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a Dep. of Plant and Soil Sciences, Univ. of Delaware, Newark, DE 19717
b Dep. of Civil and Environmental Engineering, Univ. of Delaware, Newark, DE 19717
* Corresponding author (mrad{at}udel.edu)
Received for publication June 26, 2000.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONREFERENCES |
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Abbreviations: ACN, acetonitrile • CMC, critical micelle concentration • HPLC, high performance liquid chromatography • NAPL, nonaqueous-phase liquid • PAH, polycyclic aromatic hydrocarbon • PBS, phosphate-buffered saline • PHE, phenanthrene • PPGAS, proteose peptone glucose ammonium salts
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONREFERENCES |
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Mass transfer from sorbed or insoluble phases is often considered to be the rate-limiting step in biodegradation of organic contaminants because the compounds must be released to the aqueous phase prior to entering the microbial cell and subsequent intracellular transformation by the necessary catabolic enzymes (Churchill et al., 1995; Zhang and Miller, 1994). Therefore, the bioavailable portion of hydrophobic contaminants is generally defined as the fraction that is soluble and/or readily released to the bulk aqueous phase (Cerniglia, 1992; Herman et al., 1997). However, evidence has emerged that is inconsistent with the assumption that suggests that some bacteria can degrade sorbed substrates at rates that exceed those that would be predicted by bulk aqueous contaminant concentrations (Crocker et al., 1995; Guerin and Boyd, 1992; Harms and Zehnder, 1995; Ortega-Calvo and Alexander, 1994; Osswald et al., 1996). These studies propose that while some microorganisms are limited to aqueous-phase substrates, others may be better adapted for accessing contaminants that are sorbed to soil or partitioned into a nonaqueous-phase liquid (NAPL). Ortega-Calvo and Alexander (1994) observed that an Arthrobacter sp. degraded NAPL-associated naphthalene at a rate higher than the measured rate of partitioning for the two-phase system. They concluded that there were two physiologically distinct subpopulations of this strain present in the system, one able to degrade naphthalene in solution, and the other able to degrade naphthalene at the NAPL–water interface (Ortega-Calvo and Alexander, 1994). Experiments with soil-sorbed naphthalene showed similar results (Guerin and Boyd, 1992). In these experiments, differential bioavailability of naphthalene to two naphthalene-degrading isolates was observed. In kinetic studies, P. putida strain 17484 was shown to have direct access to sorbed naphthalene whereas strain NP-Alk was limited to aqueous-phase naphthalene (Guerin and Boyd, 1992). Crocker et al. (1995) verified these results while studying the bioavailability of naphthalene with the same two bacterial strains in surfactant-modified clay–water systems.
Differential mineralization of NAPL styrene and sorbed 3-chlorodibenzofuran has also been observed, but the mechanism responsible for the enhanced bioavailability of these compounds was less clear (Harms and Zehnder, 1995; Osswald et al., 1996). These researchers postulated that bioavailability was a function of cell attachment, steep concentration gradients that occur near interfaces, or cell-mediated solubilization of the contaminant via biosurfactant production (Harms and Zehnder, 1995; Osswald et al., 1996). In any case, it is clear from these studies that the bioavailability of a contaminant is species-specific, and in many instances, not strictly limited to bulk aqueous-phase substrates.
Biosurfactants can enhance hydrocarbon desorption and solubility, but their effect on bioavailability and degradation are less straightforward. The addition of biosurfactant to soil–water systems has been shown to have conflicting effects on contaminant mineralization by different bacteria (Allen et al., 1999; Stelmack et al., 1999). The presence of surfactants at a concentration equal to one-half the critical micelle concentration (CMC) inhibited the uptake and biodegradation of NAPL extracts from creosote-contaminated soils, presumably due to reduced bacterial adhesion to the NAPL–water interface (Stelmack et al., 1999). Another study by Allen et al. (1999) showed that the surfactant Triton X-100 had contrasting effects on PAH degradation by two PAH-degrading microorganisms. The presence of the surfactant enhanced mineralization of both naphthalene and phenanthrene (PHE) by Pseudomonas strain NCIMB 9816, but in equivalent systems inhibited the degradation of the same contaminants by Sphingomonas yanoikuyae (Allen et al., 1999). Other studies have implicated biosurfactants in the alteration of cell hydrophobicity (causing cells to aggregate or form clusters) and also the disruption of bacterial attachment to soil and artificial matrices (Herman et al., 1997; Zhang and Miller, 1992). Clearly, the addition of surfactants to contaminated systems can enhance the desorption and mobility of hydrophobic contaminants, but the increased aqueous contaminant concentrations do not necessarily result in concomitant enhancement of biodegradation (Stelmack et al., 1999).
The primary goal of this study was to determine if biodegradation of soil-sorbed PHE could be accelerated through addition of biosurfactant or bioaugmentation with PHE-degrading and biosurfactant-producing bacteria. A series of batch aqueous and soil PHE mineralization experiments was conducted involving inoculation with various combinations of PHE-degrading and rhamnolipid-producing bacterial stains to investigate species-specific bioavailability and commensal interactions between the bacterial strains. We performed batch and column sorption–desorption kinetic experiments with and without biosurfactant and characterized cell surface properties to explain PHE mineralization patterns in inoculated soil microcosm studies.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONREFERENCES |
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Cultivation of Bacterial Strains
Pseudomonas aeruginosa ATCC 9027 was purchased from the American Type Culture Collection (Rockville, MD) and was cultured in proteose peptone glucose ammonium salts (PPGAS) media according to the method outlined by Zhang and Miller (1992). The PHE-degrading organisms, Pseudomonas strain R and isolate P5-2, were generously supplied by Dr. Martin Alexander (Cornell University, Ithaca, NY) and were grown at 30°C in a mineral salts medium containing PHE (1.0 g L-1) as the sole source of carbon and energy (Hatzinger and Alexander, 1995). Each organism was maintained at -70°C in frozen stock cultures containing 10% glycerol (by volume).
Surfactant Production and Extraction
Rhamnolipid was chosen for this study because it is one of the most commonly isolated and well-characterized biosurfactants and because it is more biodegradable and less toxic than synthetic surfactants. Rhamnolipid biosurfactant was produced under phosphorous-limited conditions by growing P. aeruginosa in 1.5 L PPGAS media in a 3.0 L Fernbach flask (Zhang and Miller, 1992). Rhamnolipid was extracted from the culture supernatant by precipitation with 0.25 M calcium chloride (1:2 v/v). The Ca–rhamnolipid precipitate was removed from solution by centrifugation (15 min, 19682 x g). After discarding the supernatant, the surfactant pellets were resuspended in sterile deionized water. Approximately 1.5 g W2 sodium-saturated Dowex HCR-cation exchange resin, 40-60 mesh (J.T. Baker, Phillipsburg, NJ), was added for every 100 mL solution and equilibrated overnight at 37°C to allow complete exchange of the Ca2+ for Na+ and resolubilization of the rhamnolipid molecules. After equilibration, the solution was filtered through a wire mesh filter to remove the resin.
For mineralization experiments, biosurfactant was purified by a solid-phase extraction technique using Supelclean LC-18 bulk packing material (Supelco, Bellefonte, PA). A 60-mL syringe was packed with approximately 5.0 g of the LC-18 packing material and then preconditioned with 40 mL methanol followed by one rinse with 40 mL 0.01 M phosphoric acid (pH 3.0). After the conditioning steps, 40 mL of rhamnolipid was loaded onto the packing material. The column was then washed with a mixture of 25% acetonitrile (ACN) and 75% (v/v) phosphoric acid solution (0.01 N, pH 3.1) to remove impurities. Rhamnolipid was eluted from the column with ACN and phosphoric acid (75:25% v/v). Acetonitrile was removed from the purified surfactant by evaporation. In all other experiments the rhamnolipid solution was extracted from the culture supernatant according to methods outlined above and filter-sterilized (0.22 µm), and the resulting solution was used without further purification.
Radiolabeled rhamnolipid was produced by growing P. aeruginosa in PPGAS media containing only a portion of the glucose requirement (0.027 M). The remaining glucose was added to the cultures after 24 h of incubation (37°C, 120 rpm) as d-glucose-UL-14C (0.003 M, 20 µCi per 200 mL culture broth). The culture was incubated for 9 d and then processed according to the method outlined above. Confirmation that the 14C label was associated with the rhamnolipid was achieved via reverse-phase high performance liquid chromatography (HPLC) (Thermo Separation Products, Riviera Beach, FL). The HPLC was equipped with a vacuum degasser, a CM 4100 pump, an AS 3000 autosampler, a Spectra Focus UV–VIS detector and a Packard (Meriden, CT) Model A150TR Flow Scintillation Analyzer. Samples were injected into an Alltima C18 column (Alltech, Deerfield, IL) and eluted with a gradient of ACN–phosphate buffer (0.01 M H3PO4) as follows: ACN–phosphate buffer (5:95% v/v) for the first 5 min followed by a linear gradient up to ACN–phosphate buffer (70:30% v/v) over 38 min. In order to quantify radioactivity in the eluted samples, Ultima Flo M scintillation cocktail (Packard) was delivered at a flow rate of 3 mL min-1 to provide a 3:1 (v/v) ratio of scintillant to mobile phase.
Due to its structure, rhamnolipid could not be detected and quantified spectrophotometrically. Therefore, fractions were collected from the eluted HPLC injections and the rhamnolipid concentration in each fraction was estimated by a modified orcinol assay (Koch et al., 1991). In this manner, rhamnolipid concentrations in purified extracts of culture media and the incorporation of 14C label into surfactant could be quantified.
Aqueous Phenanthrene Utilization
Aqueous PHE mineralization experiments were carried out to estimate rates of PHE degradation by isolate P5-2 and P. strain R. The cultures were incubated in a mineral salts media (Hatzinger and Alexander, 1995) and PHE was added at concentrations of approximately 0.18, 0.3, 0.45, 0.6, and 0.9 mg L-1. Prior to inoculation, the cells were prepared by washing and resuspension in phosphate-buffered saline solution (PBS) containing (per L) 8.0 g NaCl, 0.2 g KCl, 0.12 g KH2PO4, and 0.91 g Na2HPO4 to a final cell density of 1 x 107 cells mL-1.
Aqueous PHE concentrations were determined at 1-h intervals by reverse-phase HPLC. Samples (50 µL) were injected into a 10-cm Lichrosphere RP-C18 column with a particle size of 5.0 µm (EM Science, Darmstadt, Germany) and eluted with an isocratic mobile phase consisting of 80% acetonitrile and 20% water (v/v) delivered at a flow rate of 1.0 mL min-1. Eluted peaks were monitored by UV absorption at 250 nm.
Batch Sorption Isotherms and Desorption–Release Kinetics
Phenanthrene sorption isotherms and desorption–release kinetics were evaluated in 5.0-mL glass HPLC vials, with Teflon-coated closures to assure minimal PHE sorption to experimental materials. Sterile soil (1.0 g) was added to each vial and then enough PHE was added with an acetone carrier to saturate the sorption capacity of the soil and achieve a soil solution concentration of 1 mg L-1 at equilibrium. This amount (Mv) was calculated based on the organic matter content of each soil, the Koc of PHE, and a moisture content sufficient to reach field capacity (Liu et al., 1991).
The experiments were conducted using a series of PHE concentrations ranging from the maximum, Mv, to 75, 50, 25, and 12.5% of Mv. These values corresponded to a range of 0.005 to 0.280 mg PHE g-1 soil. In addition, soil free controls were used to assess the fraction of PHE sorbed to the vial and other losses during equilibration. Background solutions containing 10 mM CaCl2 and biosurfactant at 0, 2.0, 5.0, and 10.0 times the CMC (0 to 500 µg mL-1 as rhamnose equivalents) were added to reach a final soil–water ratio of 1:5 (w/v). For sorption isotherms, the samples were equilibrated for 36 h at room temperature with gyratory shaking at 140 rpm. After equilibration, the vials were centrifuged (6929 x g) and the supernatant was analyzed for PHE by reverse-phase HPLC as described previously. Phenanthrene desorption–release was assessed via destructive sampling at 12, 24, 72, and 120 h. In this experiment, soil was preequilibrated with PHE as described above. After equilibration, the tubes were centrifuged, and the equilibration solution was replaced with an equivalent volume of PHE-free solution without or with varying concentrations of rhamnolipid (2, 5, or 10 times the CMC) and PHE concentrations were analyzed by HPLC as stated previously.
Phenanthrene Mineralization
The mineralization of PHE in aqueous systems was determined by quantification of 14CO2 released from [9-14C]PHE in 125-mL flasks. Each flask contained 50 mL of mineral salts media containing PHE at a concentration of 0.1 mM and 0.05 µCi [9-14C]PHE (Hatzinger and Alexander, 1995). The flasks were inoculated with P. strain R, isolate P5-2, and/or P. aeruginosa (ATCC 9027) after the cells were cultured and washed in sterile PBS according to the methods outlined above. Filter-sterilized air was passed through a flask containing 2.0 M NaOH to remove CO2. The CO2-free air was then hydrated in a water trap before passing through the culture flasks. In this manner 14CO2 was sparged from the system into an external trap (0.5 M NaOH) that was changed daily.
Phenanthrene mineralization in the presence of rhamnolipid was assessed in 50 mM Tris-HCl buffer (pH 7.2) containing a total PHE concentration of 0.1 mM, and 0.05 µCi[9-14C]PHE. Rhamnolipid was added at concentrations of 0.5, 1.0, and 2.0 times the CMC (25 to 100 µg mL-1 as rhamnose equivalents). Each flask was equipped with a trap (1.8-mL HPLC vials suspended from rubber stoppers by copper wire) that contained 1.0 mL 0.5 M NaOH. The cultures were incubated at room temperature with gyratory shaking at 110 rpm and sampled daily by replacing the entire trap and its contents.
Serum bottles (50 mL) were used as microcosms in soil mineralization experiments and were equipped with traps as described above. Phenanthrene was added to sterile soil at a concentration of 5.0 mg kg-1 and 0.05 µCi [9-14C]PHE in an acetone carrier. The acetone was allowed to evaporate and then the microcosms were inoculated with P. aeruginosa, P. strain R, and/or isolate P5-2 to a final cell density of 2 x 106 cells g-1. Soils inoculated with a single isolate (PHE-degrading strain or the biosurfactant producer) were included as controls. Rhamnolipid biosurfactant was diluted in sterile deionized water and added at concentrations of 0, 0.5, 1.0, and 5.0 times the CMC (0 to 250 µg mL-1 as rhamnose equivalents). The microcosms were incubated at room temperature (21°C) and sampled periodically by replacing the entire trap and its contents.
Rhamnolipid Mineralization
The ability of each test organism to degrade rhamnolipid biosurfactant was analyzed in both soil and aqueous systems by monitoring 14CO2 production from 14C-rhamnolipid. The aqueous experiment was carried out in 125-mL Erlenmeyer flasks containing 50 mL of 50 mM Tris-HCl buffer (pH 7.2) and 14C-rhamnolipid (0.014 µCi) as the sole source of C and energy. Rhamnolipid mineralization in soil was assessed in microcosms identical to those used in the PHE degradation experiments except that rhamnolipid was added at a concentration of 1.5 times the CMC (75 µg g-1 as rhamnose equivalents), with 0.0025 µCi 14C-rhamnolipid as the sole source of C and energy. Inocula were prepared by washing and resuspending the test strains in PBS as stated previously and samples were taken periodically by replacing the entire trap and its contents.
All mineralization experiments were performed in triplicate in separate culture flasks. Radioactivity in the traps was determined in a Tri-Carb 2900TR Liquid Scintillation Analyzer with automatic quench correction (Packard). Samples of trapping solution were added (1:4 v/v) to Scintisafe Econo1 scintillation cocktail (Fisher Scientific, Suwanee, GA) prior to analysis.
Carbon-14-UL-Glucose Respiration in Rhamnolipid-Amended Cultures
The potential inhibition of PHE degradation due either to toxicity or preferential utilization of rhamnolipid biosurfactant as a substrate was assessed by monitoring 14CO2 production from 14C-glucose in aqueous cultures. Glucose was selected as a substrate for this purpose because solute–solute interactions between glucose and rhamnolipid were expected to be negligible. Therefore, any inhibition or enhancement in glucose mineralization in the presence of rhamnolipid could be attributed to a direct physiological effect of surfactant on the test bacterial strains. Cells were cultivated in mineral salts media by the methods previously described (Hatzinger and Alexander, 1995). Glucose was added at a total concentration of 0.1 mM, and 0.1 µCi 14C-UL-glucose. Rhamnolipid biosurfactant was added to the flasks at concentrations of 0, 0.5, 1.0, and 3.0 times the CMC (as rhamnose equivalents). The flasks were incubated at room temperature (21°C) with gyratory shaking at 120 rpm and the alkaline traps were sampled periodically as described above.
Determination of Cell Hydrophobicity
Relative cell hydrophobicity measurements were made using the bacterial adhesion to hydrocarbon (BATH) assay as described by Zhang and Miller (1994). Briefly, cells were cultured, washed twice in PBS, and resuspended to an optical density of 0.5 (660 nm). Hexadecane was added to the cell suspensions (1.0 mL hexadecane to 4.0 mL culture) in screw-top test tubes. The tubes were vortexed at maximum speed for 1.0 min (Genie 2 vortex, Setting 8; Fisher Scientific) and then the phases were allowed to separate for approximately 45 min. After separation, the aqueous phase was carefully removed and the optical density (660 nm) was determined again. Relative hydrophobicity was calculated as the percentage of cells adsorbed to the hexadecane phase.
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RESULTS |
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To assess potential inhibitory effects of rhamnolipid on the test organisms, respiration of a nonpartitioning substrate, glucose, was monitored in aqueous cultures amended with biosurfactant and 14C-glucose as a primary carbon and energy source. Glucose mineralization by isolate P5-2 was inhibited when rhamnolipid was present, particularly at the higher concentrations, suggesting that the surfactant was potentially toxic to this strain (data not shown). The effect of rhamnolipid on glucose mineralization by P. strain R was less clear. Glucose mineralization by this strain was enhanced in the presence of rhamnolipid at 0.5 times the CMC, but the presence of rhamnolipid at 1.0 times the CMC had no effect. Concentrations of rhamnolipid at 3.0 times the CMC dramatically reduced glucose mineralization by this strain (data not shown).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONREFERENCES |
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Evidence supporting the hypothesis that some organisms have access to sorbed chemicals is slowly accumulating, but the actual mechanism of uptake is still unknown. Studies conducted by Poeton et al. (1999) showed that the presence of PHE in sediment samples increased bacterial adsorption to the soil matrix by about 30%. In addition, PHE and fluoranthene degradation rates were two to five times faster in sediment–water systems than in pure aqueous systems (Poeton et al., 1999). The enhanced mineralization rates were attributed to increased sorption of bacteria onto the sediment surface, which in turn increased the likelihood of interactions between the bacteria and sorbed PAH. Harms and Zehnder (1995) observed that degradation of 3-chlorodibenzofuran from porous Teflon beads by attached bacteria resulted in enhanced desorption that exceeded the rate observed in abiotic systems where desorption was driven by continuous replacement of the aqueous phase. Similar observations have been made with organisms growing on naphthalene as a sole source of C and energy (Crocker et al., 1995; Guerin and Boyd, 1992). Crocker et al. (1995) proposed that the utilization of sorbed naphthalene by P. putida strain 17484 at the surface of the sorbent promoted the desorption and dissolution of more naphthalene from deep within the soil aggregate (Crocker et al., 1995). An alternative hypothesis also involving attachment suggests that cells are chemotactically attracted to higher concentrations of substrate dissolved in thick static boundary layers at the solid–water interface. The attached cells degrade substrate until it is depleted and then detach to seek other resources (Guerin and Boyd, 1992).
Cell–Cell Interactions
In aqueous cultures, the presence of P. aeruginosa inhibited PHE mineralization by isolate P5-2, but enhanced PHE mineralization by P. strain R. In addition, co-inoculation with the rhamnolipid-producing bacterium enhanced PHEmineralization in the low organic matter Kenansville soil inoculated with P. strain R. Moreover, when rhamnolipid biosurfactant was added to these co-inoculated microcosms PHE mineralization was significantly reduced. All of the results point to the possibility that the surfactant disrupted cell–cell interactions between the two strains or that it inhibited cell attachment to the soil particle surface. Rhamnolipid has been shown to cause cell aggregation in some exposed species and has resulted in changes in cell hydrophobicity and prevented attachment of bacteria to NAPL–water interfaces (Herman et al., 1997; Ortega-Calvo and Alexander, 1994; Stelmack et al., 1999). Based on the reduction in glucose mineralization in P. strain R cultures amended with high surfactant concentrations, rhamnolipid toxicity to P. strain R cannot be excluded as another possibility for the reduced rate of PHE mineralization observed in this system.
To our knowledge this is the first reported observation of commensal interactions between bacterial species resulting in the enhanced utilization of a sorbed substrate. Given the results of the present study, it is unknown if P. aeruginosa benefits from this interaction, and thus the relationship cannot be termed synergistic or mutualistic. Phenanthrene mineralization was minimal in control soil microcosms inoculated with P. aeruginosa alone, suggesting that the biosurfactant-producing strain contributed to PHE mineralization through an indirect mechanism rather than metabolizing PHE directly.
Effects of Rhamnolipid Additions
A primary goal for conducting parallel experiments with exogenously added surfactant was to simulate a scenario where P. aeruginosa would be releasing rhamnolipid in co-inoculated systems. The mineralization patterns in aqueous co-inoculated experiments (i.e., co-inoculation with P. aeruginosa inhibited PHE mineralization by P5-2 but enhanced PHE mineralization by P. strain R) could not be reproduced by adding rhamnolipid biosurfactant. Therefore, some other mechanism, at least in part, must have been responsible for the enhanced mineralization observed in co-inoculated experiments.
Addition of the rhamnolipid clearly enhanced release of PHE from soil and thus had the potential for enhancing PHE mineralization in the inoculated soil microcosms. Surfactant additions had little effect on the PHE mineralization rate in either soil inoculated with P. strain R alone, but reduced PHE mineralization in co-inoculated systems containing P. strain R and P. aeruginosa. These results were consistent with our hypothesis that attachment of P. strain R cells to soil particle surfaces combined with its efficient use of low concentrations of PHE enabled it to access and utilize sorbed PHE. In bacterial adhesion to hydrocarbon studies (BATH assay), approximately 36% of P. strain R cells adhered to the hexadecane–water interface. Addition of rhamnolipid resulted in a fivefold reduction in P. strain R adhesion to hexadecane, indicating that this strain had a relatively hydrophobic cell surface under the conditions of our assay. These results suggest that attachment to soil may account for the more efficient degradation of PHE by P. strain R. Co-inoculation of soil with P. strain R and P. aeruginosa enhanced PHE mineralization, suggesting that some unknown interaction was responsible for the enhancement in PHE mineralization in these systems. Although rhamnolipid concentrations were not directly measured in the co-inoculated microcosms, it is very unlikely that P. aeruginosa could have produced rhamnolipid in sufficient quantity to enhance PHE mineralization to the observed level given the experimental conditions of the study. The fact that the addition of exogenous rhamnolipid to co-inoculated systems reduced PHE mineralization to levels that were comparable with microcosms inoculated with P. strain R alone indicates that the surfactant disrupted some unknown cell–cell interaction(s) necessary for the enhanced level of PHE mineralization observed in identical soil microcosms without added surfactant.
The addition of surfactant to microcosms inoculated with strain P5-2 alone did not enhance PHE mineralization, nor did co-inoculation with P. aeruginosa. However, in the co-inoculated system containing strain P5-2 and P. aeruginosa, surfactant addition dramatically increased PHE mineralization in a reproducible fashion. In these systems, PHE mineralization increased in proportion to added surfactant concentration and so it appears that the added surfactant in some way enhanced cell–cell interactions between isolate P5-2 and P. aeruginosa, which resulted in an increased rate and extent of PHE mineralization. The results of the BATH assay revealed that isolate P5-2 possessed a relatively polar cell surface, with only 15% of the added cells adhering to the hexadecane–water interface. Rhamnolipid addition did not appear to affect attachment of strain P5-2 to the hydrocarbon–water interface, but the results were inconclusive (data not shown). At present, it is impossible to explain the observed effects of co-inoculation and the addition of biosurfactant on PHE mineralization by strain P5-2.
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CONCLUSION |
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ACKNOWLEDGMENTS |
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