Formation and Removal of Hydrocarbon Residual in Porous Media

David C. Herman, Robert J. Lenhard, and Raina M. Miller* ... The columns were then eluted with 1 mM rhamnolipid biosurfactant, which increased total r...
0 downloads 0 Views 140KB Size
Download PDF
Environ. Sci. Technol. 1997, 31, 1290-1294

Formation and Removal of Hydrocarbon Residual in Porous Media: Effects of Attached Bacteria and Biosurfactants DAVID C. HERMAN,† ROBERT J. LENHARD,‡ AND R A I N A M . M I L L E R * ,† Department of Soil, Water and Environmental Science, University of Arizona, Tucson, Arizona 85721, and Sultan Qaboos University, P.O. Box 34, Sultanate of Oman, 123

Column studies were used to investigate the fate of a representative nonaqeous-phase liquid (NAPL), hexadecane, with specific regard to (1) the effect of attached bacteria on the formation of residual saturation and (2) the role of biodegradation and biosurfactants on the removal of residual NAPL. Residual saturation of hexadecane was determined using sterile sand (40/50 mesh) columns and was found to be 19.0 ( 4.8% of the pore volume. Columns loaded with bacterial biomass (Pseudomonas aeruginosa ATCC 15442, 109 cells g-1) showed no difference in residual hexadecane formation as compared to sterile sand columns. In further column studies examining the effect of ATCC 15442 and biosurfactants on the removal of hexadecane residual, results showed that biodegradation alone removed approximately 50% of the [14C]hexadecane, in the form of 14CO2 and undefined cellular metabolites, during elution with at least 200 pore volumes of mineral salts medium. The columns were then eluted with 1 mM rhamnolipid biosurfactant, which increased total removal to 65%. Rhamnolipid addition resulted in (1) the mobilization of hexadecane free product and (2) a transitory 3-12-fold increase in the rate of hexadecane mineralization. In a separate study, the column was eluted from the beginning with a low (0.1 mM) concentration of rhamnolipid. This lower concentration of biosurfactant enhanced the removal of hexadecane by mobilization, but had no effect on the rate of biodegradation of residual hexadecane. Analysis of residual radioactivity within two columns revealed only 2% remaining as intact hexadecane. These results suggest that a combination of biodegradation and rhamnolipid treatment could be used to maximize the removal of residual NAPL from porous media.

Introduction Soil contamination by nonaqueous-phase liquids (NAPLs) is a serious problem because persistent NAPLs create a longterm source of contamination to subsurface waters. The displacement of NAPLs by water will create discrete ‘blobs’ or ganglia of NAPLs that are immobilized in the porous matrix (1, 2). The maximum amount of NAPL immobilized is a function of pore geometry and pore surface characteristics and is referred to as the residual NAPL saturation. The effect * Corresponding author telephone: 520-621-7231; fax: 520-6211647; e-mail: [email protected]. † University of Arizona. ‡ Sultan Qaboos University.

1290

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 31, NO. 5, 1997

of bacterial biomass on the formation of residual NAPL has not previously been evaluated. However, bacteria that adhere to pore surfaces can alter particle surface properties, and biomass coverage of particles has been linked to an increase in the transport of hydrophobic compounds through soil (3, 4). This may be due to a competition between bacteria and hydrophobic compounds for sorption sites. Similarly, the presence of bacteria on surfaces may affect residual NAPL formation. When components of a NAPL are suspected health hazards, actions need to be taken either to remove the source of the hazard (e.g., remove the NAPL) or to contain the contaminated area. Because the entrapped NAPL in the water-saturated zone of the subsurface is relatively immobile and discontinuous, it is very difficult and expensive to remove the NAPL hydraulically, such as with a pump-and-treat remediation technology. An alternative approach is the manipulation of the subsurface environment to create favorable conditions for biodegradation. Numerous papers have addressed biodegradation pathways and other biochemical aspects of the degradation of certain NAPLs (5, 6), but relatively little is known about the accessibility of NAPL immobilized in porous media. The first objective of this study was to quantify residual hexadecane saturation under saturated-flow conditions in sand-packed columns and to determine if the presence of attached bacteria could affect the amount of NAPL entrapped. Our second objective was to evaluate how bacteria and biosurfactants could mediate the removal of residual NAPL by biodegradation. Biosurfactants are of interest because their production is one strategy to allow the enhanced uptake and utilization of a NAPL as a carbon and energy source (79). Surfactants, including biosurfactants, can also enhance the removal of residual NAPL from porous media by mobilization (10, 11). The combined effect of mobilization and biodegradation on the removal of residual NAPL has not previously been evaluated. The experimental system chosen for this study was a biodegradable, less dense than water NAPL (hexadecane), sand-packed columns for the establishment of residual hydrocarbon under saturated flow conditions, a hydrocarbondegrading bacterium (Pseudomonas aeruginosa ATCC 15442), and a rhamnolipid biosurfactant. ATCC 15442 was chosen for this study because it has a relatively hydrophobic cell surface (8) and was expected to show strong irreversible adherence to sand particles. ATCC 15442 exhibits rapid growth on hexadecane as sole carbon source, and the rate of hexadecane degradation is increased in the presence of rhamnolipid biosurfactant.

Materials and Methods Chemicals. Hexadecane was obtained from Aldrich (Milwaukee, WI), and [1-14C]hexadecane, 2.2 mCi/mmol, was purchased from Sigma (St. Louis, MO). Radioactivity was determined using a Packard liquid scintillation analyzer Model 1600 TR (Packard Instruments, Meriden, CT). Unless otherwise noted, the scintillation cocktail used in this study was Scintiverse BD (Fisher Scientific, Pittsburgh, PA). Cultures. P. aeruginosa ATCC 15442 was obtained from the American Type Culture Collection (Rockville, MD) and stored at 4 °C on Pseudomonas agar P (Difco, Detroit, MI) plates. ATCC 15442 does not produce biosurfactants during growth on hydrocarbons as the sole carbon source (8). Bacterial suspensions used to inoculate sand columns were prepared by transferring cells from an agar plate into 100 mL of PTYG (peptone, 5 g L-1; tryptone, 5 g L-1; yeast extract, 10 g L-1; glucose 10 g L-1) medium, followed by incubation on

S0013-936X(96)00441-5 CCC: $14.00

 1997 American Chemical Society

Rhamnolipid production and purification have been described previously (7, 8).

FIGURE 1. Schematic of components used in column experiments. a shaker for 10 h at 37 °C. Cells were washed twice (4000g, 20 min) with 0.05 M NaCl, then suspended in 50 mL of 0.05 M NaCl, and placed on a shaker for at least 48 h. This washing procedure was used to obtain nutrient-deprived, glycocalyxfree cells in order to mimic subsurface conditions as closely as possible. Cell suspensions were prepared by diluting the stock culture into 0.05 M NaCl (OD450 ) 1.0, 109 cfu mL-1) or into mineral salts media (MSM, 12) to a final density of 107 cfu mL-1, depending on the experiment. Cell biomass was determined by protein analysis using the Lowry method (8). Establishing Residual Saturation. A 7 cm × 2.1 cm stainless steel column (Alltech, Deerfield, IL) was packed with sterilized, oven dried 40/50 mesh Accusand (North Kato Supply, Mankato, MN). The columns were wet packed by adding the sand (in 5-g increments) under a layer of electrolyte solution (0.01 M NaCl). Each increment was mixed thoroughly to dislodge any trapped air bubbles. The packed column was then saturated from the bottom up with 0.01 M NaCl at a flow rate of 4 mL h-1 for at least 12 pore volumes (24 h). Hydrodynamic properties of the column were determined using a conservative tracer (3H2O), and each column exibited the behavior expected for a homogeneously packed sand column. To establish residual hexadecane saturation, the column was first loaded top-down with 3 pore volumes of hexadecane using a syringe pump (Sage Instruments Model 352, Orion Research Inc.) at a flow rate of 4 mL h-1. Eluent from the column was collected in a 50-mL buret (buret A, Figure 1) in order to separately determine the volume of water and hexadecane eluted from the column. The amount of hexadecane retained within the column was calculated from the difference between the amount loaded (3 pore volumes) and the amount collected in the buret (corrected for the dead volume). The column was then flushed from the bottom up with 0.01 M NaCl for at least 3 pore volumes to establish the residual NAPL saturation. Flushing was done at a flow rate of 4 mL h-1 using a metering pump (Fluid Metering Inc., Oyster Bay, NY), and eluent was collected in a 50-mL buret (buret B, Figure 1). Residual saturation of hexadecane was calculated from the difference between the amount of hexadecane initially loaded into the column and the amount of hexadecane displaced during flushing. To determine the effect of bacterial biomass on hexadecane entrapment, sand-packed columns were loaded with a suspension of ATCC 15442 (109 cfu mL-1 in 0.05 M NaCl) from the bottom up at a flow rate of 4 mL h-1 for 4-5 pore volumes. The column was then washed for at least 3 pore volumes with 0.01 M NaCl to remove all but the irreversibly attached cells. Eluent fractions (1.5 mL) were collected, during both cell loading and washing, in test tubes containing 0.15 mL of 1 M NaOH and analyzed for protein content. Residual hexadecane saturation was then established as described above. Biosurfactant. A monorhamnolipid biosurfactant produced by P. aeruginosa ATCC 9027 was used in this study. The rhamnolipid has a pKa of 5.6 (13), a formula weight of 504, and a critical micelle concentration (cmc) of 0.1 mM (9).

Biodegradation and Mobilization of Residual Hexadecane in Column Studies. Column studies were used to evaluate the removal of immobilized hexadecane by microbial degradation and biosurfactant addition. In a preliminary experiment to evaluate residual hexadecane biodegradation, residual [14C]hexadecane saturation was established in a column as previously described. However, no mineralization was detected. Subsequent calculations showed that a prohibitively costly amount of [14C]hexadecane would be required to achieve a specific activity high enough to detect mineralization. Therefore, for the next series of experiments, residual hexadecane was established using only 10 µL of hexadecane. The columns were first packed with approximately 2 cm dry sand, and then 10 µL of [14C]hexadecane (75 µCi mL-1) was deposited onto the surface of the sand using a Hamilton syringe. The remainder of the column was packed with sand in 1-cm aliquots that were mixed with the [14C]hexadecane-wetted sand to distribute the [14C]hexadecane throughout the column. The packed columns were saturated from the bottom up with 0.01 M NaCl for at least 72 h at flow rates gradually increasing from 0.4 to 24 mL h-1. The effluent collected during the column saturation period was found to contain not more than 2% of the total radioactivity added to the column; the remainder was immobilized in the column. Following saturation, columns were loaded (24 mL h-1) with 20 pore volumes of ATCC 15442 cell suspension (107 cfu mL-1 in MSM) and then flushed with 200-300 pore volumes of MSM containing 0 or 0.1 mM rhamnolipid. Eluent fractions (12 mL) were collected in test tubes containing 1 mL of 1 M NaOH and assayed for radioactivity. After the release of radioactivity reached a plateau, MSM containing 1.0 mM rhamnolipid was flushed through the column for an additional 40-73 pore volumes. The radioactivity in column eluent fractions could have been from four possible sources; namely, (1) soluble or freeproduct [14C]hexadecane; (2) 14CO2 from the mineralization of [14C]hexadecane; (3) polar or (4) nonpolar undefined residues derived from 14C-labeled cellular metabolites or 14C-labeled cells. In order to distinguish between these forms of radioactivity, selected eluent fractions were analyzed using the following procedure. First, a 1-mL aliquot was removed for liquid scintillation analysis. The eluent sample was then acidified by addition of 1 mL of 50% (v/v) HCl and purged with air for 15 min to release all CO2. A second aliquot was removed, and the difference in radioactivity between the two subsamples was considered to be 14CO2. The remaining eluent was extracted 3 times with 10 mL of dichloromethane to separate nonpolar residues from polar residues. Polar residues were quantified by liquid scintillation analysis of an aliquot of the resulting aqueous sample. The loss of radioactivity following solvent extraction represented the amount of nonpolar residue. Finally, the amount of hexadecane in the nonpolar phase was determined by thin layer chromatography (TLC). To do this, the nonpolar phase was reduced to a volume of 0.1 mL using a rotoevaporator, and 10 µL was spotted onto a TLC plate (Whatman, Silica Gel AL-SIL-G). The plate was developed using n-pentane as the mobile phase (14); then each lane was sectioned into 1-cm strips and transferred into 20-mL scintillation vials, and the radioactivity was determined. A control lane was spotted with [14C]hexadecane to determine the migration of hexadecane. The proportion of [14C]hexadecane in the nonpolar residue fraction was determined from the ratio between radioactivity in the strip that should contain [14C]hexadecane and total radioactivity spotted in the lane. At the completion of each column investigation, the column contents were extracted 3 times with 50 mL of

VOL. 31, NO. 5, 1997 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

1291

TABLE 1. Physical Properties of Packed Columns and Determination of Residual Hexadecane Saturation within 40/50 Mesh Accusand-Packed Columns column characteristicsa

sterile (n ) 6)

with biomass (n ) 3)

(g/cm3)

1.77 ( 0.01 0.34 ( 0.01 8.1 ( 0.1 1.5 ( 0.4 19.0 ( 4.8

1.76 ( 0.01 0.34 ( 0.01 8.1 ( 0.1 1.6 ( 0.3 19.5 ( 3.9

bulk density porosity pore volume (mL) C16 residual (mL) C16 residual (% PV) a

Mean (standard deviation).

FIGURE 2. Transport of Pseudomonas aeruginosa ATCC 15442 through sand columns. For each breakthrough curve, a cell suspension (1 × 109 cfu mL-1 in 0.05 M NaCl) was loaded for 4-5 pore volumes, and then the column was washed with 0.01 M NaCl to remove nonadhering cells. dichloromethane, and the presence of hexadecane in the solvent extract was determined via TLC analysis. The extracted sand was then air-dried and oxidized (Biological Oxidizer OX400, Harvey Instrument Corp., Hillsdale, NJ) to determine sorbed 14C residues.

Results Hexadecane Residual in Sterile Sand Columns. Column packing characteristics revealed consistency in bulk density and pore volume between columns, with pore volumes ranging from 8.0 to 8.2 mL (Table 1). Residual hexadecane saturation was determined to be 1.5 ( 0.4 mL (19.0 ( 4.8% of the pore volume). Biomass Loading and Hexadecane Residual in Nonsterile Columns. Breakthrough of the bacterial cells was observed after 2-3.5 pore volumes of cell suspension had been flushed through the column (Figure 2). After loading the cell suspension for 4-5 pore volumes and then flushing for 3 pore volumes with 0.01 M NaCl, all but the irreversibly attached cells were washed from the column. The final cell retention ranged from 55 to 88% of the cells added. The sand core from one column (open circles) was divided into seven 1-cm sections; then duplicate samples from each section were vortexed in 1 mL of sterile water and plated to determine viable plate counts. Bacteria in each column section averaged 1.5 × 109 cfu g-1 (dry sand) and were distributed evenly along the length of the column (data not shown). The remaining columns (solid symbols) were used to determine the effect of bacterial biomass on formation of residual hexadecane. Results showed that the amount of hexadecane entrapped in the biomass-loaded columns was similar to the amount entrapped in sterile-sand columns, indicating that the biomass did not affect the immobilization of hexadecane (Table 1). Biodegradation and Mobilization of Residual Hexadecane. Three column experiments were performed to evaluate biologically-mediated removal of immobilized [14C]hexadecane. The physical properties of the sand-packed columns used in these experiments ranged from 1.74 to 1.77

1292

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 31, NO. 5, 1997

FIGURE 3. Cumulative percent release of radioactivity from sandpacked columns containing 10 µL of immobilized [14C]hexadecane. Each column was loaded with 20 pore volumes of cell suspension (1 × 107 cfu mL-1 in MSM). Column studies A (b) and B (9) then received MSM alone until the point indicated by the arrows (labeled As or Bs) at which point the columns received 1.0 mM rhamnolipid for 40 or 70 pores volumes, as indicated by the arrows labeled Af or Bf. Column C (0) received 0.1 mM rhamnolipid in mineral salts media until the arrow (Cs) at which time the solution was switched to 1.0 mM rhamnolipid for 73 pore volumes (Cf). g/cm3 for bulk density, from 0.33 to 0.34 for porosity, and from 8.1 to 8.3 mL for pore volume. Figure 3 shows the cumulative release of radioactivity from each column. Quantitation was based on the radioactivity in every fifth eluent fraction as an estimate of radioactivity within the two preceding and two subsequent fractions. Following inoculation with bacteria, columns A and B received MSM alone, and after a 50-pore volume lag period, removal of radioactivity from the column was evident. The removal of radioactivity reached a plateau after 52% (column A) and 48% (column B) of the radioactivity had been released from the column. At this plateau, a 1 mM rhamnolipid biosurfactant solution was introduced for 40 (column A) and 70 (column B) pore volumes, followed by MSM alone for at least 50 pore volumes. Following the addition of rhamnolipid, there was an immediate increase in radioactivity released from the column, which reached a plateau after several pore volumes. With rhamnolipid addition, the total amount of radioactivity released was 62% (column A) and 65% (columns B) of the radioactivity initially immobilized in each column. Column C was treated somewhat differently. A rhamnolipid solution (0.1 mM in MSM) was introduced into the column immediately following the loading of bacterial suspension. Again, a 50-pore volume lag phase was evident before the removal of radioactivity from the column began. The release of radioactivity from column C appeared to plateau after approximately 250 pore volumes, although the same clear plateau that occurred for columns A and B was not evident. At this point, 49% of the radioactivity within the column had been removed. The addition of 1.0 mM rhamnolipid (73 pore volumes) to column C increased the rate at which radioactivity was removed from the column, although not as dramatically as for columns A and B. When this study was terminated after more than 400 pore volumes, the total radioactivity released from the column accounted for 60% of the original radioactivity. As shown in Figure 4, the radioactivity in selected eluent fractions from columns B and C was analyzed and divided into four categories; namely, hexadecane free production (C16),

TABLE 2. Mass Balance of [14C]Hexadecane in Column Studies A, B, and C source of radioactivity 14C-C

16 removed during column saturation (%) 14C removed by biodegradation/ rhamnolipid treatment (%) 14C-C 16 remaining in column at completion of study (%) 14C metabolite residues remaining in column (%) total 14C recovery (%) a

A

B

C

1

2

1

62

65

60

NDa

2

2

ND

11

23

80

86

ND, not determined.

residues tightly adhering to the porous material. Results of the mass balance (Table 2) revealed radiocarbon recoveries of 80% and 86% for columns B and C, respectively. Both recoveries are less than 90%, which may be due to inefficient trapping of volatile materials such as 14CO2 that were formed during the experiment. However, the mass balance indicated that there was little hexadecane (e2%) remaining in the columns at the completion of either experiment.

Discussion FIGURE 4. Determination of the source of radioactivity in the column eluent. Bars show the relative proportions of carbon-14 labeled CO2, hexadecane (C16), nonpolar residues, and polar residues in selected column eluent fractions. The arrows indicate the point at which 1.0 mM rhamnolipid was loaded into the column (column B) or the point when the rhamnolipid concentration was increased from 0.1 to 1.0 mM (column C). Each bar represents the results for one or more combined eluent fractions collected at the approximate pore volume indicated. hexadecane mineralization product (CO2), nonpolar residues, and polar residues. The nonpolar and polar residues in the column eluent were from undefined sources, which we assume to be cells and cellular metabolites. In column study B, biodegradation of hexadecane, including mineralization and formation of cell materials and metabolites, accounted for all of the radioactivity released from the column until 1 mM rhamnolipid was added beginning at pore volume 220. After the addition of rhamnolipid, a significant portion of 14C removed in each fraction was mobilized hexadecane (1946%). Addition of rhamnolipid also caused an increase in release of 14CO2 (up to 12-fold), nonpolar residues (45-fold), and polar residues (5-fold) from the column. Results from column A were similar (data not shown). These results are in sharp contrast with results from column C, which was treated with a low level of rhamnolipid (0.1 mM) immediately following biomass loading (pore volume 20). In this case, both biodegradation and mobilization of hexadecane free product were involved in removal of radioactivity from the column. Mobilization of hexadecane declined with time, as did biodegradation. This column was then treated with 1 mM rhamnolipid (at pore volume 295), which stimulated further hexadecane removal from the column in a manner similar to that for column B. At the termination of each column experiment, the sand and the column itself were extracted with dichloromethane to recover any remaining [14C]hexadecane. Solvent extraction recovered less than 2% of radioactivity originally added to the column. TLC analysis revealed that 90% of this recovered radioactivity was hexadecane. Finally, to complete a mass balance, samples of the solvent-extracted sand were oxidized to quantify sorbed radioactivity. Results showed that up to 23% of the radioactivity was not extractable with dichloromethane. The source of this radioactivity may be biological

Measurement of NAPL Residual Saturation. This study describes a simple buret method to determine the residual saturation of hexadecane in small, saturated-flow, column experiments. The results indicated some variability (up to 25%) in the determination of residual hexadecane saturation even though column packing characteristics were quite consistent. Some variability between individually packed columns is not unexpected because packing of the porous media can result in unique pore sizes and arrangements. The residual saturation (VNAPL/Vvoids) of hexadecane in 40/50 mesh sand (0.19 ( 0.05) determined in this study was in the same range that has been reported previously for a variety of hydrocarbons or hydrocarbon mixtures in different soil types (2, 15). Effect of Biomass on Residual Saturation. The formation of residual NAPL will depend on porous medium physical properties, such as pore-size or pore-shape distribution, as well as the porous medium surface properties (1, 2). There are two ways in which bacteria could impact residual formation. First, given sufficient numbers or copious exopolymer production, bacteria could physically plug or block pores (16, 17). Alternatively, bacterial sorption to particle surfaces could change the properties of the surface with respect to charge or hydrophobicity. The results of these experiments indicate that there was no effect of bacterial biomass on hexadecane immobilization. This was true even at the levels of bacteria loaded (109 cells g-1), which is typical of surface soils, but represents a bacterial biomass that is 102-106 times greater than that reported within unsaturated subsurface environments (18, 19). Relating biomass to pore volume, it can be estimated that bacterial biomass would occupy only 0.5% of the column pore volume (assuming cell dimensions of 1 × 2 × 0.5 µm), and so physical interference of residual formation should not occur. However, even though a relatively hydrophobic bacterium (ATCC 15442) was chosen to attempt to impact the surface properties of the sand used in this study, no effect on hexadecane immobilization were observed. Biodegradation of Residual Hexadecane. Three separate column studies indicated that the [14C]hexadecane entrapped within the porous matrix was available for microbial utilization. Biodegradation alone (columns A and B, Figure 3) removed approximately 50% of the [14C]hexadecane initially immobilized within the column. This radioactivity was in the form of 14CO2 and undefined polar and nonpolar residues.

VOL. 31, NO. 5, 1997 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

1293

When a low concentration of biosurfactant (0.1 mM rhamnolipid) was present (column C, Figure 3), the removal of radioactivity from the column was the combined result of hexadecane mobilization and mineralization, although the overall rate of removal of radioactivity was not greater than what had been evident for columns A and B. When the release of radioactivity from columns A, B, and C had reached a plateau, a 1 mM biosurfactant treatment quickly stimulated further removal of radioactivity. This result was especially evident in columns A and B, where the higher biosurfactant treatment caused (1) the mobilization of hexadecane free product and (2) an increase in mineralization that lasted for several post-treatment pore volumes. Our results indicate that the removal of entrapped hexadecane reached a plateau before all of the hexadecane had been biodegraded. Under the flow-through conditions used in this study, the columns were not nutrient depleted and were also not oxygen depleted, as indicated by monitoring dissolved oxygen levels in the column effluent. A possible explanation is that a spatial separation developed between the degrading bacteria and the remaining residual hexadecane, thus limiting the bioavailability of a portion of the entrapped hexadecane. Given this explanation, the 1 mM biosurfactant treatment may have acted to redistribute the entrapped NAPL, thus increasing its bioavailability. However, only a portion of the mobilized hexadecane could be degraded before being removed from each column. The results from column C are more complicated. Initially, the low concentration of biosurfactant (0.1 mM) did not appear to increase hexadecane biodegradation but caused some mobilization of hexadecane (column C, Figures 3 and 4). These results correspond to the results of a batch biodegradation study that was performed to compare hexadecane mineralization in the presence of 0, 0.1, and 1.0 mM rhamnolipid. This study showed that significant enhancement of mineralization occurred with 1.0 mM rhamnolipid, but there was no effect of 0.1 mM rhamnolipid (data not shown). With respect to the mobilization, Bai et al. (11) showed that residual hexadecane is mobilized by rhamnolipid in a concentration dependent manner. At 0.1 mM rhamnolipid, removal by mobilization was slow and less complete in comparison to 1.0 mM rhamnolipid. Thus, in the case of column C, the combination of removal by mobilization and biodegradation was no better than for columns A and B prior to treatment with 1.0 mM rhamnolipid. Subsequent to 1.0 mM rhamnolipid treatment, redistribution of remaining entrapped hexadecane occurred in a manner similar to columns A and B, resulting in a short-term renewal of hexadecane mobilization and biodegradation. Carbon mass balance at the termination of column studies B and C showed recovery of 80 and 86% of radioactivity (Table 2). A combination of biodegradation and mobilization removed up to 65% of the [14C]hexadecane initially present. Approximately 2% of the radioactivity left in the column was [14C]hexadecane, and the remaining radioactivity within the column was 14C-labeled metabolites present as sorbed residues. Although the mass balance analysis could not account for all of the residual [14C]hexadecane, these results demonstrate efficient removal of the residual from sand columns using a combination of biodegradation and biosurfactant treatment. This point is especially apparent when considering that water flushing alone would have removed

1294

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 31, NO. 5, 1997

no more than 0.3% of the [14C]hexadecane in 400 pore volumes. It will require further study to determine the most appropriate application of biosurfactants in the remediation of residual NAPLs. Biosurfactants could be applied to mobilize a portion of the entrapped NAPL, thus reducing the time required for biodegradation. For example, Bai et al. (11) used 1 mM rhamnolipid to remove between 22% and 84% of residual hexadecane in sterile sand columns, depending on the grain size. The results of the current study showed that, in the case of a biodegradable NAPL, a large portion of the entrapped hexadecane was bioavailable and that biosurfactants could be applied following a period of biodegradation in order to increase the bioavailability of the remaining NAPL.

Acknowledgments This research was supported by the U.S. Department of Energy Subsurface Science Program, Office of Health and Environmental Research.

Literature Cited (1) Wilson, J. L.; Conrad, S. H.; Mason, W. R.; Peplinski, W.; Hagan, E. Laboratory investigation of residual liquid organics from spills, leaks, and the disposal of hazardous wastes in groundwater; U.S. Environmental Protection Agency: Washington, DC, 1990; EPA/ 600/6-90/004. (2) Mercer, J. W.; Cohen, R. M. J. Contam. Hydrol. 1990, 6, 107-163. (3) Bellin, C. A.; Rao, P. S. C. Appl. Environ. Microbiol. 1993, 59, 1813-1820. (4) McBride, J. F.; Brockman, F. J.; Szecsody, J. E.; Streile, G. P. J. Contam. Hydrol. 1992, 9, 133-154. (5) Watkinson, R. J.; Morgan, P. Biodegradation 1990, 1, 79-92. (6) Miller, R. M. In Biodegradation: Science and Application; Skipper, H. D., Turco, R. F., Eds.; SSSA Special Publication 43; Soil Science Society of America: Madison, WI, 1995; pp 33-54. (7) Zhang, Y.; Miller, R. M. Appl. Environ. Microbiol. 1992, 58, 32763282. (8) Zhang, Y.; Miller, R. M. Appl. Environ. Microbiol. 1994, 60, 21012106. (9) Zhang, Y.; Miller, R. M. Appl. Environ. Microbiol. 1995, 61, 22472251. (10) West, C. C.; Harwell, J. H. Environ. Sci. Technol. 1992, 26, 23242330. (11) Bai, G.-Y.; Brusseau, M. L.; Miller, R. M. J. Contam. Hydrol. In press. (12) Wyndham, R. C.; Costerton, J. W. Appl. Environ. Microbiol. 1981, 41, 783-790. (13) Ishigami, Y.; Gama, Y.; Nagahora, H.; Yamaguchi, M.; Nakahara, H.; Kamata, T. Chem. Lett. 1987, 5, 763-766. (14) Harvey, T., G.; Matheson, T. W.; Pratt, K. C. Anal. Chem. 1984, 56, 1277-1281. (15) Pennell, K. D.; Abriola, L. M.; Weber, W. J., Jr. Environ. Sci. Technol. 1993, 27, 2332-2340. (16) Vandevivere, P.; Baveye, P. Appl. Environ. Microbiol. 1992, 58, 16909-1698. (17) Vandevivere, P.; Baveye, P. Soil Sci. Soc. Am. J. 1992, 56, 1-13. (18) Bone, T. L.; Balkwill, D. L. Microb. Ecol. 1988, 16, 49-64. (19) Kieft, T. L.; Amy, P. S.; Brockman, F. J.; Fredrickson, J. K.; Bjornstad, B. N.; Rosacker, L. L. Microb. Ecol. 1993, 26, 59-78.

Received for review May 21, 1996. Revised manuscript received January 3, 1997. Accepted January 17, 1997.X ES960441B X

Abstract published in Advance ACS Abstracts, March 15, 1997.