In-Situ Biodegradation of Toluene in a Contaminated Stream. 2

In-Situ Biodegradation of Toluene in a Contaminated Stream. Part 1. Field Studies. Heekyung. Kim , Harold F. Hemond , Lee R. Krumholz , and Brian A. C...
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Environ. Sci. Techno/. 1995, 29, 117-125

hSitu Biodegradation of Toluene in a Contaminated Stream. 2. Laboratory Studies BRIAN A. C O H E N , t LEE R. KRUMHOLZ,*c* HEEKYUNG KIM, AND HAROLD F. HEMOND Ralph M. Parsons Laboratory of Water Resources and Hydrodynamics, Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139

In-situbiodegradation ratesfor a stream contaminated with toluene (95%) compartmental contributions. Rates determined in batch studies under winter conditions were 11-14% of rates determined under summer conditions. Batch studies of mineralization rates determined with [U-14C]toluene indicate that mineralization rates were 23% of biodegradation rates. The remaining toluene was converted to soluble intermediates (15%), and 62% was taken up as biomass or converted to other insoluble material. Our results support the assertion that biodegradation was the most important environmental sink for toluene, with 70% of the toluene loss due to biodegradation over the studied stream reach.

Introduction In many cases of release of xenobiotic chemicals into the environment, in-situbiodegradationby naturally occurring microorganisms is thought to play an important role in attenuating contamination. However, because of the complexity of the natural environment, the extent of these in-situprocesses is extremely hard to verify (1). Commonly, biodegradation rates are measured in laboratory microcosms; however, extrapolation of such results to the field is uncertain because of the many environmental factors (nutrients, oxygen levels, temperature, turbulence, adaptation, etc.) that are not necessarily reproduced under laboratory conditions. Comparisons between microcosm rates and actual in-situ biodegradation rates are needed to reduce these uncertainties. In this paper, laboratory experiments are presented, which include determinations of in-situ biodegradation rates of toluene in a contaminated stream. The laboratory studies are compared to the field studies on the same stream presented in the accompanying paper (2). This work provides additional evidence for in-situbiodegradationand validates an experimental approach for determining insitu biodegradation rates in the laboratory. We have also determined mineralization rates of toluene and bacterial density in different seasons in order to understand the biodegradation of toluene in the stream in more detail. Our objectives are to gain an understanding of the utility of microcosm studies in predicting in-situ biodegradation rates, to estimate the fate of toluene in a stream, and to understand the factors that may affect the biodegradation rates.

Background Biodegradation rates in streams can be expressed in a number of ways, but the most useful expressionsare those related to relevant biological and hydrological parameters such as streambed surface area, flow rates, and microbial biomass (3). It has been shown that biodegradation of a contaminant in a shallow stream is mainly accomplished by attached rather than suspended bacteria (4-9). In this studied stream, attached bacteriawerefound to be primarily responsible for toluene biodegradationbased on the results presented in this paper. In these cases, a useful formulation for expressing the biodegradation rate of toluene by attached bacteria in the studied stream is as follows (7):

where k b is the biodegradation rate constant (mlh),Cis the concentration of the toluene in the stream (mglm3 or pgl L), and (A,Iv) (m-9 is the ratio of the surface area (A,) to the volume of streamwater (v). In the accompanying paper (21,AJVwas taken as the stream depth. In this work, we employed two methods to * Corresponding author;



k”[email protected]

+ Present address: Environmental Working Group, 718 Conneticut Avenue NW, Suite 600, Washington, D.C. 20009. Present address: Department of Botany and Microbiology, 770 Van Vleet Oval, University of Oklahoma, Norman, OK 73019.



D 1994 American Chemical Society




Sampling Port

Pmrlstallic Pump

I l l Rmsonolr

Tmflon Bag

Porirtoltlc Pump

FIGURE 1. Diagram of flow-through column systems. (A) System used for experiments with unlabeled toluene. Reservoir consists of autoclaved East Drainage Ditch water continuously sparged with air. Flow rates through peristaltic pump were 3-5 ml/min. Toluene is injected from a stock solution in a syringe pump at rates ranging from 0 2 to 2 mVh. Mixing chamber is a 1 2 5 " Erlenmeyer flask. Column is 34 cm long, with a 5 cm inner diameter. Sampling ports had Luer lock fittings, and samples ware withdrawn using glass syringes. (5)Column system used with P4C]toluene. Toluene and filtered streamwater were premixed in Tedlar gas bags at a known concentration before being pumped into column. All other conditions were as described above.

estimate A,/ V; one was to measure the surface area of rock or sediment samples and the water volume in a flowthrough column setup, and the other was to measure the mass of sediment or rock samples and water volume in a batch system and then convert the mass of sediment or rock samples into a surface area which might contact the streamwater. In addition to environmental factors which may influence the biodegradation rates of both attached and suspended bacteria, mass transport of the contaminant to the attached bacteria can limit the biodegradation rates (4). Different microcosm systems will exert different types of limitation on the attached bacteria. For example, Pignatello et al. (9) observed that rock surfaces showed higher biodegradation rates of pentachlorophenol than sediment surfaces in their model stream. In this work, we determined biodegradation rate constants (kb) on different surfaces (rock, plant, sediment) using two microcosm systems. Because of rapid mixing, the batch system represents a system with little mass transport limitations, while the column system has a greater mass transport limitation.

Experimental Section Study Site. Field work and sample collection were done at the East Drainage Ditch, a small stream in Wilmington and Woburn, MA. Further details on this site are presented 118 ENVIRONMENTAL SCIENCE &TECHNOLOGY / VOL. 29, NO. 1,1995

in the accompanying paper (2). Samples were collected at an upstream location of the studied reach between the first and second sampling stations of the field study (2). SampleCollection. Rock, sediment, and plant samples collected at various times throughout the year were stored at ambient temperatures in polypropylene bottles for up to 1 h. Samples were then stored in a refrigerator at 4 "C for a maximum of 48 h before use. Biodegradation rates between freshly collected samples and samples stored for a 48-h period at 4 "C were similar (data not shown). Sediments were collected from the top 5 mm of the surface sediments, and rocks were collected from the top of the sediment surface. Plant samples were collected from a 1 m long section of dense growth of submerged plants approximately 80 m downstream from the beginning of the studied reach. These plant samples were of the species most prevalent in this stream reach. Flow-ThroughColumn Setup. Flow-through columns were set up as illustrated in Figure 1 with only a small headspace ('5% ofthe totalvolume). For columnsinwhich a.mixingchamber was used, toluene was pumped into the mixing chamber from a saturated solution with a syringe pump, and streamwater was pumped into the mixing chamber from a reservoir using a peristaltic pump. In later studies using isotopes, streamwater and solvents were premixed in Tedlar gas bags and pumped directly into the column with a peristaltic pump. All glassware and tubing in contact with streamwater were autoclaved before

experiments were begun. Sediments columns were set up horizontally as shown in Figure 1 so that flow was over a bed of sediments,while rock columnswere placed vertically. Columns were full of rocks or about half fullof sediments. Flow rates varied from 2.1 x to 2.7 x m31h,giving velocities of 0.06-0.10 mlh. These velocities, although significantly slower than those observed in the field, were the fastest that we could use while still observing a noticeable biodegradation of toluene during the course of the experiment. For sediment, surfacelvolume ratios (AJV in eq 1)were determined by first measuring the surface area of sediments, that is length of column multiplied by width of sediment layer, and the volume of water in the column containing sediments. For rocks, the surface area was estimated by measuring the sides of individual rocks with a ruler. This was possible as the sides of the rocks were relatively flat. The mass of the rock was then divided by the surface area; the rock masslsurface area ratio averaged 12 kg of rock/ m2. Rocks all had similar diameters and densities, and all the surfaces of rocks in the column were exposed to the flowing water. In every experiment, surfacelvolume ratios for rocks were determined by measuring the volume of water in the column and the mass of rocks used and then converting to the surface area. The ratios were approximately 70 m-l for rocks and 25 m-l for sediments. Column Study: Biodegradation Rates. Toluene concentrations in influent to the columns varied from 125 to 2000pglL. Retention time in columns was 3.5-4.5 h. The column were run for approximately 6-10 h at each concentration, with one retention time allowed for the column to reach equilibrium, followed by at least 2-h influent sample collections. After an additional retention time, two effluent samples were collected over a 1-hperiod. Concentrations in columns were changed monotonically upward. In control (autoclaved) columns, the difference between top and bottom concentrations did not change based on visual observation of the data after 2 or 3 h of an equilibration period at each concentration. The total time for each experiment did not exceed 24 h. Duplicate 20-mL samples were collected with a syringe from the valves near the top and the bottom of the column. These samples were added to 40 mL of EPA screw-top vials with Teflon-coatedsilicone septa. Toluene concentrations were then determined by headspace analysis as described in a later section. Control columns were run in a similar fashion, but with autoclaved sediments and streamwater. Biodegradation rates were determined by the difference between toluene concentration at the top and bottom of the column normalized to residence time and rock or sediment surface arealwater volume ratios in the column. Rates are expressed as mg of toluene (m of rockY h-l. An abiotic loss rate was determined in a similar fashion with autoclaved samples. Column Study Mineralization Rates. Mineralization rates in flow-through columns were determined using [U-14C]toluene(Sigma Chemicals, St. Louis, MO). Toluene concentrations in the column influent ranged from 80 to 300pg/L, withan activityof0.4-0.8pCilL. Eachexperiment was performed at a single toluene concentration. After flow was begun and the column was allowed 1-2 h to equilibrate, duplicate 50-mL samples were collected from the top and bottom sample ports of the column and placed in 100-mL serum bottles with Teflon-coated butyl rubber

stoppers. Residence time in the columns was between 2 and 5 h. 14C02analysis was performed by first acidifying samples and then suspending a small wire basket at the stopper in each serum bottle, which held a piece of filter paper (Whatman No. 1, 2-cm diameter) treated with phenethylamine as a COz trap (10). 14C02trapping efficiency was 100% as determined with NaH14C03. Total 14C02evolution was determined as the difference between the amount of 14Ctrapped from the influent sample and the amount of 14Ctrapped from the effluent sample. Approximately 10% of 14C activity was recovered from iduent (control)samples before biodegradation had begun. This was thought to be largelydue to [l4C1toluene volatilizing during the equilibration period and partitioning onto the phenethylamine-treated filters. The amount of 14Crecovered in effluent samples was corrected for this partitioning to determine mineralization rates. Mineralization rates in the columns were calculated from 14C02concentrations in the same way as biodegradation rates. These isotopic studies were also used as a control to determine abiotic loss rates in columns. Total 14Crecovery was determined in these columns, and abiotic losses were determined by measuring the difference between 14C activity in the influent and the effluent of the column. Batch Study: Biodegradation Rates. For biodegradation rate determinations, assays were performed in 500mL round-bottom flasks each fitted with a glass stopcock. Flasks contained 250 mL of streamwater and either 5-10 g of fresh sediments, 35-70 g of rocks, or 1-2 g of plant material. Flasks were shaken on awrist action shaker during the assays. Toluene disappearance over time was then determined by headspace analysis (see below). All assays were run in duplicate, with one additional flask either autoclaved or poisoned with 15 mglL HgC12 to serve as controls. These two sterilization techniques gave similar results. Assays performed in the summer and fall were run at 22 "C, and all assays run in winter were run at 5 "C. Toluene and other solvents (trichloroethylene (TCE), chloroform or chlorobenzene) were added from saturated solutions to a final concentration ranging from 100 to 1000 pg/L. Other solvents were added to determine the effects of a mixture of contaminants on biodegradation rates of toluene. No detectable biodegradation was observed in flasks incubated with only streamwater and solvents. Biodegradation rates were determined as initial degradation rates based on the initial slope ofthe concentration vs time curve. This was measured at the point where degradation was observed to begin. In cases where concentrations of toluene was above 390pglL, 1-4 h lags were observed before degradation began, and these were not included in calculations. Slopes of live samples were corrected for abiotic losses in killed controls. Biodegradation rates were observed to be proportional to the mass of rocks or sedments in the flask (data not shown). Biodegradation rates in batch studies were expressed as mg of toluene (kg of sediment or rocks)-' h-l. Batch Study Mineralization Rates. Mineralization rates of [14C]toluene to 14C02were also determined in batch cultures. Two grams of wet sediment were added to 20 mL of streamwaterin 100-mLserum bottles with Teflon-coated butyl rubber stoppers. l4CO2analysis was performed as previously described in an earlier section. Either 20 000 counts per minute (cpm) (250pglLsamples) or 10 000 cpm (100 pglL samples) [U-l4C1toluenewas added to reaction vessels, and unlabeledtoluene was added to bring the final VOL. 29, NO. 1, 1995 / ENVIRONMENTAL SCIENCE & TECHNOLOGY


concentration to either 100 pg/L or 250 pg/L. Identical autoclaved controls were prepared at the two different concentrations. At 2 h intervals, duplicates were killed by the addition of HzS04 to pH 2.0, and the amount of I4CO2produced was determined as described above. Mineralization rates were determined from the slope of l4CO2production vs time and were corrected for radioactivity measured in control samples. In parallel to the batch mineralization studies, a set of serum bottles was set up in the identical fashion with unlabeled toluene. These were analyzed for biodegradation via toluene loss as described below. Determination of Toluene Metabolites. In order to determine the ultimate microbial fate of toluene, an attempt was made to determine the amount of 14Cremaining in solution as metabolites. Samples of 250 pglL toluene, including [l4C1toluene,were incubated for 3-24 h until toluene levels were below detection limits (15pglL). The mixture was acidified to allow COZto escape, and samples were centrifuged at 12 000 rpm for 30 min. Total 14C remaining in solution was determined by liquid scintillation counting. Headspace Analysis of Toluene and Other Solvents. Sampleswere analyzed for the disappearance of toluene or other solvents (TCE, chloroform, and chlorobenzene) over time. Analysis was performed by injecting a 1-mL headspace sample withdrawn from serum bottles or flasks,using a gas-tight syringe, to a gas chromatograph [Carlo Erba HRGC 5300 Mega series gas chromatograph (Fisons Instruments, Valencia, CA)] fitted with a Restek RTX-5 capillary column and a flame ionization detector. Oven temperature was held at 40 "C for 2 min and increased to 150 "C by 10 "C/min. Detection limits were 200 pglL for chloroform and TCE and 15 pg/L for chlorobenzene. Calculation of Stream Biodegradation Rate Constant. The stream biodegradation rate constant was calculated from batch and column studies of rock and sediments based on the composition of streambed surface. Streambed characterization was performed in the summer months. Observationswere made every3 m, startingat the beginning of the studied reach and continuing for 100 m. The composition of the streambed (percentage covered by rocks or sediments) was determined by visual estimation as 85% sediment and 15% rocks. To be consistent with the way the field rate constant was calculated,the rate constants in laboratory studies were obtained by assuming that the biodegradation rates were first-order with respect to toluene concentration. Solving eq 1 for Cyields

where kl is a first-order biodegradation rate constant (h-l), C,, is the initial concentration of toluene, and C, is the toluene concentration after a period of time, z. In batch studies, the first-order biodegradation rate constant (kl) was taken as the slope from natural logarithm of concentration vs time curves at the initial concentrations of 100pglL. The range of initial concentrations chosen for the calculation was based on the finding that biodegradation rates were first-order with respect to toluene concentration below 2OOpglL as discussed in the Discussion section. The first-order biodegradation rate constant then was divided 120


by the mass of sediment or rocks used (kg of rocks or sediment) and multiplied by the water volume (m3of water) in the flasks, resulting in the biodegradation rate constant in units of m3 (kg of sediment or rock)-' h-l. This biodegradation rate constant was converted to a rate constant expressed as m/h (kb)by being multiplied by the masslsurface area ratio for rocks or sediments. For rocks, the mean masslsurface area ratio was 12 kg of rocks/m2, determined as described previously. For sediments, the masslsurface area ratio was obtained on the assumption that the aerobic toluene-degrading population was contained in the top sediment from streambed surface to the depth of 0.2-0.5 cm and that the capacity for toluene degradation is homogeneous throughout the depth. This depth is based first on the findings of Pignatello et al. ( I ] ) , who found that the aerobic degradation zone in a PCPcontaminated stream extended to a depth of 0.5 cm. Visual inspection of East Drainage Ditch sediments indicated that the sandy, lighter colored sediments that were indicative of the aerobic zone extended to this depth. Several in-situ measurements of oxygen profiles of the sediments in this stream, using an oxygen microelectrode, showed that the aerobic depth in this site was in the range of 0.2-0.5 cm (data not shown). Using these assumptions, the mass/ surface area ratio for sediment was obtained as the value in the range of 4-10 kg of sediment/m*by multiplying the wet sediment density (2 g/cm3) by the depth of the active zone (0.2-0.5 cm). In column studies, the biodegradation rate constant (kb) was obtained from eq 2 by substituting C, with the concentration of influent, C, with the concentration of effluent of the column, and z with the residence time in the column. Loss rate constants obtained from control columns were subtracted from rate constants of live columns. Determination of Biomass and Activity. Samples were frozen, stored for up to 3 months, and later analyzed to determine biomass and fluorescein diacetate hydrolysis (FDA) activity. For cell counts, cells were dislodged from sediments by treatment with a surfactant followed by sonication (12). Cells were then counted by DAPI staining under an epifluorescence microscope. At least 20 grids and 300 cells were counted for each sample. All samples were run in duplicate, and cells attached to particles that were visualized were counted twice, assumingthat the same number of cells would be attached and uncountable underneath the particle. Microbial activity determinations were based upon fluorescein diacetate hydrolysis rates (13). Assays were performed in 100-mL plastic capped vials with 5 g of sediment, 30 g of rocks, or 1 g of plants. Vials were shaken on a wrist action shaker during the incubations. FDA hydrolysis rates were observed to be linear with both biomass and time under the experimental conditions.

Results Biodegradation Rates: Column Studies. For columns with rocks, biodegradation rates appeared to be mixed or first order with respect to concentration at concentrations up to 640pglL (Figure 2B). Abiotic losses as determined from control columns account for 37% of total losses. For sediments, toluene biodegradation rates in live columns appeared to be mixed or first order with respect to concentration at concentrations up to 2200 pg/L (Figure 2A). The mixed-order system exists because microorganisms in the deeper sediment may be exposed to a lower


l2 lo

c -

1 5 -

8 6


Sediment in Batch 10-


- i

2 0 .&


-21 0














B. 2.0,





Batch 6


Toluene Concentration (pg/L)









I l


Toluene Concentration (pg/L)

FIGURE 3. Toluene biodegradation rates in batch cultures of rocks and sediments. Rates of toluene loss in shake-flasks with sbeemwater are detemined by the disappearance of toluene as measured by gas chromatography. Error bars represent standard deviations of duplicate samples. TABLE 1

Comparison of Toluene Biodegradation Rates for Batch Studies in Winter and Summer with Contaminant Mixtures assay type'

-0.5 0







Toluene Concentration (pg/L)

FIGURE 2 Toluene biodegradation rates in sediment (A) and rock (B)columns. Rates of toluene loss in columns with streamwater and sediment were determinedas described in the ExperimentalSection. Error bars represent standard deviations of at least three samples from the top and bottom of columns. Control column data was collected from two differenttypesof controls: an autoclaved control analyzed for toluene disappearance by GC and a live control in which a mass balance was performed with ["Cltoluene.

level of toluene than microorganisms in the surface sediment. Abiotic losses account for 32% of total losses. Biodegradation Rates: Batch Studies. Toluene biodegradation rates were determined under summer conditions (22 "C) for concentrations between 110 and llOOpg/L (Figure 3). These rates appeared to be first order with respect to concentration over the concentrations up to 200 pg/L. In contrast to column results, however, biodegradation rates in batch appear to approach zero order with respect to concentrations higher than 400 pg/L. For concentrationsbetween 0 and 25Opg/L, no lag was observed before biodegradation began. At higher concentrations, lags of between 2.7 and 4.5 h were observed. V,, for sediments was 1.4 mg of toluene (kg of sediment)-' h-l (6-14 mg of toluene (m2 of sediment surface)-'h-l); for rocks, Vm,was 1.5 x lo-' mgoftoluene (kg of rock)-' h-l (1.8 mg of toluene (m2of rock surface)-' h-l. These results indicate that at concentrations in excess of 400 pg/L, biodegradation rates dependent only upon surface arealvolume ratios would be observed: dC/dt = Vm,(A,/ V)

where V,,


(mg of toluene m-2 h-'1 is the maximum

condition summer summer, mixtureb summer, with CBc winter winter, mixture

sediment [mg of toluene Ikg of sediment)-l h-ll 1.6 f 0.23 1.5 f 1.2 0.23 0.16

f 0.13 f 0.06

rock [mg of toluene Ikg of rock)-' h-'1

0.15 f 0.018 0.068 f 0.013 0.03€Id 0.017 f 0.007 0.026 f 0.017

a Results are from shake-flask assays at toluene concentrations of 250pg/L. Summer assays were performed at 22 "C in July and August and winter assays at 5 "C in January and February. Errors represent standard deviations of multiple samples. Mixture consists of 1000 pg/L each of chloroform and trichloroethylene and l0OpgR chlorobenzene. CB = sample with chlorobenzene and toluene, 100 pg/L each. Standard deviation unavailable; result is from one sample.

biodegradationrate observed for microorganisms attached to stream surfaces. In order to determine seasonal factors affecting biodegradation rates, batch studies were performed with samples collected in winter and incubated at typical ambient (5 "C) temperatures. These studies were run only at concentrations of 250 pglL. These assays indicated significantlyslower (10%-33%) biodegradation rates than samples under summer conditions (Table 1). In addition, assays at toluene concentrations of250pglL were performed in summer months at 22 "C for rooted submerged macrophytes and for streamwater with no amendments. Microorganisms attached to macrophytes effected a toluene biodegradation rate of 26 & 10 mg of toluene (kg of plant material-' h-l, order of magnitude higher than sediments on a per gram basis. For samples in which toluene was added to 250 mL of streamwater with no sediments, rocks, or plants, no biodegradation was observed over a 24-h period. VOL. 29, NO. 1, 1995 / ENVIRONMENTAL SCIENCE i 3 TECHNOLOGY





I 2.5



Control Toluene








Time (hour)

FIGURE 4. Comparison of biodegradation and mineralization rates by stream sediments. Assays were performed at 22 "C in 100-mL serum bottles containing 30 mL of sneamwater and 2 g of sediment. Closed squares represent biodegradation.determined by measuring toluene disappearance by GC in replicatesamples. Closed triangles representmineralization rates, determined by evolution of"C02from ['%]toluene, and are plotted as the concentration of toluene mineralized. Open triangles and squares represent autoclaved controls. Error bars represent standard deviation of duplicates.

Degradation rates were also determined for samples where toluene was added with a suite of other organic compounds [chloroform(1000pglL). TCE (lOOOpg/L),and chlorobenzene (100 pg/L)]. In all samples to which the organic mixture was added, chlorobenzene was observed to degrade to below detection limits. In samples in which chlorobenzene was added with no toluene present, no chlorobenzene biodegradation was observed. No significant biodegradation of TCE or chloroform was observed. Although the latter information is not surprising, it is interesting that toluene was required for the induction of the chlorobenzene degradation system in the resident microflora. Mineralization Rate: ColumnStudies. Mineralization rates of toluene by rock and sediment samples were also determined in columns using ['4Cltoluene. Mineralization rates were observed to be first order with respect to concentration. Mineralization rates for rock columns are 2.7 x mg of toluene (m2of rock surface)-' h-' at the influent concentration of 280 pg/L and 1.7 x mg of toluene (m2of rock surface)-' h-' at the influent concentrationof 150pg/L. Forsedimentcolumns, amineralization rate of 1.6 x IO-' mg oftoluene (m2of sediment surface)-l h-I was observed at the influent concentration of2lOpglL and a rate of 8.6 x mg of toluene (m2 of sediment surface)-' h-' was observed at the influent concentration of88pglL. Bytaking the slope ofthe plots ofmineralization rate vs influent concentration, the mineralization rate constants were determined to be 1.1 x m/h for rocks and 8.4 x m/h for sediments. Mineralization Rate: Batch Studies. Toluene mineralization rates for stream sediments were determined by measuring 14C02 evolution from [U-14C]toluene. Mineralization rates were found to be first order with respect to toluene concentration at concentrations of 100 and 230 pg/L. Mineralization rates were not determined at higher concentrations. A plot comparing biodegradation to mineralization at the initial concentration of 230 pg/L is presented in Figure 4, where biodegradation rate is 0.1 mg 122 m ENVIRONMENTAL SCIENCE &TECHNOLOGY I VOL. 29. NO. 1.1995

Dale RGURE5. Seasonal variations of microbial activity and cell numbers. Microbial activily results are based upon fluorescein diacetate hydrolysis rates in sediment samples. Cell counts are based upon direct counts of OAPl stained sediment samples pretreated with Tween 80 and sonicated to dislodge anached microorganisms. Error bars represent standard deviations of duplicate assays.

of toluene (kg of sediment)-' h-' and mineralization rate is 2.5 x mg of toluene (kg of sediment)-l h-'. Mineralization is observed to beginwithno lag after toluene is added to samples. After centrifugation and evolution of 14C02,15%of the isotope remained in solution as dissolved intermediates at the time that the toluene had degraded to below detection limits. Seasonal Variation of Biomas and Activity. Seasonal variations of microbial biomass were determined hy cell counts oforganisms attached to sediment duringsampling periods throughout the year. Cell densities varied 5-fold over the course of the year, with maximum cell densities occurring inlate spring andremainingat highlevelsthrough the summer, and minimum cell densities occurring in late winter (Figure 5). Counts of cells attached to rocks in June indicated a microbial cell density of (3.5 f 1.1)x lo6 cells/ cm2 of rock surface. Counts of plant surfaces indicated a cell density of 2.9 x 108 cells/g. Fluorescein diacetate hydrolysis activity also exhibits a seasonal dependence (Figure5). FDAhydrolysis ratesvary by a factor of 3.5. Rates are high throughout the summer and into the fall and reach a peak in October. These rates are at a minimum in February, remaining low through the winter, and returning to higher levels in the late spring. FDA hydrolysis measurements of rocks in June have an activity of (1.8 0.2) x absorbance units (cm2of rock surface)-' h-1 while activity measurements of plants indicate an activity of (2.9 i0.07 x lo-' absorbance units (cm2of plant surface)-' h-l. No detectable activity was observed with 10 mL of streamwater alone after 16 h of incubation.


Discussion Zn-Situ Biodegradation Rate Constant: Comparison of Laboratory, Field, and Column Studies. During the summer, rate constants based on field data ranged from 0.08 to 0.40 mlh, averaging 0.21 f 0.12 mlh (2). The rate constant calculated using data from column studies was 3.9 x mlh, 2 orders of magnitude lower than the rate constant determined in field studies (Table 2). The rate constant determined with batch cultures at a similar temperature ranged from 0.06 to 0.16 m/h depending on the depth of active zone assumed, comparahle to in-situ


Biodegradation Rate Coustants DeterminBd with Batch, Column, and lnSifu Mass Balance Studies biodegradation rate constant kb (m/h) surface type





7.4 x 10-2-1.9 x lo-’ 9.7 10-3 6.4 x 10-2-1.6 x 10”

4.3 x 1.4 10-3 3.9 x

2.1 x IO-’

rocks streamb

a Ref 2. Rate constant is the average summer imsitu biodegradation rate constant. *Rate constants calculated assuming a stream bottom made up of 85% sediments and 15% rocks.

biodegradation rate constants determined in field studies (Table 2). It should be noted that there are some uncertainties in converting the batch rate constant to the field rate constant. The deeper portion of sediment probably does not degrade toluene as rapidly as the top portion of sediment in the stream. Nonetheless, our studies showed that the system of shake-flaskswas a reasonable microcosm to represent this fast-flowing stream. Batch studies in which flasks are constantly shaking are models of physical systems where mass transport of substrate to the degrading organisms is not likely to be a limiting factor, while column studies, with slow flow rates and no shaking, are models of a physical system where mass transport may be limiting. In an effort to quantify the extent of the mass transport limitation, Reynolds numbers (Re),a measure of the turbulence within a system, were calculated for the stream and the column (14):

Re = ULJv


where U is the flow velocity, L, is the characteristic length, and Y is the kinematicviscosityofwater. The characteristic length for streams is the hydraulic radius (13,and that for columns is the diameter of the column. The Reynolds number was 94 000 for the stream, a range where turbulent flow occurs. Reynolds numbers for the columns were 2, indicating nonturbulent flow. The shake-flasks are, by inspection, a fully turbulent system. These results indicate that mass transport to the sediment surface is probably not a factor-limitingtoluene biodegradation in the stream but may be limiting in the laboratory columns. Column and batch studies indicate significantly different behavior with regard to biodegradation kinetics as well. Biodegradation rates in the column although not proportional to toluene concentration over the entire range of concentrations presented increased up to the highest concentration tested, while batch studies with both sediments and rocks exhibit biphasic behavior, with first-order (substrate limited) biodegradation rates at concentrations up to 200 pglL and zero-order rates that are independent of concentration above 400,uglL. The biodegradation rate constant in column studies with rocks is 14% of the rate constant in batch studies, while that in the column with sediments is 2% of the rate constant in batch (Table 2). The lower biodegradation rates and the fact that no V , was observed in column studies are likely due to the differences in turbulent mass transport. Previous studies of herbicide biodegradation by stream bacteria indicated that laboratory microcosms exhibited increased biodegradation rates as stirring speeds of these microcosms were increased (16). The relatively high mass/volume ratios, slow flow rates, and lack of turbulence in columns means

that zones of decreased toluene concentration may form at the waterldegradation surface, causing biodegradation rates to be partly controlled by diffusive mass transport. Using the data from our batch studies and our cell counts, we calculate a V,, O f 5 x 10-l2mg of toluene cell-’ h-I for sediments and a Vm, of 5 x lo-” mg of toluene cell-’ h-’ for bacteria on rocks. These rates are significantly lower than the range of 2 x 10-2 to 5 x io-5 mg of toluene cell-’ h-l reported by Robertson and Button (17) for a variety of pure marine cultures. This difference is expected, given that our estimates include all cells counted. Not all of these cells are toluene degraders, and not all are metabolically active. Moreover, in our batch cultures there are more than one species of organism capable of toluene biodegradation (Tay, personal communication). Thus, the observed biodegradation rates are probably due to a combination of different organisms, each with different kinetic parameters for toluene biodegradation. The rate for bacteria on rocks is an order of magnitude higher than the rate for bacteria on sediment on a per unit cell basis, presumably because of their greater exposure to oxygen and toluene than experienced by bacteria in the sediments. Component Contributions to Biodegradation Rates. On a per unit area basis, the biodegradation rate constant with rocks is 4% (batch)or33% (column)of the rate constant with sediments (Table 2). Biodegradationrates in the water column are negligible. Thus, because of their relatively high biodegradation rates and the large surface area (85%) that they cover, sediment-dwelling bacteria appear to be responsible for the majority (’90%) of biodegradation in the stream. Bacteria attached to rocks are responsible for significantlylessdegradation. Although they have a higher degradation rate on a per cell basis than sediment, because of their smaller cell number per unit surface area and the smaller surface area of rocks, they are only responsible for a small fraction of the biodegradation. On a per mass basis, biodegradation rates of plants are an order of magnitude higher than rates of sediments; however, they are not responsible for most biodegradation because of their small surface area (1%of stream surface) and short growing season. Plants emerge in May or June and have died back by the end of August. This may account for part of the difference ( ~ 1 0 %between ) winter and summer rates. Environmental Fate of Toluene. Results from the field studies (2)indicate that volatilization and biodegradation account approximately for 30% and 70%, respectively, of total toluene losses from the studied section of the East Drainage Ditch. Abiotic loss in the column studies accounted for 35% of total loss. It is likely that sorption of toluene into sediment was the primary abiotic sink; batch studies showed negligible abiotic losses. This latter information is further evidence that batch studies better reflect field conditions as sorption was not an important sink in both batch and field systems. Evolution of 14C02 from batch cultures indicates that mineralization rates are approximately23% of biodegradation rates, while measured mineralization rates in columns range from 12% (rocks) to 25% (sediments) of biodegradation rates. Comparingbiodegradation rates in batch studies to mineralization rates leaves 77% of the biotransformed toluene unaccounted for. Since 15% of the initial [14C]toluene was converted to soluble intermediates, approximately 62% of the initial toluene concentration has presumably been converted to biomass or insoluble intermediates. VOL. 29, NO. 1, 1995 /ENVIRONMENTAL SCIENCE &TECHNOLOGY



Environmental Fate of Toluene in a Reach of East Drainage Ditch fate


total inputa volatilization converted to COZ biomass/insoluble intermediates soluble intermediates

97 13



8 17b 4


of yearly input 100 13 8 18b 4 57

a Based on yearly averages of flow rates and toluene concentration at the beginning (input) and the end (outflow) of the studied reach in field studies (2).*Most of this would be expected to be ultimately mineralized.

In assays with a marine organism at toluene concentrations up to 1 g/L, Robertson and Button (17)reported that 23%was converted to COZ,10%converted to biomass, and 67% remained as organic products. It appears that the East Drainage Ditch is more efficient in producing biomass and as efficient in converting toluene to COz. This may be explained by the fact that there are likely a number of different organisms present in this consortium, and they are each likely to be able to convert different intermediates to C 0 2 or biomass. Our assays were performed over an 8 h time period, and it is likely that more toluene would have been completely mineralized over a longer time period. Although mineralization efficiencywas high, 15%of the toluene which was initially present in solution was recovered as soluble intermediates in these experiments. These intermediates are expected to consist of nonvolatile oxidized aromatic and straight chain hydrocarbon species, including catechol, methylcatechol, benzoic acid, and cis-muconic acid (18). This general type of situation where biodegradation competes with volatilization as a sink for the compound may be a case where biodegradation is actually a drawback to removing hazardous contaminants from surface waters. If volatilization is the only sink, then no intermediates are left behind. However, if biodegradation occurs, approximately 15%of the toluene is left behind as nonvolatile and possibly harmful intermediates (although these may later be converted to COz). Thus, we hypothesize that in these types of situations biodegradation may actually increase the residence time of potentially harmful contaminants. The environmental fate of toluene in the studied section of the East Drainage Ditch has been determined (Table 3). The results show the impact of biodegradation as well as the influence of toluene on the microbial community of biomass. It also illustrates the cleansing effects of a short stream reach on the level of solvent in the stream. Seasonal Variations of Biodegradation Rates and Biomass. Cell counts in general show an agreement with measurements of in-situ biodegradation rates, which show a pattern of high rates in summer and low rates in winter. Cell counts show that highest cell numbers are observed in May and are maintained over the entire summer. Biodegradation rates determined in winter are 11% (rocks) to 14% (sediments) of biodegradation rates in the summer (Table l), consistent with field results (2). The decrease in biodegradation rates in the Mrinter is attributable to two effects: a decrease in cell population and a decrease in the reaction rate of the biodegradation reaction because of the decrease in temperature. Only a 2-fold decrease in 124


cell numbers is observed between counts in summer and winter. This indicates that a decrease in microbial activity is the main cause of the decrease in biodegradation rate. The observed enzyme effect is consistent with the work of Button and Robertson (19),who report that a 10"C decrease in temperature approximately halves biodegradation rates for marine organisms. Measurements of microbial FDA hydrolysis activity indicate that metabolic activity peaks in late autumn and then decreases 5-fold, reaching a minimum in late winter/ early spring. Interestingly, microbial FDA hydrolysis activity reaches its peak in October, after the warmest part of the season has passed. This behavior does not correspond to measured in-situ biodegradation rates (21, which tend to be lower in the fall than the summer. The observed discrepancy between microbial activity and in-situ biodegradation rate may be due to a large influx of dissolved organic matter into the stream in the form of falling leaves and senescent vegetation. Bott and Kaplan (20) and Lewis et al. (6)have both reported a positive correlation between organic matter and microbial activity. Thus, as autumn comes, we would expect to see an increase in DOC input to the stream and a large increase in microbial activity. However, if this organic matter were more easily degradable than toluene, it would be preferentially degraded, and one might expect to see a decrease in toluene biodegradation rate. Dibble and Bartha (21)have reported that the addition of a large amount of easily degradable organic matter decreased biodegradation rates for oil sludges by sediment microorganisms. It is possible that the decreased in-situ biodegradation rates that we observed during September were due to this same type of diauxic effect.

This work was supported by NIEHS Superfund Basic Research Program Grant 5P42ES04675-06. We thank John MacFarlane for his graphical services and technical assistance.

literature Cited (1) Madsen, E. L. Environ. Sci. Technol. 1991, 25, 1663-1673. (2) Kim, H.; Hemond, H. F.; Krumholz, L. R.; Cohen, B. A. Environ. Sci. Technol. 1995, 29, 108-116. (3) Wuhrmann, K. In Water Pollution Microbiology; Mitchell, R. M., Ed.; John Wiley and Sons: New York, 1972. (4) Gantzer, C. J. Ph.D. Dissertation, UniversityofIllinois at UrbanaChampaign, Urbana, 1986. (5) Gantzer, C. J.; Kollig, H. P.; Rittmann, B. E.; Lewis, D. L. Water Res. 1988, 22, 191-200. (6) Lewis, D. L.; Freeman, L. F., 111;Watwood, M. E. Environ. Toxicol. Chem. 1986, 5, 791-796. (7) Lewis, D. L.; Gattie, D. K. Appl. Environ. Microbiol. 1988, 54, 434-440. (8) Lewis, D. L.; Kellogg, R. B.; Holm, H. W. In Validation and

predictability of laboratory methods for assessing the fate and effects of contaminants in aquatic ecosystems; Boyle, T. P., Ed.; American Socity for Testing and Materials: Baltimore, MO, 1985; p 233. (9) Pignatello, J. J.; Johnson, L. K.; Martinson, M. M.; Carlson, R. E.; Crawford, R. L. Appl. Environ. Microbiol. 1985, 50, 127-132. (10) Knaebel, D. B.; Vestal, J. R. J. Microbiol. Methods 1988, 7, 309317. (11) Pignatello, J. J.; Johnson, L. K.; Martinson, M. M.; Carlson, R. E.; Crawford, R. L. Can.J. Microbiol 1986, 32, 38-46. (12) Yoon, W. B.; Rosson, R. A. Appl. Environ. Microbiol. 1990, 56, 595-600. (13) Schnurer, J.; Rosswall, T. Appl. Environ. Microbiol. 1982, 43, 1256-1261. (14) Rouse, H. Elementary Mechanics of Fluids; Dover Publications, Inc.: New York, 1978.

(151 Chow, V. T.;Maibent, D. R.; Mays, L. W.Applied Hydrology; McGraw-Hill,Inc.: New York, 1989. (161 Lewis, D . L.; Kollig, H. P.; Hall, T. L. Appl. Enuiron. Microbiol. 1983,46,146-151. (17)Robertson, B. R.; Button, D. K. Appl. Enuiron. Microbiol. 1987, 53,2193-2205. (18)Smith, M.R. Biodegrchtion 1990,1, 191-206. (19)Button, D.K.; Robertson,B. R. Limnol. Oceanogr. 1986,31,101-

(21)Dibble, J. T.;Bartha, R. Appl. Enuiron. Microbiol. 1979,37,729739.

Received for reviewApril 5, 1994. Revised manuscript received September 8, 1994. Accepted September 15, [email protected] ES940209A


(20)Bott,T.L.;Kaplan, L. A. Appl. Enuiron. Microbiol. 1985,50,508522.

@Abstractpublished in AduunceACSAbstructs, November 1,1994.