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Applied and Environmental Microbiology, May 2002, p. 2509-2518, Vol. 68, No. 5
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.5.2509-2518.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Assessing the Role of Pseudomonas aeruginosa Surface-Active Gene Expression in Hexadecane Biodegradation in Sand

P. A. Holden,1* M. G. LaMontagne,1 A. K. Bruce,1 W. G. Miller,2 and S. E. Lindow3

Donald Bren School of Environmental Science & Management, University of California, Santa Barbara,1 USDA Research Agricultural Service, Albany,2 Department of Plant and Microbial Biology, University of California, Berkeley, California3

Received 19 October 2001/ Accepted 24 January 2002


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ABSTRACT
 
Low pollutant substrate bioavailability limits hydrocarbon biodegradation in soils. Bacterially produced surface-active compounds, such as rhamnolipid biosurfactant and the PA bioemulsifying protein produced by Pseudomonas aeruginosa, can improve bioavailability and biodegradation in liquid culture, but their production and roles in soils are unknown. In this study, we asked if the genes for surface-active compounds are expressed in unsaturated porous media contaminated with hexadecane. Furthermore, if expression does occur, is biodegradation enhanced? To detect expression of genes for surface-active compounds, we fused the gfp reporter gene either to the promoter region of pra, which encodes for the emulsifying PA protein, or to the promoter of the transcriptional activator rhlR. We assessed green fluorescent protein (GFP) production conferred by these gene fusions in P. aeruginosa PG201. GFP was produced in sand culture, indicating that the rhlR and pra genes are both transcribed in unsaturated porous media. Confocal laser scanning microscopy of liquid drops revealed that gfp expression was localized at the hexadecane-water interface. Wild-type PG201 and its mutants that are deficient in either PA protein, rhamnolipid synthesis, or both were studied to determine if the genetic potential to make surface-active compounds confers an advantage to P. aeruginosa biodegrading hexadecane in sand. Hexadecane depletion rates and carbon utilization efficiency in sand culture were the same for wild-type and mutant strains, i.e., whether PG201 was proficient or deficient in surfactant or emulsifier production. Environmental scanning electron microscopy revealed that colonization of sand grains was sparse, with cells in small monolayer clusters instead of multilayered biofilms. Our findings suggest that P. aeruginosa likely produces surface-active compounds in sand culture. However, the ability to produce surface-active compounds did not enhance biodegradation in sand culture because well-distributed cells and well-distributed hexadecane favored direct contact to hexadecane for most cells. In contrast, surface-active compounds enable bacteria in liquid culture to adhere to the hexadecane-water interface when they otherwise would not, and thus production of surface-active compounds is an advantage for hexadecane biodegradation in well-dispersed liquid systems.


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INTRODUCTION
 
Natural biological attenuation is the present strategy for remediating greater than 25% of soil sites contaminated with petroleum hydrocarbons (33). Because of insufficient knowledge about in situ controls on bioavailability and biodegradation of low-solubility hydrocarbon pollutants in soil, it is difficult to predict cleanup end points (33) and to select appropriate remediation strategies. Reported mechanisms for bacterial metabolism of sparingly soluble hydrocarbons are (i) dissolution and diffusion of dissolved hydrocarbons to cells with uptake via active or passive transmembrane transport, (ii) invagination of hydrocarbon non-aqueous-phase liquid (NAPL) into cells (9) with subsequent intracellular metabolism of the hydrocarbon inclusions, and (iii) bacterial production of surface-active compounds such as surfactants (biosurfactants) and emulsifiers (bioemulsifiers) that increase the local pseudosolubility of hydrocarbons and thus improve mass transfer to biodegrading bacteria (3, 20, 29). Compared with synthetic surfactants, which can be either stimulatory below the critical micelle concentration (23) or inhibitory above the critical micelle concentration (28), biosurfactants reduce the interfacial tension in two-phase NAPL-water mixtures, while being nontoxic and fully biodegradable (40). Biosurfactants have been studied largely in liquid culture with a common soil bacterium, Pseudomonas aeruginosa. Many other soil bacteria produce surface-active compounds, but little is known regarding the production of biosurfactants in soils (29). More knowledge regarding in situ production of surface-active compounds would enhance our understanding of natural bioattenuation. Furthermore, if surface-active compounds are produced in soil and could be stimulated in situ, engineered bioremediation could be enhanced.

P. aeruginosa produces rhamnolipids that improve biodegradation of various hydrocarbons in liquid culture (17, 27, 50-52) when either N (11, 12, 31), P (11, 32), or Fe (11) is limiting and with a variety of carbon substrates, including glucose (11, 12) and hexadecane (17, 22, 44). Rhamnolipid synthesis in P. aeruginosa is encoded by genes in the rhl operon (35) which are positively regulated by rhlR, a luxR homolog (36). P. aeruginosa also produces a bioemulsifying protein, PA (18), which is encoded by the pra gene (14); PA is produced under similar conditions of nutrient limitation.

Rhamnolipids purified from liquid culture, when amended to soil, improve residual hydrocarbon recovery (5, 6, 15, 48) and biodegradation (16, 24). However, if soil bacteria produce rhamnolipids in situ, the costs associated with laboratory production and soil amendment could be deferred. To date, there is little direct evidence that either biosurfactants or bioemulsifying proteins are produced in situ in hydrocarbon-contaminated porous media, e.g., soil. In fact, it has been suggested that microbial surfactants in liquid culture may be simply cell surface components that are dispersed into solution following cell lysis or excreted during unbalanced growth (13, 20, 40, 41). The dispersive conditions of well-mixed liquid cultures do not occur in soil, so these compounds may play a different, and perhaps cell surface-limited, role in interactions of cells with insoluble hydrocarbons. In this study, rather than infer from liquid culture experiments the possible production of biosurfactants in soil, we used expression vectors to determine whether P. aeruginosa transcribes genes for surface-active compounds when cultured in unsaturated porous media. Furthermore, we asked whether the ability to produce surface-active compounds at native levels in unsaturated porous media enhances hexadecane biodegradation. Our results provide direct evidence that genes for surface-active compounds are transcribed by P. aeruginosa in porous media. However, while genes that encode or regulate surface-active compounds were expressed in P. aeruginosa in unsaturated porous media, gene expression was not tied to measurably improved hexadecane biodegradation rates or end points.


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MATERIALS AND METHODS
 
Chemicals, media, strains, and plasmids.
The strains and plasmids used in this study are summarized in Table 1. All strains were maintained at -80°C and cultured on Luria-Bertani agar (LA) for 24 h prior to use. When used, kanamycin, ampicillin, and tetracycline (TET) were added at final concentrations of 50, 150, and 30 µg/ml, respectively.


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  		TABLE 1. Strains and plasmids used in this study

Chemicals were purchased from Sigma Chemical Co. (St. Louis, Mo.). The basal medium was either half-strength 21C (19) or a variation that replaced ammonium N with equimolar nitrate N that we named CW21C. Filter-sterilized (0.2-µm-pore-size filter) hexadecane was provided as the sole carbon source. All other hydrocarbons used in green fluorescent protein (GFP) induction studies were filter sterilized. The stock solutions for the liquid-culture GFP induction studies were filter-sterilized glucose (0.33 g/ml), NH4Cl (50 mg/ml), and NaNO3 (79 mg/ml). The bacterial suspension used in drop slides was prepared in 1 mM phosphate buffer (P buffer) consisting of 4 ml of 0.2 M K2HPO4 and 1 ml of 0.2 M KH2PO4 per liter (pH 6.8). Restriction enzymes were purchased from Promega (Madison, Wis.), Roche Molecular Biochemicals (Indianapolis, Ind.), or Stratagene (La Jolla, Calif.). DNA sequencing as well as synthesis of the inverse PCR and pra probe oligonucleotides were performed at the University of California, Berkeley (http://mcb.berkeley.edu/barker/dnaseq). All other oligonucleotides were synthesized by Operon Technologies (Alameda, Calif.).

Acquisition of the pra promoter fragment by inverse PCR.
Although a short region upstream of the structural gene for the PA protein (pra) in PG201 had been characterized (14), the complete promoter element for pra, including all putative regulatory elements, was not available. Therefore, inverse PCR (34) was used to generate a complete pra promoter-containing fragment. The pra gene contains several unique restriction sites, including an NaeI site. To determine if this NaeI site would be suitable for inverse PCR, total chromosomal DNA was extracted from PG201 by using the QIAamp system (Qiagen, Santa Clarita, Calif.) and digested with NaeI. The digest was Southern blotted and probed with a digoxigenin-labeled PCR product. The probe, labeled with the Genius kit (Roche Molecular Biochemicals), contains a portion of the pra gene amplified from upstream of the NaeI site with the oligonucleotide primers PROBE-5' (5'-CGG-ACG-GGA-AGG-CAC-AAG-GTG-GTC-G-3') and PROBE-3' (5'-CCG-GAT-TAC-GGG-TTG-ACT-ACG-3'). We confirmed by positive hybridization that a 500-bp NaeI fragment contained part of the pra coding region.

NaeI-digested PG201chromosomal DNA was circularized in a 20-µl ligation reaction mixture (12 h, 18°C) containing 18 µl of 10x ligation buffer, 1 µl of DNA, and 1 µl of ligase (34). The circularized DNA was purified and concentrated by ethanol-sodium acetate precipitation prior to use as template DNA in the inverse PCR (34). The pra 5' flanking region, containing the putative complete pra promoter, was amplified using the oligonucleotide primers INVERSE-5'(5'-GCG-GAT-CCT-GCC-TGA-GCG-TTT-CGT-CC-3'; bp +49 to +63) and INVERSE-3' (5'-CCG-GAT-CCG-ATT-TCA-TTG-TTT-TGT-ATC-C-3'; bp -12 to +8), where bases in boldface are identical to sequences in the pra region and numbers are relative to the start (+1) of pra. Inverse PCR was conducted (1.5 mM final MgCl2 concentration) in a DNA Thermal Cycler (Perkin-Elmer Cetus) using 35 cycles of 94°C for 1 min, 50°C for 2 min, and 72°C for 3 min. The PCR product was cloned into pCR2.1 (Invitrogen, Carlsbad, Calif.) and transformed into DH5{alpha}. Plasmid DNA was extracted from colonies containing the insert (as determined by blue-white screening) and digested with EcoRI to establish which clones contained the ca. 500-bp NaeI pra fragment. A 500-bp fragment was recovered from one selected clone and sequenced (Fig. 1) to confirm that it contained the 5' end of pra in addition to a significant region upstream of the structural gene. The sequence of the upstream region indicated that the INVERSE-5' primer hybridized to a secondary location, resulting in a smaller-than-expected PCR product. Thus, because the INVERSE-5' primer hybridized downstream of the NaeI site and the INVERSE-3' hybridized upstream, the amplified product contains no NaeI site. The EcoRI-ended gene fragment containing the putative pra promoter was cloned into the EcoRI site of pUC1813 (42). In one orientation (pUC1813PRA), the promoter fragment contains upstream and downstream restriction sites for HindIII and KpnI, respectively.



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  		FIG. 1. Sequence of the inverse PCR product containing the putative promoter for pra, a structural gene encoding the PA bioemulsifying protein. An 8-bp region homologous to the pra structural gene (GenBank accession no. L08966) is italicized. The Shine-Dalgarno region (S/D) is underlined. PCR primers as described in the text are in boldface.

Plasmid construction and transformation.
The regulatory element of either pra or rhlR was cloned upstream of gfp in the promoter probe vector pPROBE-TT (30); these fusions were transformed into DH5{alpha}. To construct the Ppra-GFP fusion, pPROBE-TT was digested with HindIII and KpnI and the putative pra promoter fragment was ligated into the multicloning site by using standard protocols; the ligation product, plasmid pHX1, was transformed into competent DH5{alpha} (Gibco, Rockville, Md.). Transformants were selected from colonies that developed on LA amended with TET (LA-TET). The rhlR promoter region (1.1 kb) was amplified from PG201 DNA using primers FORRHL (5'-CAG-AAG-CTT-TTC-CTG-CAA-TCC-GAC-3') and REVRHL (5'-CCG-GTA-CCT-GGT-CGA-TGT-GAA-A-3'), which contain restriction sites for HindIII (forward) and KpnI (reverse), according to standard protocols. The rhlR promoter-containing fragment was cloned into pPROBE-TT as before to produce pRhlPGB. Based on prior work with Pseudomonas species (7), gfp-containing plasmids were not expected to significantly burden cells.

Late-exponential-phase, Luria-Bertani broth (LB)-grown PG201 cells were made competent for transformation by treatment with 100 mM MgCl2 (30 min, 4°C) followed by resuspension in 100 mM CaCl2. Competent cells were combined with purified plasmid DNA (pHX1, pRhlPGB, or pPROBE-TT) and then incubated (4°C, 30 min) and heat shocked (2 min, 42°C) prior to addition of fresh LB and a second incubation (60 min, 30°C). Cells were spread at several dilutions on LA-TET, and the plates were incubated (32°C, 48 h). Transformants were confirmed to harbor the gfp-containing plasmid by colony PCR of LA-TET-grown cells with previously published gfp primers (47). The stabilities of the pHX1 and pRhlPGB plasmids in PG201 were determined by cultivating the engineered strains overnight (150 rpm, 30°C) in LB-TET, resuspending cells in fresh LB, then subsampling over several days and spread plating samples onto LA-TET and LA.

Liquid and sand culturing.
All culturing was at 30°C. Liquid culturing was performed in shaking, gas-tight, 500-ml triple-baffled Nephelo flasks (Bellco, Vineland, N.J.) with Teflon-lined top caps and 18-mm Mininert (Valco Instruments, Baton Rouge, La.) side caps for gas-tight sampling. Liquid cultures were prepared with 90 ml of basal medium (either half-strength 21C or CW21C), 10 ml of Hutner's mineral solution (19), 200 µl of hexadecane, and 100 µl of inocula prepared by suspending cells (from overnight solid-medium culture) to similar optical densities (ODs) in basal medium. Liquid cultures were inoculated with 100 µl of cell suspension. At various times during cultivation, 1 ml was sampled for liquid tensiometry. Population density was monitored by culture absorbance at 660 nm.

Sand cultures were prepared using 100 g of autoclaved (three times, 30 min) clean quartz sand with grains that were previously classified as being between 212 and 180 µm in diameter (sizes 80 and 70 U.S. standard sieves, respectively). Culture bottles were 250-ml amber glass (Boston rounds; Fisher Scientific, Tustin, Calif.). Sand cultures were inoculated with a dilute 3.4-ml bacterial suspension, which resulted in an unsaturated porous medium. The inocula for the sand cultures were similar concentrations of washed (two times in basal medium) exponential-phase cells grown in LB. Uninoculated controls and inoculated sand cultures were mixed by rolling the reactor bottles for 30 min on a roller mill and then incubated overnight (30°C) prior to addition of hydrocarbon substrates. Addition of hexadecane (7 µl) resulted in an initial C/N ratio of 12 (equivalent to liquid culture). Gas and sand samples were removed periodically for analysis. Prior to sampling, the bottle contents were mixed completely on a roller mill for 20 min. Headspace sampling was through a 24-mm Mininert top cap. Sand was sampled using a sterile scoop after purging (1.2 liter/min for 3 min) the bottle with sterile, moist (100% relative humidity) air in a laminar flow hood. Sand samples (1 g) were preserved with 2 mg of dissolved NaN3, refrigerated (4°C for less than 48 h), and then hexane extracted.

gfp induction in well-mixed liquid broth cultures.
LB-TET-cultivated cells were used as inocula for studying GFP induction over time because they were initially nonfluorescent. Overnight LB-TET cultures were subsampled (10 ml), and the remaining culture volume (approximately 90 ml) was washed twice and resuspended in P buffer. The washed suspensions were divided into 9- or 10-ml subsamples, with a total of eight treatments. Amendments were made to the 10-ml washed subsamples, resulting in the following treatments: negative control (P buffer only), 2 µl of hexadecane per ml, 18.3 mM glucose, 36.7 mM glycerol, 9.3 mM NH4Cl, and 9.3 mM NaNO3. The proportion of N provided as either NH4Cl or NaNO3 was the same as that provided in our growth studies of P. aeruginosa PG201 and its mutants PG201pra-, PG201rhlA- and PG201pra-rhlA- (Table 1). The remaining two treatments consisted of 9 ml of washed cell suspension plus 100 µl of Hutner's mineral solution and 900 µl of either half-strength 21C or CW21C. The amount of nitrogen provided in the last two treatments was slightly less than that provided in our studies of PG201 and its mutants. The amended suspensions were shaken (30°C, 150 rpm) in covered 50-ml Erlenmeyer flasks. Over a 48-h period, 10 µl was periodically sampled (three times) from each suspension, a wet mount on a clean glass microscope slide was prepared, and at least 10 fields of view per slide were visualized first by epifluorescence microscopy for GFP and second by phase contrast for total cells. The apparent brightness and abundance of GFP-producing cells relative to the controls were noted, and qualitative scores were assigned for the purposes of comparing treatments.

gfp expression in drop slides.
Drop slides consisted of a coverslip overlying a special glass microscope slide with 10 wells created by a hydrophobic barrier (Precision Lab Products, Middleton, Wis.). A 0.5-µl drop of an overnight culture in LB-TET (washed twice and resuspended in P buffer) of PG201(pHX1), PG201(pRhlPGB), or PG201(pPROBE-TT) was dispensed into duplicate wells. A 0.5-µl drop of hexadecane was added adjacent to a P-buffer cell suspension drop prior to applying the coverslip. When the coverslip was lowered, both drops spread within the hydrophobic barrier. To minimize dehydration, the edges of the coverslip were sealed to the slide with a thin film of silicone lubricant (Dow, Midland, Mich.). The slides were monitored over time for GFP fluorescence by both epifluorescence microscopy and confocal laser scanning microscopy (CLSM).

Microscopy and fluorescence measurements.
GFP faded rapidly under the UV light source; therefore, epifluorescence microscopy was performed prior to phase-contrast imaging. GFP induction in liquid was determined using a Nikon E800 epifluorescence microscope with a GFP filter set (Chroma, Brattleboro, Vt.). The positive control for GFP was PG201(KtKan), which was cultivated on LA-kanamycin and suspended in P buffer to a cell density similar to that of the samples. The negative control was P. aeruginosa PG201 grown in LB, washed (two times), and resuspended. The relative amount of GFP produced per cell was inferred by the brightness of GFP-producing cells compared to positive and negative controls. The proportion of total cells expressing GFP was estimated by observing, for each field of view, the abundance of cells by phase contrast.

For visualizing GFP produced by sand-cultured bacteria, 1 g of moist sand was vortexed for 10 s with 12 ml of sterile H2O. The supernatant was allowed to clear for 1 h, and 5 ml was suction filtered onto a 13-mm-diameter, 0.2-µm-pore-size black Isopore membrane (Millipore, Bedford, Mass.). A second 1-ml aliquot from the same 12-ml suspension was stained with a nucleic acid-specific stain (SYBR-gold; Molecular Probes, Eugene, Oreg.), filtered, and visualized for total bacteria. Images of sand-cultivated gfp-expressing bacteria and total bacteria were captured using a DE750 color video camera (Optronics, Goleta, Calif.) mounted on an Olympus BX60 epifluorescence microscope with a fluorescein isothiocyanate filter set.

Drop slides were screened for GFP production by epifluorescence microscopy with a Nikon E800 microscope and then imaged in the x-y and x-z planes with a TCS SP2 CLSM (Leica, Mannheim, Germany) with a 63x water immersion objective and an Ar 488-nm excitation laser. The thicknesses of the z scans were approximately 0.2 µm.

Environmental scanning electron microscopy (ESEM) was performed in wet mode using a gaseous secondary electron detector in an FEI Co. XL30 ESEM FEG (Philips Electron Optics, Eindoven, The Netherlands). Images of PG201 colonizing sand under the specified experimental conditions were acquired for at least 10 subsamples of a 2-day-old culture.

Surface tension, CO2, and hexadecane measurements.
The air-water interfacial (surface) tension of liquid cultures was measured for 1.5 ml of either whole broth cultures or cell-free culture broth (supernatant from centrifugation at 10,000 x g for 10 min), using a liquid tensiometer (model KT-10; Krüss USA, Charlotte, N.C.), 10-ml (concentrated-H2SO4-washed) beakers, and a small-volume plate. A reduction in surface tension was indicated by any measurement lower than that of uninoculated media (72 mN/m).

Headspace CO2 analysis of sand cultures was performed with a GC14 gas chromatograph (GC) (Shimadzu, Kyoto, Japan) using a Porapak R column (Supelco, Bellefonte, Pa.), a TCD detector, and high-purity helium (30 ml/min) as the carrier gas. Concentrated hexane extracts of sand cultures were analyzed for hexadecane content using an HP 6890 GC with a mass spectrometer (MS) (Agilent Technologies, Palo Alto, Calif.). Unknowns were compared to a minimum of 15 standards spanning three calibration ranges for hexadecane in hexane. The GC-MS operating conditions were as follows: 1-µl sample volume, capillary column (Alltech AT-1), splitless injection, 250°C injector temperature, ultra-high-purity helium carrier gas at 30 cm s-1, initial oven temperature of 40°C (4 min), temperature increased by 10°C min-1 to 270°C then held (3 min), and detector temperature of 275°C. The GC-MS signal was automatically integrated using HP G1034C software for MS ChemStation (Agilent).

Data analysis.
Means from replicated measurements were compared using the method of Tukey (8) with a confidence interval of 0.05. Hexadecane depletion data from sand cultures were analyzed by curve fitting to determine the best fit to either zero-, first-, or second-order mathematical models (46). Depletion rate coefficients were calculated by linearizing the data for the appropriate mathematical model and performing regression analysis with Excel 2000. For the sand culture experiments, the biomass C originating from the utilization of hexadecane was calculated by subtracting the total CO2 carbon (mole basis) from the molar amount of hexadecane C utilized (total added hexadecane C). The molar fraction of carbon fixed was then calculated by dividing the calculated biomass C (molar basis) by the total molar addition of hexadecane C.


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RESULTS
 
Putative pra promoter.
The sequence of a region 5' to the pra gene in PG201, which includes the pra promoter, is provided in Fig. 1. A 459-bp region of the putative promoter-containing sequence was 99% identical with a homologous region in the P. aeruginosa PAO1 complete genome (GenBank accession no. AE004091). While not containing a consensus Shine-Dalgarno sequence, a region that could serve as a ribosome binding site was recognized (Fig. 1).

Growth and surface tension decrease in liquid culture.
We cultured PG201, PG201rhlA-, PG201pra-, and PG201pra-rhlA- with either NH4+ N or NO3- N to determine the effects of nitrogen on growth rate and the accumulation of surface-active compounds in liquid culture. Production of rhamnolipids and/or PA protein (and any other surfactant) will decrease liquid surface tension. The surface tension was lowest in late-exponential-phase cultures (Fig. 2). Surface tension was the same for whole cultures and cell-free supernatant (data not shown). When NH4+ was the N source, the strains had similar lag times and grew to similar maximum cell densities (Fig. 2) at similar rates (Table 2). Also when NH4+ was the N source, the amount of surface-active compounds was significantly smaller for cells deficient in rhlA, while deficiency in pra did not alter surface tension (Table 2).



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  		FIG. 2. OD at 660 nm (circles), liquid surface tension (squares), and headspace parts per thousand carbon dioxide (triangles) (data for PG201 and PG201pra- only) with either ammonium (closed symbols) or nitrate (open symbols) in liquid cultures of P. aeruginosa. Strains are either wild type (PG201) or deficient in PA protein synthesis (PG201pra-), in rhamnolipid synthesis (PG201rhlA-), or in both (PG201pra-rhlA-). Hexadecane is the carbon source. Each point represents the average of three or more independent replicates.


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  		TABLE 2. First-order growth rate coefficients and liquid culture density-normalized maximum liquid surface tension decrease for P. aeruginosa with hexadecane as the sole carbon source and either ammonium or nitrate as the nitrogen source

When NO3- was the nitrogen source, lag times were longer for all strains and PG201 grew to higher culture densities than the other strains (Fig. 2). The growth rates for PG201 with either NO3- or NH4+ were similar, but the growth rates for the other strains with NO3- were significantly lower (Table 2). While liquid surface tension decreased for all strains (Fig. 2), the decrease in surface tension for PG201, on a per-cell basis, was similar whether NH4+ or NO3- was the N source (Table 2). With NO3- N, the decrease in surface tension per cell was similar across strains (Table 2).

Hexadecane biodegradation in sand culture.
Relative to uninoculated controls, where no biodegradation activity was observed, GC analysis of hexane-extracted sand samples over time showed that hexadecane was rapidly and completely biodegraded in moist sand by all cultures, including strains deficient in pra and rhlA (Fig. 3). Lag times, hexadecane depletion, and CO2 generation patterns were similar for all strains. Among the rate models tested, hexadecane depletion best fit a first-order process (r2 > 0.9; r2 values for zero- and second-order models were lower). First-order depletion coefficients and the fraction of hexadecane C fixed into cellular material were statistically similar (n >= 3; {alpha} = 0.05) across the strains and averaged 0.093 h-1 and 0.35, respectively. ESEM images indicated that PG201 was dispersed on sand grains and did not appear as thick biofilms (Fig. 4).



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  		FIG. 3. Depletion of hexadecane (closed symbols) and accumulation of CO2 (open symbols) by four strains of P. aeruginosa PG201 in sand culture. Hexadecane is the sole C source; N is provided as either NO3- (top) or NH4+ (bottom). Strains are either wild type (PG201) or deficient in PA protein synthesis (PG201pra-), in rhamnolipid synthesis (PG201rhlA-), or in both (PG201pra-rhlA-). Points represent either the average of three independent replicates (NO3-) or single measurements (NH4+) at each time point.



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  		FIG. 4. ESEM images of P. aeruginosa PG201 in moist sand after 2 days of cultivation. NO3- was the N source and hexadecane was the carbon source. Magnifications, x2,500 (A) and x3,500 (B). Arrows point to typical cells or cell clusters.

Expression vector stability and gfp induction in liquid culture.
Significant GFP induction was observed in PG201 strains carrying pra or rhlR gene fusions when washed cells were subjected to a range of nutritional conditions. The expression vector was reasonably stable (95% after 2 days). Strain PG201(pPROBE-TT), harboring a promoterless gfp reporter gene only, was nonfluorescent (Table 3). Strain PG201(pHX1), in which gfp was driven by the pra promoter element, fluoresced brightest in the presence of hexadecane (Table 3). Strain PG201(pRhlPGB) fluoresced brightest and equally in the presence of either hexadecane, glucose, or ammonium-containing P buffer in the absence of a C source (Table 3). It is noteworthy that the fluorescent cells appeared to be mostly adherent to oil drops.


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  		TABLE 3. GFP fluorescence after 48 h in liquid culture in P. aeruginosa harboring fusions of promoters for pra and rhlR to a gfp reporter plasmid

gfp expression in sand culture.
We cultured PG201(pHX1), PG201(pRhlPGB), and PG201(pPROBE-TT) in moist sand to determine if the promoters for rhlR and pra are active in unsaturated porous media. We used nutritional conditions (hexadecane C and NO3-) that, in liquid culture, appeared to stimulate the production of surface-active compounds (Fig. 2 and Table 2). Epifluorescence microscopy of eluted cells revealed that PG201(pHX1) and PG201(pRhlPGB) produced GFP in moist sand culture with hexadecane as the sole C source and with nitrate N. No visible GFP fluorescence was observed in PG201(pPROBE-TT), which lacked a promoter sequence 5' to the gfp reporter gene. By directly counting total eluted Sybr-Gold-stained cells, we observed that approximately 3% of PG201(pHX1) and 1% of PG201(pRhlPGB) exhibited GFP fluorescence after 1 week of cultivation in sand. Similar numbers of total cells were present in each of the cultures at the time of sampling.

gfp expression in drop slides.
In our liquid culture GFP induction studies, we observed the highest fluorescence in cells exposed to hexadecane; most of those fluorescent cells were adherent to hexadecane drops. This led us to hypothesize that transcription of surface-active genes occurs mostly at the hydrophobic hexadecane-water interface due to a lack of availability of the substrate in the water phase. To test our hypothesis, we worked with static drop slides that would allow us to better visualize the spatial pattern of GFP production. As in sand cultures, PG201 (pPROBE-TT), the promoterless control strain, did not produce GFP in the drop slides (data not shown). Epifluorescence microscopy of the slides containing PG201(pHX1) and PG201(pRhlPGB), (i.e., examination from plan view), was inadequate to demonstrate where GFP was produced, because when viewed from the top, it appeared that green cells were contained both in water and in oil. This is because an oil-water-glass interface was formed by water wicking under the hydrophilic coverslip, on top of the oil, where the water resided as a thin film between the glass and oil. With CLSM, we observed that PG201(pHX1) and PG201(pRhlPGB) expressed gfp at the hexadecane-water and the hexadecane-water-glass interfaces (Fig. 5). An angle of approximately 60° was formed between the coverslip and the gfp-expressing cells, which were oriented along the hexadecane-water interface in the x-z direction by CLSM.



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  		FIG. 5. CLSM of drop slides containing P buffer, hexadecane, and P. aeruginosa strains PG201(pHX1) (A) and PG201(pRhlPGB) (B). Top frames are plan views; bottom frames are x-z sections. Fluorescence is from transcription of plasmid-borne gfp controlled by the putative promoters for pra (A) and rhlR (B). Bars, 20 µm.


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DISCUSSION
 
An objective of this study was to assess in situ production of surface-active compounds in porous media. Although a number of soil isolates produce surface-active compounds in broth culture (see, e.g., references 1, 2, 38, and 45), direct evidence of related gene transcriptional activity in sand or soil has been lacking, as is an understanding of the ecology of surface-active compound production in soil. Here we report the transcriptional activity in P. aeruginosa sand culture of a gene encoding PA emulsifying protein. However, despite this transcriptional activity of pra in sand culture, PG201 hexadecane biodegradation in sand was unaffected by the absence of intact pra or rhlA genes.

In liquid culture with NO3- as the sole N source, the deletion of either pra or rhlA slows growth at the expense of hexadecane by PG201 (Fig. 2) (14). Therefore, we expected that PG201 using NO3- in sand culture would degrade hexadecane faster than its mutants (with mutations in pra, rhlA or both). Instead, the biodegradation patterns were not different in sand culture whether PG201 was with or without intact genes encoding surface-active compounds. Our explanation is that the direct association between PG201 and hexadecane, which is facilitated by hexadecane spreading on sand surfaces where cells are adherent (Fig. 4), reduces the role for surface-active compounds in sand culture and favors the biodegradation mechanism of direct hydrocarbon uptake by PG201. Upon considering the spatial pattern of gfp expression, and taking into account prior reports, it is possible that different predominant uptake mechanisms govern hexadecane biodegradation in liquid versus sand.

Studies with liquid cultures of cells harboring a pra-gfp fusion revealed that surface-active gene expression by P. aeruginosa using hexadecane is spatially limited to the hexadecane-water interface. This may be because PG201 adheres to hexadecane, where transcription of genes (e.g., pra) for surface-active compounds is induced. Bacterial adherence to hydrophobic interfaces, i.e., the oil-water or the (also hydrophobic [21]) air-water interface (39, 43, 49), occurs with hydrophobic cells (43, 44). Cells may be hydrophobic already, or, in the case of P. aeruginosa, may become hydrophobic from their rhamnolipid production (50), which reduces the amount of exposed cell surface lipopolysaccharide (4). The transcriptional activity of rhlR in liquid media lacking alkanes was shown to be nearly constitutive at relatively low levels, with only a slight increase (twofold) upon entry of cells into the stationary phase with relatively high cell concentrations in cell broth (36). rhlR, by acting as a transcriptional activator of rhlA and rhlB, which are involved in rhamnolipid biosynthesis, thus is required for synthesis of this surfactant (27, 44) and also for adherence to hexadecane drops (44). We examined rhlR transcription as a way to ensure that cells were sufficiently active in our sand system to enable rhamnolipid synthesis. Indeed, we found that there was little variation in rhlR transcription under the various conditions tested here (Table 3) and that cells expressed considerable rhlR activity in both liquid and sand culture. Thus, the rhlR fusion served as a positive control for gfp expression, while transcriptional activity also suggests favorable conditions for cellular adhesion to hexadecane. On the other hand, the pra fusion appears to report specific transcriptional activity at the hexadecane-water interface. Intact pra and rhlA genes make PG201 grow faster with hexadecane and NO3- (14), indicating that these genes facilitate the use of hexadecane C by PG201 in liquid. Additionally, pra is inducible by hexadecane but not by glycerol or glucose (14). Taken together with our observations that pra promoter-driven gfp fusions fluoresced only at the hexadecane-water interface, these findings indicate that hexadecane appears to be a transcriptional activator for pra. Because hexadecane is so sparingly soluble, cellular adherence to the oil is a logical prerequisite for hexadecane to cause such transcriptional activity.

The involvement of rhamnolipids in cellular adhesion to hexadecane and the transcriptional activity of pra at the oil-water interface have different implications in liquid versus sand culture, which may explain why we observed relatively short lag times in sand culture as well as similar biodegradation patterns for strains proficient and deficient in production of surface-active compounds. We propose that cells in sand simply do not need surface-active compounds to facilitate rapid uptake of hexadecane because they are already in direct contact with the hydrocarbon. In sand culture, assuming a mean grain diameter of 200 µm and well-distributed hexadecane over all sand surfaces (a condition in our preparations), the total surface area of hexadecane is approximately equal to the sand surface area of ca. 1.2 x 1012 µm2. In liquid culture, this same surface area would arise from breaking the total amount of hexadecane into 1-µm-diameter drops. In liquid culture, aqueous media and oil are mixed via shaking, which is mainly intended to facilitate aeration. However, we typically observe unstable oil-in-water emulsions before the onset of bacterial growth. The emulsion is a distribution of oil drops of different sizes dispersed into the aqueous medium. The mean drop size of oil in an oil-water emulsion correlates positively with approximately the square root of oil-water interfacial tension, the mixing intensity, and the volume fraction of the dispersed phase (10, 37), which in the case of our PG201 liquid cultures would be 0.002. For chlorobenzene in water (interfacial tension of 0.0334 N/m), where the volume fraction of chlorobenzene was 0.005 and the stirring rate was 180 min-1, the mean oil droplet diameter was 215 µm (37). Hexadecane interfacial tension is more like that of n-dodecane, also a low-solubility alkane, which has an oil-water interfacial tension of 52.8 N/m (43). Considering that our liquid cultures were mixed and had volume fractions of oil similar to those in a previous study in which a more soluble hydrocarbon resulted in larger drops (37), it is inconceivable that the conditions of liquid culture we used could result in a hexadecane drop diameter of as small as 1 µm. The result is that the exposed surface area of hexadecane is lower in liquid culture than in sand culture. It would seem that making surface-active compounds, which both facilitates cellular adhesion to oil and improves oil bioavailability, is relatively advantageous to cells in liquid.

In sand culture, cells are distributed over the sand surface that is coated with hexadecane, and thus nearly every cell is in direct contact with oil. The high spatial opportunity for cellular adherence to hexadecane is advantageous to cells in that direct adherence is probably a faster route for hexadecane uptake. In liquid culture, there is a long lag time prior to hexadecane-associated growth (Fig. 2), which contrasts with the short lag time preceding hexadecane biodegradation in sand (Fig. 3). Also in liquid culture, the provision of nitrate as the N source, which has been shown to stimulate surface-active compound production, corresponds to rapid growth (Table 2). In both sand and liquid culture, hexadecane biodegradation is complete, but the patterns of biodegradation suggest differing mass transfer mechanisms. The data support the idea that cells in sand were already adherent to hexadecane while cells in liquid were dependent on their production of surface-active compounds to make them adherent and to increase the availability of hexadecane in solution. In contaminated soil, hydrocarbon spreading will enhance the contact area between bacteria and insoluble NAPLs. Our work supports the idea that bacterium-oil contact is a relatively more important factor for accelerating hexadecane biodegradation in unsaturated porous media than is the indigenous production of surface-active compounds.


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ACKNOWLEDGMENTS
 
Funding for this work was provided by National Science Foundation Award DEB-9805946 (to P. A. Holden), by National Science Foundation Awards BES-99772 and DMR-9724254, by U.S. Environmental Protection Agency Cooperative Agreement AERL9405 (to M. K. Firestone and J. R. Hunt, University of California, Berkeley), and by EPA/DOE/NSF/ONR Bioavailability Program Award R827133-01 (to P. A. Holden and A. Keller).

ESEM was performed by Jose Saleta in the MicroEnvironmental Imaging and Analysis Facility at the University of California, Santa Barbara (UCSB). CLSM was performed by Mario Yasa in the laboratory of Cyrus Safinya, Department of Physics and Materials Research Laboratory, UCSB. Joshua Schimel (UCSB) provided the instrumentation for CO2 analysis. We acknowledge James R. Hunt for helpful criticisms and the assistance of Portia Zamos, Claire Eustace, Cindy Wu, Sam Paik, Jennifer Lau, Perrin Pellegrin, Jennifer Guigliano, Don Herman, and Allen Doyle.


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FOOTNOTES
 
* Corresponding author. Mailing address: Donald Bren School of Environmental Science & Management, 4670 Physical Sciences Building North, University of California, Santa Barbara, CA 93106. Phone: (805) 893-3195. Fax: (805) 893-7612. E-mail: holden{at}bren.ucsb.edu. Back


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Applied and Environmental Microbiology, May 2002, p. 2509-2518, Vol. 68, No. 5
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.5.2509-2518.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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