Journal of Environmental Quality 31:1824-1830 (2002)
© 2002 American Society of Agronomy, Crop Science Society of America, and Soil Science Society of America
TECHNICAL REPORTS
Bioremediation and Biodegradation
Biodegradation during Contaminant Transport in Porous Media
V. The Influence of Growth and Cell Elution on Microbial Distribution
Irfan Yolcubala,
Shelley A. Pierceb,
Raina M. Maierb and
Mark L. Brusseau*,a,b
a Dep. of Hydrology and Water Resources, Univ. of Arizona, 429 Shantz, Tucson, AZ 85721
b Dep. of Soil, Water, and Environmental Science, Univ. of Arizona, 429 Shantz, Tucson, AZ 85721
* Corresponding author (brusseau{at}ag.arizona.edu)
Received for publication September 6, 2001.
 |
ABSTRACT
|
|---|
This study investigated the interaction between microbial growth and cell elution, and their influence on resultant microbial distribution between the aqueous and solid phases during solute transport in a sandy, low-organic-carbon-content porous medium. Miscible displacement experiments were conducted with salicylate as the model compound, and with different initial conditions (e.g., substrate concentrations and cell densities) to attain various degrees of microbial growth. For each experiment, salicylate and dissolved oxygen concentrations as well as cell densities were monitored in the column effluent. Cell densities were also measured in the porous medium at the beginning and end of each experiment. Total microbial growth was determined in two ways, one based on a cell mass balance for the system and the other based on total amount of salicylate degraded. For conditions yielding a considerable amount of microbial growth, the majority of the biomass was associated with the aqueous phase (68–90%). Conversely, under minimal-growth conditions, most cells (approximately 60–70%) were attached to particle surfaces. Significant cell elution was observed for most conditions, the rate of which increased in the presence of the substrate. The results suggest that the increase in aqueous-phase cells observed for the experiments exhibiting the greatest growth is associated with the production of new cells, and that under appropriate conditions aqueous-phase biomass can contribute significantly to contaminant biodegradation.
Abbreviations: CFU, colony forming units • MSB, mineral salts broth
|
INTRODUCTION
|
|---|
THE INCREASING cost and inefficiency of current remediation technologies has led to considerable interest in the use of indigenous microorganisms to remediate organic-contaminated sites (National Research Council, 1993). The level of microbial activity at a site, as well as the biodegradation potential, is influenced by numerous factors, including the type and quantity of microorganisms and their disposition in the subsurface environment. Given that the majority of microbial biomass is usually attached to solid-phase surfaces (Harvey et al., 1984; Godsy et al., 1992; Albrechtsen, 1994), it is generally assumed that aqueous-phase biomass does not contribute significantly to biodegradation. This is likely to be the prevailing condition for typical soil systems with high fractions of clay and organic matter (Murphy and Ginn, 2000). Conversely, many aquifer systems are comprised of sandy materials that have low clay and organic-matter contents. The aqueous-phase fraction of the biomass may be of greater significance in such systems. However, relatively little research has examined this question in detail.
Biodegradation processes induced by the presence of organic contaminants and attendant microbial growth may influence the distribution of biomass between the solid and aqueous phases (Harvey and Barber, 1992). Increases in aqueous-phase cell densities under microbially active conditions have been reported previously for laboratory systems (Murphy et al., 1997; Shaw and Burns, 1998; Murphy and Ginn, 2000) as well as for contaminated-aquifer field sites (Aamand et al., 1984; Harvey et al., 1984; Harvey and Barber, 1992; Godsy et al., 1992; Department of Energy, 1993). For example, a five-fold increase in biomass concentrations in ground water at the Department of Energy Savannah River site was attributed to degradation of tetrachloroethene (PCE) and trichloroethene (TCE) enhanced by the addition of oxygen and methane (Department of Energy, 1993). Harvey et al. (1984) reported that the aqueous biomass concentrations increased by an order of magnitude while soil-phase biomass concentrations remained the same in a contaminated portion of the well-known Cape Cod site. For the Cape Cod site, Harvey and Barber (1992) also reported that free-living bacteria constituted >31% of the total biomass in at least one part of the sewage-contaminated plume. They showed that the abundance of free-living bacteria correlated strongly with total dissolved organic carbon in the contaminated ground water. In a creosote-contaminated aquifer in Florida, Godsy et al. (1992) showed that up to 49% of methanogens were associated with the aqueous phase in the contaminated portion of the aquifer, whereas the microbial populations were predominantly attached to solid surfaces in the uncontaminated portion of the aquifer, similar to what was observed in the uncontaminated portion of the Cape Cod site (Harvey et al., 1984).
The results presented above indicate that aqueous-phase biomass can represent an important fraction of the total microbial population in aquifer systems. However, the relationship between the aqueous-phase biomass and biodegradation of substrates is not well understood. This study evaluates the coupled interaction of microbial growth and cell elution, and their resultant influence on the distribution of biomass between aqueous and solid phases during transport of a model hydrocarbon (salicylate) under saturated-flow conditions. In these experiments, initial conditions (e.g., substrate concentration and cell density) of the system were varied to attain various degrees of microbial growth, and for these conditions, the fractions of solid- and aqueous-phase biomass were measured by determining cell mass balances.
|
MATERIALS AND METHODS
|
|---|
Solutions and Sorbent
Salicylate (obtained from Sigma, St. Louis, MO), an intermediate compound in the degradation of naphthalene and other polycyclic aromatic hydrocarbons (PAHs), was used as a model substrate serving as the sole carbon and energy source for the organism. It is not sorbed by the porous medium used for these experiments, which eliminates bioavailability constraints (Yolcubal, 2001). Mineral salts broth (MSB) was used as the nutrient–electrolyte solution for the column experiments. The MSB contained (per liter) 1.5 g KH2PO4, 0.5 g Na2HPO4, 0.2 g MgSO4·7H2O, 2.5 g NH4Cl, 0.3 mg FeCl3, and 13.2 mg CaCl2·2H2O, and was adjusted to pH 7.0.
A well-sorted sand (20/30 mesh) with a mean particle diameter of 700 µm, an organic carbon content of 0.04%, and a cation exchange capacity of 0.57 cmol kg-1 was chosen as a porous medium for the column studies (North Kato Supply, Mankato, MN). The properties of the model porous medium are similar to those of the aquifer materials associated with the well-known Borden and Cape Cod sites. For example, Borden aquifer material (Ontario, CA) is composed of clean, well-sorted, fine- to medium-grained sand (70–690 µm), with a low clay and organic matter content (0.02%) and a cation exchange capacity of 0.52 cmol kg-1 (Mackay et al., 1986). Cape Cod aquifer material is largely composed of sand and gravel outwash (500-µm mean particle diameter) with a less than 1% clay and silt content (Le Blanc et al., 1991).
Bacterial Strain and Culture Conditions
The bacterium used in this study was Pseudomonas putida RB1353, kindly provided by Dr. Robert Burlage (Oak Ridge National Laboratories, Oak Ridge, TN). This organism contains a stable plasmid, NAH7, that encodes for naphthalene degradation. The bacterial strains were cultured and maintained in Luria Broth, which consists of (per liter) 10 g tryptone, 5 g yeast extract, and 10 g NaCl, and is adjusted to pH 7. Agar plates were made by adding 15 g L-1 Bacto-agar (Difco Laboratories, Detroit, MI) to the Luria Broth medium.
The bacterial strains were stored frozen in 12% glycerol. A frozen stock was used to inoculate a preculture for each experiment. The preculture was inoculated into a 250-mL flask containing 25 mL of Luria Broth and placed on a shaker (120 rpm) for 24 h maintained at 24°C. Growth cultures were prepared at a cell density of 105 colony forming units (CFU) mL-1 from the preculture, placed on a shaker, and allowed to grow to stationary phase. This required approximately 48 h, based on the results of a growth study (Neilson et al., 1999). A 20-mL aliquot of cell solution was taken from the growth culture, centrifuged for 10 min at 9000 rpm to pellet the cells, washed once in 20 mL of 0.85% saline, and then resuspended in 20 mL of sterile MSB solution, which resulted in a final suspension of approximately 109 CFU mL-1. Following cell harvesting, serial dilutions were prepared to obtain a desired cell density for each column experiment.
Column Experiments
Initial system conditions (i.e., substrate concentration and cell density) were varied to attain a range of microbial growth. Substrate experiments were conducted with salicylate concentrations of 5.6, 6.7, 20.7, 21.8, and 89.7 mg L-1 at an initial cell density of approximately 1 x 107 CFU g-1 dry sand and at an average pore-water velocity of 9 cm h-1 (Table 1, Experiments 3–7). Experiments were also conducted with initial cell densities of 2.8 x 106, 1.4 x 107, and 2.5 x 108 CFU g-1 dry sand. A salicylate concentration of 20 mg L-1 and an average pore-water velocity of 9 cm h-1 were used for these experiments (Table 1, Experiments 5, 8, and 9).
A stainless steel column (ModCol, St. Louis, MO) with a 5-cm i.d. and 10-cm length was used for these experiments. Column effluent was monitored for salicylate, dissolved oxygen, and cell density. The column was connected to a single-piston pump (SSI [Deerfield, IN] Accuflow Series II) with teflon tubing with a two-way valve to switch between MSB and salicylate solutions.
Prior to each experiment, a 2% bleach solution was used to sterilize the system lines. After treatment with bleach, the system lines were flushed with several pore volumes of a 0.01% sterile sodium-thiosulfate solution to neutralize the bleach. Finally, the system lines were flushed with several pore volumes of sterile deionized, distilled water. All the glassware, solutions, column, and the porous medium were sterilized by autoclaving for 30 min.
For each column experiment, 420 g of sterile sand was inoculated to a desired cell density with P. putida RB1353. The inoculated sand was mixed thoroughly under a laminar flow hood to ensure homogeneous distribution of bacteria. Three subsamples (approximately 1 g each) were collected, and cells were extracted with saline solution added at a 1:10 sand to solution ratio. Each sample was vortexed for one minute, serially diluted, and plated on Luria Broth agar in triplicate for determination of initial cell density. The columns were then packed in incremental steps with the inoculated sand under sterile conditions to obtain uniform bulk density, and saturated from the bottom for about 17 h (approximately 15 pore volumes) with sterile MSB solution at a flow rate of 1 mL min-1. The MSB and salicylate solution reservoirs were continuously sparged with oxygen during both saturation and substrate injection to avoid oxygen limitation. Following saturation of the column, a salicylate pulse was injected into the column at a flow rate of 1 mL min-1 at the concentration of interest.
Solution samples (1 mL) were periodically collected for analysis of salicylate and dissolved oxygen concentrations in the effluent. Dissolved oxygen measurements were obtained with a micro-oxygen electrode connected to an oxygen meter (Microelectrodes, Bedford, NH). One-hundred microliters of 2.75 M NaOH were added to each sample to inhibit microbial activity immediately after the dissolved oxygen measurements were taken. Solution samples were stored at 4°C until analysis. Prior to salicylate analysis, samples were centrifuged at 10 000 x g for 10 min to pellet cell debris, and then the salicylate concentration was determined with UV/VIS spectrophotometry (Shimadzu [Kyoto, Japan] UV-1601) at 231 nm. Samples collected from the reservoir prior to and after each experiment were analyzed to ensure constant initial salicylate concentration. Effluent samples were periodically plated on Luria Broth agar to monitor cell elution behavior of the P. putida RB1353 and to aid in calculating cell mass balances. After the salicylate pulse was completed, the columns were flushed with two pore volumes of MSB at a flow rate of 1 mL min-1. Finally, the column was disassembled, and subsamples of porous media (approximately 1 g) were obtained from three locations within the column (near inlet, midpoint, and near outlet) and treated as described above to determine final solid-phase cell density.
The initial total biomass was calculated as the product of the initial solid-phase cell density (CFU g-1 dry sand) and the mass of dry sand packed in the column (Table 2). The total amount of cells eluted during each experiment was calculated by integrating the area under the cell elution curve. The final amount of biomass remaining in the sand was calculated by multiplying the final solid-phase cell density by the mass of sand packed in the column. The final solid-phase cell densities were estimated by interpolating between sampling locations, assuming a linear distribution between each point. While it is possible that the actual distributions are not piece-wise linear, this assumption is probably a sufficient approximation for our case given the relatively short length of the column. This would be especially true for those systems experiencing minimal growth. As will be discussed below, the growth estimates based on cell mass balance calculations are generally similar to those based on mass of salicylate degraded, which supports the validity of this assumption for our experiments. The total microbial growth in the column was calculated by subtracting initial solid-phase biomass from the sum of the final solid-phase biomass and the total amount of cells eluted. The fractions of solid- and aqueous-phase biomass were calculated for each experiment by dividing the final solid-phase biomass or total amount of cells eluted by the final total biomass in the system (Table 2).
An estimate of total biomass produced was also calculated based on the mass of salicylate degraded (i.e., theoretical growth yield). The mass of salicylate degraded for each column experiment was calculated by subtracting the total mass of salicylate eluted from the total input mass. The estimated biomass produced was determined with a cell yield coefficient of 0.22 g cells produced per gram salicylate consumed, obtained from batch studies (Sandrin et al., 2001). This cell mass was then converted to cell number (CFU) assuming an individual cell weight of 9.5 x 10-10 mg (Neidhardt et al., 1990). The rate of increase in biomass was calculated by dividing the total number of cells produced by the total number of cells remaining in the porous medium before the initiation of the salicylate pulse (Table 3).
Control experiments were conducted with an average pore-water velocity of 9 cm h-1 and an average cell density of 2 x 107 CFU g-1 dry sand to determine if microbial growth occurred in the absence of salicylate (MSB only) (Table 1, Experiments 1 and 2). The MSB solution was continuously injected into the column for approximately 15 or 57 pore volumes. The effluent was periodically monitored for cell concentration. In addition, spatial distribution of the solid-phase cell density at the end of each experiment was determined as for the other column experiments. The control experiments were repeated to evaluate reproducibility.
|
RESULTS AND DISCUSSION
|
|---|
Microbial Growth
The degree of microbial growth for each experiment was determined based on cell mass balance as well as the mass of salicylate degraded in the column system (Tables 2 and 3). Cell mass-balance calculations for the control experiments indicated that there was no measurable increase in cell numbers in the absence of salicylate, suggesting that the contribution of MSB to microbial growth is insignificant. For the salicylate experiments, the magnitude of total microbial growth was dependent upon extant conditions. As was expected, minimal growth was observed for the experiments in which low initial salicylate concentrations were used (Experiments 3 and 4), as well as for the experiment conducted with a large initial cell density (Experiment 9). Conversely, a relatively large degree of growth was observed for the other experiments (5–8), especially for the one for which a small initial cell density was used (Table 3).
Growth estimates based on the mass of salicylate degraded were generally comparable with those obtained from cell mass-balance calculations (Table 3). For example, for the experiment for which a small initial cell density was used, the growth was calculated to be 1634% based on the cell mass balance versus 1529% based on the mass of salicylate degraded. For two of the three experiments yielding minimal total microbial growth (Experiments 4 and 9), the growth calculated from the cell mass balance was negative, compared with an approximate 10% increase based on the mass of salicylate degraded. This difference probably reflects the difficulty in measuring small increases in cell numbers with the cell mass-balance method.
Salicylate Transport Behavior
The salicylate breakthrough curves obtained for different substrate input concentrations and initial cell densities are shown in Fig. 1
. The relative salicylate concentrations for all cases reached values close to 1 within the first pore volume, indicating that microbial lag affected transport behavior. Prior experiments have shown that the delay in substrate metabolism (i.e., metabolic lag) for this system is due to the time required for induction and synthesis of appropriate enzymes required for biodegradation of salicylate (Sandrin et al., 2001).
View larger version (32K):
[in this window]
[in a new window]
|
Fig. 1. Salicylate breakthrough curves as a function of (A) input salicylate concentration and (B) initial cell density. The arrows correspond to the completion of salicylate injection. A * indicates the estimated end of the effect of lag on salicylate transport behavior, denoted by the distinct changes in the slope of salicylate breakthrough curves.
|
|
After a few pore volumes, the salicylate concentrations declined sharply as the bacterial population exited the lag phase and attained maximum biodegradation potential. As shown in Fig. 1a, the effect of lag on salicylate transport behavior ended at approximately 4, 6, and 12 pore volumes for the 5.6, 20.7, and 89.7 mg L-1 experiments, respectively. This is qualitatively denoted by the distinct change in slope of the salicylate concentration curves (Fig. 1a). These values are not necessarily an exact measure of the length of the lag phase; however, they indicate the ending of the effect of lag and the beginning of the influence of microbial growth on salicylate transport behavior.
After the effect of lag on salicylate transport behavior became insignificant, the relative salicylate concentration remained essentially constant at approximately 0.4 for the lowest Co experiment (Experiments 3, Fig. 1a). Similar behavior was also observed in the duplicate experiment (Experiment 4, data not shown). This steady state transport behavior indicates that there was minimal or no measurable microbial growth during these experiments. This is substantiated by both the cell mass-balance calculations and the growth estimates based on the salicylate degraded for the 5.6 and 6.7 mg L-1 experiments, which showed only 12 to 22% total microbial growth (Table 3, Experiments 3 and 4).
In contrast to the low-Co experiments, the salicylate concentration exhibited a moderate decline after completion of the lag-phase effects for the intermediate-Co experiments. This nonsteady transport behavior indicates there was a continued increase in substrate demand with time, due presumably to an increase in cell density (i.e., population growth). This is consistent with the microbial growth estimates, which showed increases of 166 to 219% (Table 3, Experiments 5 and 6).
For the high-Co experiment, salicylate transport was influenced by the presence of oxygen-limiting conditions despite the fact that solutions were continuously sparged with O2. The dip and rebound in salicylate concentration observed in the breakthrough curve for the Co = 89.7 mg L-1 experiment is an indication of the influence of oxygen constraints on salicylate degradation. This was verified by measuring dissolved oxygen concentrations in effluent samples, which decreased from 32 to 2 mg L-1 in approximately 12 pore volumes and remained constant thereafter. In comparison with the low-Co experiments, nonsteady transport observed in the high-Co experiment was caused by a combination of lag, growth, and dissolved oxygen limitation.
A significant period of nonsteady transport was observed for the lowest cell-density experiment. As seen in Fig. 1b, the relative salicylate concentration decreased significantly starting at approximately 15 to 20 pore volumes, following a period wherein the salicylate concentration was relatively constant. The drop in relative salicylate concentration correlates directly with an increase in the number of cells eluted (Fig. 2)
. This suggests that the increase in substrate utilization was due to microbial growth. The occurrence of significant growth for this experiment is supported by the results of the cell mass balance, which indicated a 1634% increase in cell density for the entire experiment, equivalent to a 27% increase per pore volume. Another period of quasi–steady state transport behavior occurred after approximately 30 pore volumes. This is consistent with the constant cell elution behavior observed during this period (Fig. 2), and indicates the substrate concentration was not sufficient to support continued microbial growth.
Significantly different behavior was observed for the experiment conducted with the highest initial cell density (Fig. 1b). In this case, the relative salicylate concentration decreased rapidly as the system exited the lag phase, and then leveled off at a constant concentration. The steady state behavior observed for the salicylate breakthrough curves after five to eight pore volumes indicates that there was no significant increase in substrate demand after this time. This suggests that there was minimal or no measurable net microbial growth, which is supported by both the cell mass-balance calculations and the growth estimate based on the salicylate degraded (Table 3, Experiment 9).
Cell Elution and Microbial Distribution
Cell elution was observed in all experiments. As seen in Fig. 3 , cell elution behavior during MSB saturation (-15 to 0 pore volume) was similar for all cases, indicating that cell elution is reproducible. Following the MSB saturation, cell numbers in the effluent increased for about 5 to 10 pore volumes with the initiation of the salicylate pulse and then reached a relatively constant concentration. For the smallest cell density experiment, an increase in effluent cell density continued up to 30 pore volumes (Fig. 3b). The increase in the amount of cells eluted corresponded well with an increase in total amount of microbial growth for both the different Co and cell-density experiments, suggesting that the increase in cell elution is associated with growth, rather than a hydrodynamic effect such as shearing-induced detachment. This is supported by the observed increase in cells eluted with a decrease in pore-water velocity (Yolcubal, 2001).
View larger version (34K):
[in this window]
[in a new window]
|
Fig. 3. Cell elution behavior as a function of (A) salicylate input concentration and (B) cell density. Zero pore volumes represents the initiation of the salicylate pulse.
|
|
Microbial distribution between the aqueous and solid phases in the column was determined for each experiment. Under conditions where there was minimal or no measurable total microbial growth in the system (e.g., Table 3; Experiments 1–4 and 9), the majority of the cells were associated with the solid phase. For example, in the control experiments (Table 2, Experiments 1 and 2), approximately 70% of the total cells were attached to solid surfaces in the system. Similar distributions were observed for the low-Co experiments (Table 2, Experiments 3 and 4), where the majority of cells (54–61% of the total cells in the system) were associated with the solid phase. For the largest cell-density experiment (Table 2, Experiment 9), however, cell mass-balance calculations indicated that the majority of cells in the system (79%) resided in the aqueous phase. This is probably due to the use of an initial biomass loading that was larger than the carrying capacity of the system, which resulted in rapid elution of the excess biomass. This was evidenced by the fact that a large portion (82%) of the total cells eluted were eluted prior to the initiation of the salicylate pulse. In fact, excluding the cells eluted prior to the injection of salicylate results in a distribution of 47% associated with the aqueous phase and 53% associated with the solid phase for this experiment.
In contrast to the results obtained for the minimal-growth experiments, the majority of cells were associated with the aqueous phase for the experiments conducted under conditions for which significant growth was observed (e.g., Table 2, Experiments 5–8). In these experiments, approximately 68 to 90% of the total cells were eluted from the column. The fraction of aqueous-phase biomass increased with an increase in total microbial growth. For example, 90% of the total biomass in the system resided in the aqueous phase for the experiment conducted with the small initial cell density (Experiment 8), which yielded the largest degree of growth among all experiments conducted. Given the large growth in this experiment, the results indicate that the majority of the new cells produced during population growth (cell division) were associated with the aqueous phase rather than the solid phase. These results suggest that aqueous-phase biomass can contribute considerably to the biodegradation of contaminants under conditions wherein microbial growth is significant.
|
CONCLUSIONS
|
|---|
Characterizing the processes influencing biodegradation potential under conditions relevant to solute transport in porous media is essential to developing accurate models for predicting contaminant transport and to designing successful in situ bioremediation systems and risk assessment strategies. This study investigated the interaction between microbial growth and the attendant cell elution behavior during transport of a highly soluble model hydrocarbon in a homogeneous, saturated porous medium. The results presented herein illustrate the significance of aqueous-phase biomass in a system comprised of low organic-matter and clay content. These results are consistent with observations reported for contaminated-aquifer systems under similar conditions. This work reinforces the concept that the contribution of aqueous-phase biomass to biodegradation processes should be considered in evaluating contaminant transport and remediation.
|
ACKNOWLEDGMENTS
|
|---|
This research was supported by grants provided by the USEPA Joint Bioremediation Program and the National Institute of Environmental Health Sciences Superfund Basic Research Program.
|
NOTES
|
|---|
I. Yolcubal, current address: Dep. of Geology, Kocaeli Univ., Vinsan Kampusu, 41040, Kocaeli, Turkey.
|
REFERENCES
|
|---|
- Aamand, J., C. Jorgensen, E. Arvin, and B.K. Jensen. 1984. Microbial adaption to degradation of hydrocarbons in polluted and unpolluted groundwater. J. Contam. Hydrol. 4:299–312.
- Albrechtsen, H.J. 1994. Distribution of bacteria, estimated by a viable count method, and heterotrophic activity in different size fractions of aquifer sediments. Geomicrobiol. J. 12:253–264.
- Department of Energy. 1993. Cleanup of VOCs in non-arid soils—The Savannah River Integrated Demonstration. WSRC-MS-91-290, Rev. 1. Restoration and Waste Manage. Office of Technol. Dev., Washington, DC.
- Godsy, E.M., D.F. Goerlitz, and D. Grbic-Galic. 1992. Methanogenic biodegradation of creosote contaminants in natural and simulated ground-water ecosystems. Ground Water 30:232–242.
- Harvey, R.W., and L.B. Barber II. 1992. Associations of free-living bacteria and dissolved organic compounds in a plume of contaminated groundwater. J. Contam. Hydrol. 9:91–103.
- Harvey, R.W., R.L. Smith, and L. George. 1984. Effect of organic contamination upon microbial distributions and heterotrophic uptake in a Cape Cod, Mass., aquifer. Appl. Environ. Microbiol. 48:1197–1202.[Abstract/Free Full Text]
- LeBlanc, D.R., S.P. Garabedian, K.M. Hess, L.W. Gelhar, R.D. Quadri, K.G. Stollenwerk, and W.W. Wood. 1991. Large-scale natural gradient tracer test in sand and gravel, Cape Cod, Massachusetts. 1. Experimental design and observed tracer movement. Water Resour. Res. 27:895–910.
- Mackay, D.M., D.L. Freyberg, and P.V. Roberts. 1986. A natural gradient experiment on solute transport in a sand aquifer. 1. Approach and overview of plume movement. Water Resour. Res. 22:2017–2029.
- Murphy, E.M., and T.R. Ginn. 2000. Modeling microbial processes in porous media. Hydrogeol. J. 8:142–158.
- Murphy, E.M., T.R. Ginn, A. Chilakapati, C.T. Resch, J.L. Phillips, T.W. Wietsma, and C.M. Spadoni. 1997. The influence of physical heterogeneity on microbial degradation and distribution in porous media. Water Resour. Res. 33:1087–1103.
- National Research Council. 1993. In-situ bioremediation: When does it work? National Academy Press, Washington, DC.
- Neidhardt, F.C., J.L. Ingraham, and M. Schaechter. 1990. Physiology of the bacterial cell. A molecular approach. Sinauer Assoc., Sunderland, MA.
- Neilson, J.W., S.A. Pierce, and R.M. Maier. 1999. Factors influencing the expression of luxCDABE and NAH7 genes in Pseuodomonas putida RB1353 (NAH7, pUTK9). Appl. Environ. Microbiol. 65:3473–3482.[Abstract/Free Full Text]
- Sandrin, S.K., F.L. Jordan, R.M. Maier, and M.L. Brusseau. 2001. Biodegradation during contaminant transport in porous media: 4. Impact of microbial lag and bacterial cell growth. J. Contam. Hydrol. 50:225–242.[Web of Science][Medline]
- Shaw, L.J., and R.G. Burns. 1998. Biodegradation–transport interactions of pulse applied 2,4-D in repacked soil column microcosms. J. Environ. Qual. 27:1472–1478.[Abstract/Free Full Text]
- Yolcubal, I. 2001. In-situ monitoring of microbial activity and biodegradation during solute transport in porous media. Ph.D. thesis. Univ. of Arizona, Tucson.
TOP
ABSTRACT
INTRODUCTION
