World Journal of Microbiology & Biotechnology 19: 571–581, 2003. 2003 Kluwer Academic Publishers. Printed in the Netherlands.
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Bioremediation of creosote-contaminated soil: a pilot-scale landfarming evaluation Harrison Ifeanyichukwu Atagana School of Earth Sciences, Mangosuthu Technikon, P.O. Box 12363, Jacobs, Durban 4026, South Africa Tel.: þ27-31-907-7477, Fax: þ27-319-072-892/31-907-2892, E-mail:
[email protected] Received 7 May 2002; accepted 13 March 2003
Keywords: Bioaugmentation, biodegradation, bioremediation, biostimulation, creosote-contaminated-soil, landfarming
Summary A pilot-scale landfarming investigation of the effects of biostimulation and bioaugmentation on a creosotecontaminated (258.3 g kg)1) mispah form (FAO: lithosol) soil, with a view to developing a cost-effective bioremediation methodology for creosote-contaminated soils was conducted in nine duplicate reactors, including two controls (Treatments 1 and 2). Treatments 3–9 were watered and aerated daily and Treatment 4–9 were monthly amended with mono-ammonium phosphate. Treatment 5–9 received further amendments as follows: Treatment 5, hydrogen peroxide; Treatment 6, indigenous microbial biosupplement; Treatment 7, sewage sludge; Treatment 8, cow manure; Treatment 9, poultry manure. Residual concentrations of creosote ranged between 29 and 215 g kg)1 after sixteen weeks. The phenolics and the 2- and 3-ringed polyaromatic hydrocarbons (PAHs) were removed below detectable levels or to very low levels. The 4- and 5-ringed PAHs were removed by between 68 and 83%. Indigenous microbial biosupplement and sewage sludge were the most effective in creosote removal. Hydrogen peroxide did not significantly enhance microbial population and creosote removal. There was no significant difference between the results obtained from the treatments amended with organic manures. However, there was a significant difference between the effects of the organic manures and the indigenous microbial biosupplement. Results from this study suggests that a combination of the two treatment techniques (biostimulation and bioaugmentation) would be a better approach to treating soil contaminated with very high concentrations of creosote.
Introduction Landfarming, an above-ground bioremediation technology for contaminated soils has been successfully used in treating petroleum-contaminated sites (USEPA 1995). The tilling process used in landfarming usually results in volatilization of lighter fractions of petroleum, while heavier fractions are degraded by biological processes (Sims et al. 1989; USEPA 1990; Wang et al. 1990; Testa et al. 1991; Baker & Herson 1994; USEPA 1995). Landfarming has been applied to soils contaminated with a wide range of other chemical pollutants (Stroo et al. 1989; Lajoie et al. 1994). However, the success of the technique may not necessarily be repeated with other pollutants. Mixed results have been obtained from landfarming of creosote-contaminated soils (Mueller et al. 1991) possibly due to the wide range of compounds present in creosote. The objective of the present study is to develop a costeffective remediation strategy for the treatment of creosote-contaminated sites in South Africa. Hence it is looking at employing landfarming, in conjunction
with biostimulation and bioaugmentation, to treat creosote-contaminated soils. It will also compare the efficacies of biostimulation and bioaugmentation in the landfarming process.
Materials and methods Soil samples About 500 kg of mispah form (FAO: Lithosol) soil heavily contaminated with creosote (258.3 g kg)1) was collected from a wood treatment site in KwaZulu–Natal midlands in South Africa. The samples were bulked and mixed in an electric concrete mixer before being stored at 4 C. A 1 kg sample of the soil was taken for creosote analysis. Experimental design Eighteen polyvinyl chloride (PVC) vessels of about 30 l capacity each, with a diameter of 56 cm and a depth of
572 22 cm, were filled with 25 kg of the creosote-contaminated soil described above. To each of the soil reactors was added 175 g of agricultural lime (CaCO3) to raise the soil pH to approximately 7. Table 2 shows the experimental design and the different treatments applied. All treatments were duplicated. To the soil in Treatment 1 was added 57 g of sodium azide to establish a sterile control. Sodium azide was applied twice, at the beginning of the experiment and in the seventh week of the experiment. The application was done by dissolving sodium azide in distilled water and diluting to 2 l before mixing thoroughly with the soil. The method was adapted from Snyman (1996). The pH of the soil in the reactors was measured weekly and when necessary, agricultural lime (7 g kg)1) was added to raise the pH to about 7. The pre-treatment level was about 5. Ambient air temperature and temperature of the soil in the reactors was measured daily at noon. Between 250 and 500 ml of water were added to the replicate reactors in Treatments 3–9 every 2 days to keep the soil moisture at about 70% field capacity, as determined by mass. Aeration of soil in Treatments 3–9 was effected by tilling the soil daily with a hand trowel. Mono-ammonium phosphate (MAP) fertilizer (1 g kg)1 soil) was added to Treatments 4–9 to raise the nitrogen level of the soil. The choice of MAP was made on the basis of previous results in crude oilcontaminated soil (Snyman 1996). The application of MAP, which changed the CNP ratio from about 250:1:4 to about 180:1:2 after the first application was repeated once every month for the duration of the experiment. As an alternative source of electron acceptor, hydrogen peroxide was introduced into the soil in Treatment 5, at a concentration of 500 mg l)1 (600 ll of a 39% solution) once a week (Thomas & Ward 1989). Treatment 6 received a weekly dose of 50 ml of an indigenous microorganism-biosupplement, which was prepared by adding 2 g of a creosote-contaminated soil sample to an enrichment broth containing (g l)1) NaNO3, 2; KCl, 0.5; MgSO4 Æ 7H2O, 0.5; Na2HPO4, 1; FeSO4 7H2O, 0.01; sucrose, 3; 30 ml creosote [adapted from Harrigan & McCance (1966)]. The mixture was homogenized in a blender for about 20 s. A second creosote-contaminated soil sample (2 g) was inoculated into 1 l of soil extract broth (Parkinson et al. 1971) which contained 30 ml creosote as additional carbon source. The broth was sonicated to release soil-adsorbed creosote and disperse the creosote properly before inoculation. The microbial suspension was incubated and aerated by means of a sintered glass diffuser at 25 C for 4 days in two 50 ml glass filter funnels with taps. Microbial populations were estimated by plate counts on nutrient agar. Treatment 7 received a 10% (w/w) addition of sewage sludge from a combined domestic and industrial waste water treatment plant which was dosed twice, the first at the beginning of the experiment and the second in the
H.I. Atagana seventh week, during the treatment period. Treatments 8 and 9 received 10% (w/w) cow manure and poultry manure respectively. Both treatments were applied at the beginning and in the seventh week during the duration of the experiment. Samples were taken at weekly intervals from all the treatments for analysis. The soil samples were analysed for changes in concentration of creosote and for changes in concentration of selected creosote components. Counts of microorganisms were made on nutrient agar and mineral salts agar supplemented with 30 ml l)1 creosote. Antibiotics and fungicides were incorporated into the media for isolating fungi and bacteria, as described by Atlas & Bartha (1972). Soil nutrient analyses Determination of total nitrogen was done by a soil digestion method as described by Forster (1995). Ammonium in the extract was determined by distillation and titration. Isolation and characterization of indigenous soil microorganisms Soil samples of 1 g each were diluted serially (10)1– 10)9), using aqueous physiological saline solution (0.85% m/v) and plated out on soil extract agar and nutrient agar to estimate total colony forming units (c.f.u.). Soil extract agar was prepared as described by Parkinson et al. (1971). The dilutions were also plated on potato dextrose agar (PDA), which was formulated as described by Davis & Westlake (1978). Plates were incubated at 25 C for 24 h for bacteria and at 25 C for 4 days for fungi. Pure cultures were obtained from colonies on the plates by means of dilution streaking. Gram stains and light microscope examinations were carried out on all bacterial cultures. Cultural and morphological characteristics of all fungal isolates were used to identify the fungi to genus level (Raper & Thom 1968). Plate counts of total heterotrophs and creosote-degrading species A 1 g sample of soil was taken from each reactor on a weekly basis and analysed for the total number of microorganisms present, using standard dilution plating techniques on nutrient agar. The soil was also analysed for total number of creosote-degrading microorganisms by inoculating a modified version of the medium, described by the Organisation for Economic Co-operation and Development (OECD), containing 30 ml creosote oil as the sole carbon source. The OECD medium was prepared by adding 17 g agar, 4 ml FeCl3 (0.25 g l)1), 1 ml each of MgSO4 Æ 7H2O (22.5 g l)1), CaCl2 (27.5 g l)1) and (NH4)2SO4 40 g l)1 to 2 ml of the following mixture: KH2PO4 (8.5 g l)1), K2HPO4 (21.75 g l)1), NaH2PO4 Æ 7H2O (33.4 g l)1) and NH4Cl
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Bioremediation of creosote-contaminated soil (1.7 g l)1) and diluting to one litre with distilled water. The mixture was autoclaved at 121 C for 20 min and cooled. Creosote oil was filtered through a hydrophilic membrane (0.4 lm) pore filter and agar was added to it before the plates were poured. Determination of concentrations of creosote and selected creosote components in the soil Total creosote concentration in the soil was analysed using the USEPA 4181.1 (1983) method as follows: creosote-contaminated soil (2 g) and 2 g anhydrous Na2SO4 were placed in a 30 ml amber glass vial. Carbon tetrachloride (10 ml) was added and the vial was sealed with a teflon-lined screw cap. The sealed vial was vortexed for 15 s and then placed in a sonicating bath (Whaledent Biosonic) for 15 min before remixing on the vortex mixer for about 15 s. It was then placed in the sonicating bath for another 15 min. The solvent was transferred to a clean, dry vial containing 1 g activated Florosil (Sigma) and 0.6 ml water [i.e. 6% water (w/ w)]. The sealed vial was shaken for 1 min and allowed to stand overnight at ambient temperature. This silica ‘clean-up’ procedure was used to remove interfering humic materials (USEPA 1986). The extract was finally filtered through a Whatman GF/C glass fibre filter. The filtrate was made up to 10 ml in a volumetric flask and the absorbance determined with a Nicolet Avater 320 Infra-red spectrophotometer at wave numbers between 2760 and 3070 cm)1 and an integration value for the absorbance peak area was automatically generated. Results from the samples that were amended with 10% (w/w) organic manure were corrected with a correction factor of 0.043, which compensated for the change in mass due to the addition of 5 kg of manure used as amendment. Determination of changes in the concentration of selected creosote hydrocarbons was done by Soxhlet extraction and GC/FID. The gas chromatograph was a Varian-3800 with argon as the carrier gas and fitted with a 30 m capillary column with 0.25 mm internal diameter and 0.25 lm film thickness, and a flame ionization detector (FID). Two temperature programmes were run in order to obtain good separation and quantification of the more volatile compounds. The first temperature
programme was: 60 C, 4 min, followed by ramping at 10 C/min up to 235 C, maintained for 40 min; injector temperature 220 C. The second temperature programme was used for the analysis of the more volatile compounds, viz. 20 C; 1 min 40 C, 1 min, 10 C/min, ramping up to 200 C, maintained for 20 min; injector temperature 220 C (Eriksson et al. 2000).
Results and discussion Characteristics of the contaminated soil The concentration of creosote in the composite soil sample was 258.3 g kg)1 [25.83% (w/w)]. Individual samples from the study area had creosote concentrations varying from 180 (18%) to 380 g kg)1 (38%). The extractable phosphorus content of the soil was 4.75 mg kg)1 soil. This meant that the phosphorus content of the soil had to be increased, because the typical phosphorus concentration of a soil ranges from 12 to 18 mg kg)1 soil (Snyman 1996). Total nitrogen was found to be 0.081%. Inorganic nitrogen normally represents about 2% of the total nitrogen in soils. This meant that additional nitrogen was necessary in the soil. On drying at 105 C, 37.3% of the total mass of the sample was lost (Table 1). This represents mainly water
Table 1. Characteristics of the creosote-contaminated soil studied. Parameters
Means
Total creosote pH Total nitrogen Extractable phosphorus Clay Fine silt Sand Total exchangeable cations Exchange acidity Acid saturation Los on drying at 105 C (H2O and Volatile organics) Non-volatile fractions and water of crystallization at 800 C Total heterotrophic microorganisms
258.3 g kg)1 5.45 0.08% 4.75 mg kg)1 18.8% 18.8% 62.5% 4.61 cmol/l 0.19 cmol/l 12.5% 37.3% 17.5% 2.18 · 106 c.f.u. g)1
Table 2. Experimental design and treatments for each soil reactor. Duplicate Treatments (A and B)
Water
Sodium azide
MAP
Tilling (daily)
H2O2
Indigenous microbial culture
Sewage sludge
Cow manure
Poultry manure
1 2 3 4 5 6 7 8 9
No No Yes Yes Yes Yes Yes Yes Yes
Yes No No No No No No No No
No No No Yes Yes Yes Yes Yes Yes
No No Yes Yes Yes Yes Yes Yes Yes
No No No No Yes No No No No
No No No No No Yes No No No
No No No No No No Yes No No
No No No No No No No Yes No
No No No No No No No No Yes
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and volatile organic matter. A further 17.5% was lost on ignition at 800 C representing the non-volatile organic matter. The soil pH in most of the treatments fluctuated between 7 and 5.5. However, higher pH values occurred in Treatments 6, 7 and 9, where the highest recorded values were 7.5 in Treatments 6 and 7, and 7.8 in Treatment 9. The lowest pH encountered outside this range was 4.7 in Treatment 5, 5.0 in Treatment 3 and 5.1 in Treatment 4. These pH values are, however, not outside the conventional values reported to sustain microbial activity in the soil (Alexander 1999). Hence microbial activity in the reactors continued to flourish during the period of the experiment. Liming, however, kept the pH at about 7 during the experimental period. There was a general decrease in pH in this experiment. The decrease in pH in Treatment 1 (sterile control) can be associated with very slow degradation that occurred naturally through some other action such as photoxidation. Soil temperatures were observed to be slightly below the mean day air temperature, in some cases. However, in a few instances where biosupplements were used, soil temperature was slightly above the air temperature but usually by not more than 1 or 2 C. The overall temperature pattern in the soils followed the mean day temperature of the air, with the lowest temperature (21 C) recorded in the late winter months and the highest (31 C) in the summer months. Degradation of creosote The decrease in creosote concentration in Treatment 1 (sterile control) from 258.3 to 215.2 g kg)1, representing a 16.7% loss in creosote content of the soil (Figure 1) could not be attributed to microbial action considering
that microbial counts continued to be about zero throughout the experimental period because of the addition of sodium azide. However, it indicates that a large amount of volatile components of creosote remained in the soil despite its age, as this reduction can only be attributed to volatilization. Another possible explanation is that seepages from the discharge unit, which still held a large quantity of creosote, may have continued to contaminate the soil further with the lower molecular mass, more volatile fractions of creosote. It is also possible that the volatile fractions may have been ‘locked up’ in interstitial spaces and closed pore spaces in the soil matrix and only became released when the soil was turned over and homogenized. The process of aeration and homogenization may have resulted in the oxidation and/or breakdown of higher molecular mass components into volatile fractions or undetected intermediates or end-products. Volatilization was not expected to be high in the controls as it was assumed that the bulk of creosote compounds left in the soil would be the higher molecular mass fractions that are not volatile and known to degrade slowly. In Treatment 2, (natural control), the decrease in creosote concentration from 258.3 to about 156 g kg)1 represents a 39.5% loss in concentration (Figure 1). This increased decline in creosote concentration, which is 23% larger than observed in Treatment 1 (sterile control) can be attributed to biodegradation, as can be seen from the results of the plate counts of the microorganisms in this treatment where the numbers of total heterotrophic microorganisms and hydrocarbon degraders increased substantially (Figures 2 and 3). These increases soon levelled off, maintaining a relatively unchanging population size, probably as a result of stabilization of conditions such as aeration and moisture, since this treatment was neither aerated nor
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Figure 1. Changes in creosote concentration (g kg)1) in creosote-contaminated soil during pilot-scale landfarming. Values are means of two 1 standard error.
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Bioremediation of creosote-contaminated soil 4.50E+07 4.00E+07 3.50E+07 3.00E+07 2.50E+07 2.00E+07 1.50E+07 1.00E+07 5.00E+06 0.00E+00 0
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Figure 2. Counts of heterotrophic microorganisms (c.f.u. g)1) in creosote-contaminated soil during pilot-scale landfarming. Values are means of two 1 standard error.
watered. The natural soil conditions prevailing in Treatment 2 allowed an increase in the microbial population at the expense of the creosote hydrocarbons. Pre-treatment such as homogenization must have enhanced aeration in the soil and/or the accessibility of hydrocarbons to the microbial population and thus promoted creosote degradation (Figure 1). Treatment 3, which was regularly watered and aerated throughout the treatment period, but did not receive any nutrient supplement, showed improved microbial activity (Figures 2 and 3), as well as improved creosote
reduction (from 258.3 to 107.5 g kg)1) representing 58.4% loss in concentration (Figure 1), compared to Treatments 1 and 2 (p 0.05) (Figure 1). These results establish that watering and mechanical tilling are effective means of increasing soil aeration and moisture, which in turn increases creosote degradation. Poor natural degradation of creosote compounds in the soil may have been caused by their inaccessibility to the microbial biomass by being held within the structure of soil aggregates. This is common when relatively immiscible liquid phases such as dense non-aqueous phase
2.60E+06 2.40E+06 2.20E+06 2.00E+06 1.80E+06 1.60E+06 1.40E+06 1.20E+06 1.00E+06 8.00E+05 6.00E+05 4.00E+05 2.00E+05 0.00E+00 0
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Figure 3. Counts of creosote degraders (c.f.u. g)1) in creosote-contaminated soil during pilot-scale landfarming. Values are means of two 1 standard error.
576 liquids (DNAPL) become trapped in interstitial pores (Kimball 1994). Hydrocarbons adsorbed to soil particles that later become very tightly compacted become less available to microbial biomass on the inaccessible surfaces. Aeration and moisture availability in such a soil matrix become limited and thus inhibit further microbial activity (Alexander 1999). Tillage would have broken up aggregates, thus exposing the hydrocarbons to microbial attack. The results from these treatments also establish that the high concentration of creosote found in the soil after a long period of disuse of the facility was due to poor aeration and/or physical inaccessibility, which has hindered the biological breakdown of the creosote. Treatments 4–9 received fertilizer treatment in the form of MAP to supplement the nitrogen and phosphorus contents of the soil. They were all aerated daily and supplied with between 250 and 500 ml of water every 2 days, as determined by the water content of the soil throughout the experimental period. Treatments 5–9 received additional nutrient supplementation. In Treatment 4, creosote concentration decreased from 258.3 to 95.7 g kg)1 representing a 62.9% loss in concentration. This increase above treatment 3 is attributed to the nutrient supplementation. Microbial populations were also considerably higher than in Treatment 3 (Figures 2 and 3). This initial deficiency in nitrogen and phosphorus in the soil was thought to be one of the factors responsible for the poor natural degradation that took place in the soil over the long period of contamination. Thus the enhancement of microbial activity and the consequent improved creosote degradation in Treatment 4 (Figure 1) was attributed to the supplementation of the soil in the reactors with MAP. The weekly dose of hydrogen peroxide (H2O2) added to Treatment 5 provided an alternative source of electron acceptor. However, only a concentration reduction of 61.6%, representing a comparatively small improvement (3.2%) compared to Treatment 3, which showed a reduction of 58.4% and a lower reduction compared to Treatment 4, which showed a reduction of 62.9% (p 0.05) was observed. Hydrogen peroxide was meant to supply large amounts of oxygen for the biodegradation process. However, H2O2 is known to be more reactive than oxygen (Fiorenza & Ward 1997) and as a result it subject to decomposition by the action of light, metal ions and enzymes. The soil in the reactor was supplied with about 500 ml of water and aerated daily throughout the experimental period. Results from this treatment showed that hydrogen peroxide as an alternative electron acceptor did not appreciably enhance biodegradation of the creosote. The result in this treatment suggests that daily tillage was sufficient to provide an adequate supply of oxygen for creosote degradation. In addition, some of the lower molecular mass components may have been lost due to continued tilling of the soil. Toxicity and consequent poor adaptation of the resident microorganisms to the hydrogen peroxide could also have occurred, since it is known to
H.I. Atagana cause toxicity to microorganisms during decomposition as a result of the free hydroxyl (OH•) radical being formed (Halliwell & Gutteridge 1985). This action has been reported to damage DNA (Ananthaswamy & Eisenstark 1977; Demple & Linn 1982; Demple et al. 1986). However, the slight improvement over Treatment 3 is possibly due to the additional oxygen from the hydrogen peroxide. Treatments 6 and 7 were supplemented with additional nutrients in the form of indigenous microbial biosupplement and sewage sludge, respectively. Both treatments contained the highest counts for total heterotrophic and creosote degrading microorganisms. (Figures 2 and 3). Microbial counts increased rapidly between the second and third week of experimentation and continued to increase more slowly until the 16th week. A rapid increase in counts of creosote-degrading microorganisms was observed to continue in Treatment 7 until the fourth week, reaching a peak of 2.1 · 106, before declining slightly over the next 3-week period and then becoming relatively stable for the remaining period of the experiment (Figure 3). This rapid increase indicates a fairly rapid adaptation of microorganisms from the sewage sludge to the contaminating creosote. Although there was a slight fall in microbial population, which coincided with a period during which most of the cocci became scarce, the counts continued to increase gradually afterwards. The increase in numbers of creosote-degrading microorganisms in Treatment 6 continued slowly after the third week. Results in Figures 2 and 3 show that the organisms from the sewage sludge inoculum proliferated more rapidly than those from the enriched indigenous soil biosupplement. This may be due to the conditions established in the enrichment reactor to improve the growth of the organisms, which were not present in the pilot-scale soil reactor system. These two treatments (6 and 7) showed substantial degradation of creosote and concentrations decreased from 258.3 to 29.1 and 36 g kg)1 respectively. This represents a total reduction of 88.7 and 86.1% respectively (Figure 1). Enrichment cultures have been used in different soil reactors to accelerate degradation of hydrocarbons (Snyman 1996), in many cases acting purely as microbial inocula for the biodegradation process. Even though microbial activity in the reactor supplemented with sewage sludge was slightly higher than that in the treatment with indigenous biosupplement, the latter showed a higher percentage reduction (p 0.05) in creosote content of the soil. This can be attributed to the specificity of the indigenous soil organisms to creosote hydrocarbons compared to those of sewage origin, which must still adapt to the new hydrocarbon source. Treatments 8 and 9, which received cow and poultry manure, respectively, showed substantial microbial activity and reduced creosote concentration from 258.3 to 38.2 and 40.1 g kg)1, respectively, representing a total reduction of 85.2 and 84.5%, respectively (Figure 1). These decreases are comparable (p 0.05) with those
Bioremediation of creosote-contaminated soil observed in Treatments 6 and 7, which showed reductions of 88.7 and 86.1%, respectively. The decrease in creosote concentrations in Treatments 8 and 9 was rapid up to the ninth week, reaching 60.4 and 69 g kg)1, respectively. The degradation rate then slowed until the 16th week, with only very slight decreases being recorded over the remaining 7 weeks of the experiment. This slow degradation rate over the second half of the experiment can be attributed to the very low concentration of hydrocarbon remaining in the soil, which led to carbon limitation for the degrading microorganisms. Microbial utilization of carbon as a growth substrate in the soil was not expected to deplete only the carbon supply of the soil but also other utilizable soil nutrients such as nitrogen. These conditions could lower the microbial population of the soil, leading to reduced degradation of the creosote. Low concentrations have been reported to hinder microbial utilization of hydrocarbons in the environment (Alexander 1999). Another possible explanation is that the metabolites of the degrading microorganisms may have accumulated in the soil and become inhibitory to the organisms (Veldkamp & Jannasch 1972). This is, however, less likely because the microbial population continued to increase, although very slowly until the end of the experiment. The results from the treatments supplemented with organic fertilizers (sewage sludge, cow manure and poultry manure) demonstrate creosote degradation where the active microorganisms already have a substantial amount of utilization carbon supplied in the manure. These substrates also have high amounts of nitrogen and phosphorus and other required mineral nutrients that will tend to stimulate microbial activity. This nutritionally rich environment will naturally cushion the organisms from the initial shock experienced on transfer to a medium containing a new recalcitrant carbon source. Adaptation to the new substrate would, therefore, be achieved in a reasonably short time. The variety of organisms isolated from these reactors in the first week of operation and the high level of creosote degradation observed in these reactors suggest that the organisms only require a short period of adaptation to the new carbon substrate. This may have been due to cometabolism of the new carbon substrate along with the carbon substrates available in the manure. This could have been achieved by the production of enzymes necessary for the breakdown of the creosote substrate while the microorganisms were still metabolising the original substrate. It is possible that such enzymes required for the breakdown of the polluting creosote hydrocarbons have been produced as constitutive enzymes (Alexander 1999) while metabolizing the manure substrate. This situation may have reduced the acclimation period significantly. Another possibility is the production of inducible enzymes as a result of the introduction of the creosote hydrocarbons. These enzymes may have been present in very limited amounts, typically because their production has been suppressed by the products of enzymes for the catabolism of
577 another substrate (Alexander 1999). Such enzymes can be readily induced by the abundant presence of their specific substrate since enzyme induction is usually carried out in very short periods of time; it is commonly completed in minutes (Richmond 1968). The other possibility is the creation of new environmental conditions brought about by the mixing of the creosotecontaminated soil and the manure. This may have affected the nutrient and substrate composition available to the microbial community and consequently created an entirely new environment that may have encouraged new species to invade and colonize the substrate. Since there was no taxonomic characterization of the isolates, it was not possible to confirm whether new species colonized the substrate or not. From a comparison of Treatment 1 with the others (p 0.05), it can be seen that the volatile fractions of the pollutant, which were removed by volatilization, constituted a reasonable proportion of the total contaminating hydrocarbons. However, the non-volatile fractions had to be degraded by soil microorganisms, as is evident from the results from Treatments 2–9, which showed continued microbial activity and further creosote degradation. This latter effect was, however, not observed in Treatment 1 (sterile control), in which only 43.1 g kg)1 of the original creosote present was removed. There was no significant difference at p 0.05 in reduction in creosote concentration in Treatments 7–9. There was, however, a significant difference in concentration reduction between the indigenous biosupplement and the organic manures. Analyses of selected creosote components monitored in the degradation studies, showed that phenol, o-cresol, p-cresol and m-cresol, were reduced to undetectable levels (
