. 1996. TCE remediation using in situ, resting-state bioaugmentation. Environ. Sci. Technol. 30:1982-1989. [Google Scholar]
- 12.Ehlers, L. J., and E. J. Bouwer. 1999. RP4 plasmid transfer among species of Pseudomonas in a biofilm reactor. Water Sci. Technol. 39:163-171. [Google Scholar]
- 13.Flemming, H.-C. 1994. Biofilme, Biofouling und mikrobielle Schädigung von Werkstoffen. Forschungs- und Entwicklungsinstitut für Industrie- und Siedlungswasserwirtschaft sowie Abfallwirtschaft e. V., Universität Stuttgart, Habilitationsschrift, Stuttgart, Germany.
- 14.Flemming, H.-C. 1995. Sorption sites in biofilms. Water Sci. Technol. 32:27-33. [Google Scholar]
- 15.Frank, N., A. M. Simao Beaunoir, M. A. Dollard, and P. Bauda. 1996. Recombinant plasmid DNA mobilization by activated sludge strains grown in fixed-bed or sequenced-batch reactors. FEMS Microbiol. Ecol. 21:139-148. [Google Scholar]
- 16.Geisenberger, O., A. Ammendola, B. B. Christensen, S. Molin, K.-H. Schleifer, and L. Eberl. 1999. Monitoring the conjugal transfer of plasmid RP4 in activated sludge and in situ identification of the transconjugants. FEMS Microbiol. Lett. 174:9-17. [DOI] [PubMed] [Google Scholar]
- 17.Gianfreda, L., and P. Nannipieri. 2001. Basic principles, agents and feasibility of bioremediation of soil polluted by organic compounds. Minerva Biotechnol. 13:5-12. [Google Scholar]
- 18.Hajji, K. T., F. Lepine, J. G. Bisaillon, R. Beaudet, J. Hawari, and S. R. Guiot. 2000. Effects of bioaugmentation strategies in UASB reactors with a methanogenic consortium for removal of phenolic compounds. Biotechnol. Bioeng. 67:417-423. [DOI] [PubMed] [Google Scholar]
- 19.Hausner, M., and S. Wuertz. 1999. High rates of conjugation in bacterial biofilms as determined by quantitative in situ analysis. Appl. Environ. Microbiol. 65:3710-3713. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20.James, G. A., L. Beaudette, and J. W. Costerton. 1995. Interspecies bacterial interactions in biofilms. J. Ind. Microbiol. 15:257-262. [Google Scholar]
- 21.James, G. A., D. R. Korber, D. E. Caldwell, and J. W. Costerton. 1995. Digital image analysis of growth and starvation responses of a surface-colonizing Acinetobacter sp. J. Bacteriol. 177:907-915. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 22.Janssen, D. B., and J. P. Schanstra. 1994. Engineering proteins for environmental applications. Curr. Opin. Biotechnol. 5:253-259. [DOI] [PubMed] [Google Scholar]
- 23.Juni, E. 1978. Genetics and physiology of Acinetobacter. Annu. Rev. Microbiol. 32:349-371. [DOI] [PubMed] [Google Scholar]
- 24.Juni, E., and A. Janik. 1969. Transformation of Acinetobacter calcoaceticus (Bacterium anitratum). J. Bacteriol. 98:281-288. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 25.Karanth, N. G. T., P. G. Deo, and N. K. Veenanadig. 1999. Microbial production of biosurfactants and their importance. Curr. Sci. 77:116-126. [Google Scholar]
- 26.Keen, N. T., S. Tamaki, D. Kobayashi, and D. Trollinger. 1988. Improved broad-host-range plasmids for DNA cloning in gram-negative bacteria. Gene 70:191-197. [DOI] [PubMed] [Google Scholar]
- 27.Korber, D. R., J. R. Lawrence, M. J. Hendry, and D. E. Caldwell. 1992. Programs for determining statistically representative areas of microbial biofilms. Binary 4:204-210. [Google Scholar]
- 28.Kuehn, M., M. Hausner, H.-J. Bungartz, M. Wagner, P. A. Wilderer, and S. Wuertz. 1998. Automated confocal laser scanning microscopy and semiautomated image processing for analysis of biofilms. Appl. Environ. Microbiol. 64:4115-4127. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 29.Kwok, W. K., C. Picioreanu, S. L. Ong, M. C. M. van Loosdrecht, W. J. Ng, and J. J. Heijnen. 1998. Influence of biomass production and detachment forces on biofilm structures in a biofilm airlift suspension reactor. Biotechnol. Bioeng. 58:400-407. [DOI] [PubMed] [Google Scholar]
- 30.Lewis, K. 2001. Riddle of biofilm resistance. Antimicrob. Agents Chemother. 45:999-1007. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 31.Lorenz, M. G., D. Gerjets, and W. Wackernagel. 1991. Release of transforming plasmid and chromosomal DNA from two cultured soil bacteria. Arch. Microbiol. 156:319-326. [DOI] [PubMed] [Google Scholar]
- 32.Lorenz, M. G., K. Reipschläger, and W. Wackernagel. 1992. Plasmid transformation of naturally competent Acinetobacter calcoaceticus in non-sterile soil extract and groundwater. Arch. Microbiol. 157:355-360. [DOI] [PubMed] [Google Scholar]
- 33.McClure, N. C., A. J. Weightman, and J. C. Fry. 1989. Survival of Pseudomonas putida UWC1 containing cloned catabolic genes in a model activated sludge unit. Appl. Environ. Microbiol. 55:2627-2634. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 34.McClure, N. C., J. C. Fry, and A. J. Weightman. 1991. Survival and catabolic activity of natural and genetically engineered bacteria in a laboratory-scale activated sludge unit. Appl. Environ. Microbiol. 57:366-373. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 35.Nielsen, K. M., A. M. Bones, and J. D. van Elsas. 1997. Induced natural transformation of Acinetobacter calcoaceticus in soil microcosms. Appl. Environ. Microbiol. 63:3972-3977. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 36.Nüßlein, K., D. Maris, K. Timmis, and D. F. Dwyer. 1992. Expression and transfer of engineered catabolic pathways harbored by Pseudomonas spp. introduced into activated sludge microcosms. Appl. Environ. Microbiol. 58:3380-3386. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 37.Palmen, R., P. Buijsman, and K. J. Hellingwerf. 1994. Physiological regulation of competence induction for natural transformation in Acinetobacter calcoaceticus. Arch. Microbiol. 162:344-351. [Google Scholar]
- 38.Palmen, R., B. Vosman, P. Buijsman, C. K. D. Breek, and K. J. Hellingwerf. 1993. Physiological characterization of natural transformation in Acinetobacter calcoaceticus. J. Gen. Microbiol. 139:295-305. [DOI] [PubMed] [Google Scholar]
- 39.Parkins, M. D., M. Altebaeumer, H. Ceri, and D. G. Storey. 2001. Subtractive hybridization-based identification of genes uniquely expressed or hyperexpressed during biofilm growth. Methods Enzymol. 336:76-84. [DOI] [PubMed] [Google Scholar]
- 40.Pleshakova, E. V., A. Y. Muratova, and O. V. Turkovskaya. 2001. Degradation of mineral oil with a strain of Acinetobacter calcoaceticus. Appl. Biochem. Microbiol. 37:342-347. [PubMed] [Google Scholar]
- 41.Ravatn, R., A. J. Zehnder, and J. R. van der Meer. 1998. Low-frequency horizontal transfer of an element containing the chlorocatechol degradation genes from Pseudomonas sp. strain B13 to Pseudomonas putida F1 and to indigenous bacteria in laboratory-scale activated-sludge microcosms. Appl. Environ. Microbiol. 64:2126-2132. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 42.Rittmann, B. E., B. F. Smets, and D. A. Stahl. 1990. The role of genes in biological processes. Environ. Sci. Technol. 24:23-29. [Google Scholar]
- 43.Roanne, T. M., K. L. Josephson, and I. L. Pepper. 2001. Dual-bioaugmentation strategy to enhance remediation of cocontaminated soil. Appl. Environ. Microbiol. 67:3208-3215. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 44.Roberson, E. B., and M. K. Firestone. 1992. Relationship between desiccation and exopolysaccharide production in a soil Pseudomonas sp. Appl. Environ. Microbiol. 58:1284-1291. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 45.Rouse, J. D., D. A. Sabatini, J. M. Sulfita, and J. H. Harwell. 1994. Influence of surfactants on microbial degradation of organic compounds. Crit. Rev. Environ. Sci. Technol. 24:325-370. [Google Scholar]
- 46.Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed., Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
- 47.Saravanane, R., D. V. Murthy, and K. Krishnaiah. 2001. Treatment of anti-osmotic drug based pharmaceutical effluent in an upflow anaerobic fluidized bed system. Waste Manag. 21:563-568. [DOI] [PubMed] [Google Scholar]
- 48.Späth, R., H.-C. Flemming, and S. Wuertz. 1998. Sorption properties of biofilms Water Sci. Technol. 37:207-210. [Google Scholar]
- 49.Timmis, K. N., and D. H. Pieper. 1999. Bacteria designed for bioremediation. Trends Biotechnol. 17:200-204. [DOI] [PubMed] [Google Scholar]
- 50.Van Limbergen, H., E. M. Top, and W. Verstraete. 1998. Bioaugmentation in active sludge: current features and future perspectives. Appl. Microbiol. Biotechnol. 50:16-23. [Google Scholar]
- 51.van Loosdrecht, M. C. M., C. Picioreanu, and J. J. Heijnen. 1997. A more unifying hypothesis for biofilm structures. FEMS Microbiol. Ecol. 24:181-183. [Google Scholar]
- 52.Williams, H. G., M. J. Day, J. C. Fry, and G. J. Stewart. 1996. Natural transformation in river epilithon. Appl. Environ. Microbiol. 62:2994-2998. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 53.Wuertz, S. 2002. Gene exchange in biofilms, p. 1408-1420. In G. Bitton (ed.), Encyclopedia of environmental microbiology, vol. 3. John Wiley and Sons, New York, N.Y.
- 54.Wuertz, S., L. Hendrickx, M. Kuehn, K. Rodenacker, and M. Hausner. 2001. In situ quantification of gene transfer in biofilms. Methods Enzymol. 336:129-143. [DOI] [PubMed] [Google Scholar]