Biodegradation of heavy oil from the Nakhodka oil spill by indigenous microbial consortia

Biodegradation of heavy oil from the Nakhodka oil spill by indigenous microbial consortia

Siti Khodijah Chaerun


A biodegradation process of heavy oil from the Nakhodka oil spill by indigenous microbial consortia was monitored over the 429-day at laboratory scale. The indigenous microbial consortia consisted of bacteria and fungi from the Nakhodka oil spill in the Sea of Japan as well as the bacterium Pseudomonas aeruginosa isolated from Atake seashore, Ishikawa Prefecture, Japan. Both bacteria and fungi had a significant role in the observed biodegradation of heavy oil during the 429-day bioremediation of the Nakhodka oil spill with respect to the pH of the solution. Hydrocarbon-degrading bacteria had tendency to play greatest role at the neutral-alkaline condition (pH: 7~7.8). On the contrary, when pH shifted to acidic (pH: 2~4), the fungi took over to degrade heavy oil. Fungal succession during the 429-day bioremediation was predominated by Rhinocladiella sp., while all fungi which survived in degrading heavy oil were identified as Rhinocladiella sp., Aspergillus sp., Acremonium sp., and Penicillium sp. During that period, the aliphatic hydrocarbons were reduced significantly, whereas the aromatic hydrocarbons remained relatively constant, and there was the production of metabolites after 429 days of bioremediation. These results suggest that both bacteria and fungi as indigenous mixed microbial consortia may contribute to the biodegradation of heavy oil from the Nakhodka oil spill. In addition, the benevolent and favorable microbial interactions in bioremediation are essential and requisite to completely biodegrade oil spills for bioremediation success.

Keywords: Bioremediation; Hydrocarbon-degrading bacteria; Rhinocladiella sp.; Aspergillus sp.; Acremonium sp.; aliphatic hydrocarbons; aromatic hydrocarbons.


In situ bioremediation processes represent a complex ecosystem within which many microbial interactions are occurring. Since microorganisms compete with each other in the uptake of substrate for their growth, the favorable microbial interactions in bioremediation processes to remove environmental pollutants are important for indigenous microorganisms in contaminated sites. The microbial interactions are categorized with regard to the effects that the organisms have on one another: neutral, benevolent, or antagonistic [1]. Neutral interactions are those which have no observable effect, i.e., no real interaction. Benevolent interactions (i.e., commensalism and mutualism) are those in which one organism is aided without any harm to the other. Antagonistic interactions involve a detrimental effect upon one organism [1]. There is no one species that is responsible for bioremediation of a contaminated site. The synergism between indigenous microorganisms is very significant to in situ bioremediation because of their ecological relationship within bioremediation consortiums in enhancing the overall rate of degradation of contaminants.

Pollution at most sites has existed for long periods. Oil spills (e.g., the Nakhodka oil spill) provide adequate time for the natural development of seed organisms that respond to the available energy of the spill [2]. For most bioremediation, proper environmental conditions must be maintained for microbial growth. Microorganisms are principally sensitive to temperatures, pH, contaminant toxicity, contaminant concentration, moisture content, nutrient concentrations, and oxygen concentration [3].

The aim of this study is to investigate the interactions of hydrocarbon degrading bacteria with fungi, as natural mixed cultures of microorganisms, in biodegrading heavy oil from the Nakhodka oil spill at laboratory scale (ex situ bioremediation). The experimental conditions were designed as natural as possible without any supplement to reflect in situ bioremediation processes, as well as the true complexity of petroleum hydrocarbon degradation in natural environments where the compounds are present in multicomponent mixtures. Changes in microbial community during a bioremediation processes were monitored, in particular fungal community. Identification of fungi was also conducted, as well as the dominating fungus for fungal succession. In addition, the metabolic products as a result of heavy oil biodegradation were investigated. In this study, the different treatments were also carried out for the mixing conditions to reflect marine and coastal environments that are significantly influenced by wind and wave actions.

Materials and Methods

Bacterial Strain

Apart from microorganisms inhabiting natural seawater (NSW) and heavy oil, a bacterium capable of degrading heavy oil was also added as an inoculum in order to promote the degradation of the heavy oil. The bacterium was isolated from Atake seashore, Ishikawa Prefecture, Japan. Using 16S rDNA sequencing, this bacterium was affiliated to Pseudomonas aeruginosa (98% similarity). This bacterium was selected for study because it produced high extracellular emulsifying activity after 2 days of incubation in the presence of heavy oil (2% v/v) and 1 g/l of yeast extract [4, 5, 6].

Microbial Culture and Batch Experimental Conditions

Batch bacterial experiment was conducted with two successive different treatments: with shaking for 105 days (0-105 days) and without shaking for 324 days (105-429 days). The growth medium contained the necessary components needed for bacterial growth. The following was included for each litre of medium: 25 g of N[H.sub.4]N[O.sub.3], 0.5 g of Fe[C.sub.6][H.sub.5][O.sub.7] x n[H.sub.2]O (ferric citrate), and 0.5 g of [K.sub.2]HP[O.sub.4] per litre of distilled, deionized water (NDW). Natural seawater (NSW) without filtration (collected from the Sea of Japan) was also added to each flask as additional medium [6]. Briefly, the batch reaction vessel consisted of acid-washed 1000 ml Erlenmeyer flask containing 600 ml of growth medium: 480 ml of NSW was mixed with 120 ml of an autoclaved NDW. The pH of the medium was adjusted to pH 7.8 using 1N NaOH solution. Before introducing the inocula, which originated from the heavy oil, and the NSW medium into the flask, the flask, together with its contents, was autoclaved, allowed to cool to room temperature (24[degrees]C), and then inoculated with NSW medium, the stock liquid culture of bacterial strain Pseudomonas aeruginosa (4.5% v/v), and heavy oil (collected from the Nakhodka oil spill) to a final concentration of ~150 g/l, serving as the sole carbon and energy source [2, 7, 8]. Culture was incubated for 429 days (~1.5 years) at room temperature (24[degrees]C) with shaking at 125 rpm for 105 days, allowing microorganisms to grow, then without shaking (a static condition) for another 324 days. Samples were removed periodically at sampling periods of 0, 36, 64, 105, 239 and 429 days. Separate sets of samples were made up and prepared for analysis by gas chromatography (GC), scanning electron microscopy (SEM), and transmission electron microscopy (TEM). In addition, the liquid medium of experimental system was supplemented with 1g/l of yeast extract after 36 days of the course of the experiment [6].

Electron Microscopy

For SEM observation, freeze drying was used for sample preparation [9]. Briefly, the suspension samples were fixed with 2.5% (vol/vol) glutaraldehyde, pipette-drawn, mounted on 0.22 [micro]m membrane filter, washed and fixed with t-butyl alcohol, subsequently frozen in liquid nitrogen, and dried up with low-vacuum SEM. After freeze-drying completely, samples were transferred to the brass-stub with double-sided adhesive carbon tape, coated with carbon, and then observed by using a scanning electron microscope (JEOL JSM-5200 LV). For TEM observation, samples were fixed with 2.5% v/v glutaraldehyde for 2 h at 4[degrees]C, mounted on copper specimen grids, allowed to dehydrate at room temperature, and then viewed using a JEOL JEM-2000EX transmission electron microscope [6].

Isolation and Identification of Fungi

At the end of this experiment, fungi were isolated from the solution and the plate floating on water surface of the solution by plating on Malt Extract agar plus 0.85% NaCl, and 1 g/l yeast extract. For fungal identification, microcultures were performed by using malt extract agar media. Microcultures were examined macroscopically and microscopically (40x and 100x with cotton blue) for their identification.

Analytical Techniques

The Eh and pH in solution were monitored using pH meter and Eh meter (Horiba) at set time intervals. To determine the total cells, 1 ml of the homogenized solution containing cells was placed in vials, serially diluted with fresh minimal salts media, and plated in nutrient agar (NA) plates supplemented with 1 g/l yeast extract and 0.85% (w/v) NaCl, then incubated at 25[degrees]C for 3 to 5 days. For statistics, sample analyses were performed in triplicate. The data are presented as the arithmetic mean [+ or -] standard deviation of the mean [6].

The hydrocarbon compounds of heavy oil at the onset (0 day) and the end of experiment (429 days) were determined by gas chromatography (GC) analysis with a flame ionization detector (FID) using a HP-1 capillary column. Approximately 1.0 g of heavy oil-cell complexes were extracted with 5.0 ml of hexane in 10-ml vials with Teflon-lined screw caps by shaking at 150 rpm in darkness for 24 hours. Extracts were filtered through glass wool to remove particulates, and mixed with a small amount of copper to remove sulphur. The purified extract was injected to and percolated over a silica gel column (70/230 mesh, 0.5 by 4 cm) until reaching the height of 5 mm above silica gel column, then eluted with: 2 ml of hexane to obtain the n-alkane fraction, 3 ml of hexane-benzene (3:1) to obtain the aromatic fraction, and 3 ml of benzene-methanol (4:1) to obtain the ketone fraction of the heavy oil. For the aromatic and ketone fractions, the residual solvents (i.e., benzene and methanol) were removed by a gentle stream of nitrogen, and 3 ml of hexane were added immediately. 1[micro]l of each fraction was injected by a cool on-column inlet with helium as carrier gas, and the oven was temperature programmed as follows: at initial temperature of 50[degrees]C, followed by 30[degrees]C/min to 120[degrees]C, then 5[degrees]C/min to 310[degrees]C. The major hydrocarbon compounds of heavy oil were identified on the basis of their retention time.


Microbial Growth

Plate counting showed that the numbers of cells (CFU) in the solution increased slightly after 36 days of incubation from an initial value (at the onset of the experiment) of 53 x [10.sup.5] [+ or -] 10 x [10.sup.5] CFU/ml to 65 x [10.sup.5] [+ or -] 5.7 x [10.sup.5] CFU/ml (Table 1). After 36 days of incubation, the liquid medium was supplemented with 1 g/l yeast extract. The supplement of yeast extract did not result in an increase in microbial cell number. Subsequently, after 64 days of incubation up to the end of experiment (at 429 days), cell numbers appeared to decline markedly: 25 x [10.sup.5] [+ or -] 7.3 x [10.sup.5] CFU/ml at 64 days, 9 x [10.sup.5] [+ or -] 2.2 x [10.sup.5] CFU/ml at 105 days, 8 x [10.sup.5] [+ or -] 3.1 x [10.sup.5] CFU/ml at 239 days, 4 x [10.sup.5] [+ or -] 1.0 x [10.sup.5] CFU/ml at 429 days. These low cell number densities indicated that the cells utilized the heavy oil as growth substrate very slowly due to a high concentration of heavy oil (~ 150 g/l). In addition, TEM images showed that the spherical and rod-shaped bacterial cells (some of which were dividing) appeared to be covered with and surrounded by heavy oil and other amorphous material, assumed to be extracellular polymeric substances (EPS) (Fig. 1), while SEM images showed the bacterial and fungal cells growing well and multiplying in colonies (Fig. 2).

pH and Eh

The pH of the solution remained relatively constant (maintaining the initial pH = ~8) during the first 36 days of incubation after which time, microbial growth significantly reduced the pH of the aqueous solution to: 3.7 [+ or -] 0.2 at 64 days, 3.4 [+ or -] 0.1 at 105 days, 2.0 [+ or -] 0.3 at 239 days, 1.9 [+ or -] 0.1 at 429 days (Table 1). Over the course of the experiment, bulk Eh values ranged from 179 [+ or -] 1.5 to 362 [+ or -] 2.0, indicating that the biodegradation of heavy oil took place under aerobic condition (Table 1).



Microbial Community and Succession During Biodegradation of Heavy Oil

Over a period of 429 days, both bacteria and fungi played a significant role in degrading heavy oil (Table 1). When the solution pH was neutral-alkaline condition (pH 7~8), the bacterial growth predominated: the bacterial growth percentage was in the range of 98.5-100%, while the fungal growth percentage was in the range of 0-1.5%. Conversely, when the pH of solution decreased to be less than 5, the fungal growth was dominant: the fungal growth percentage was in the range of 99.5-100%, while the bacterial growth percentage was in the range of 0-0.5%. Two fungal families were found; Dematiaceae (consisting of 1 genus: Rhinocladiella sp.) and Moniliaceae (consisting of 3 genera: Acremonium sp., Aspergillus sp., and Penicillium sp.). Rhinocladiella sp., Acremonium sp., Aspergillus sp., and Penicillium sp. were characterized by black, white, yellow and pink colored colonies on agar plate, respectively. However, Rhinocladiella sp. appeared to be predominant over the 429-day biodegradation as the pH of solution was in the range of 2 to 4.

Biodegradation of Heavy Oil Over A 429-Day Bioremediation

The gas chromatographic profiles of heavy oil as n-alkane fraction, aromatic fraction, and ketone fraction are given in Figs. 3-5. The aliphatic hydrocarbons were reduced significantly after 429 days (Fig. 3), whereas aromatic hydrocarbons remained relatively constant and new aromatic hydrocarbons were formed (Fig. 4). In addition, the formation of ketone was also observed, indicating the metabolites of heavy oil biodegradation (Fig. 5). Analysis of the heavy oil as n-alkane fraction at the onset of the experiment (0 day) showed that the Nakhodka oil spill mostly contained n-alkane of [C.sub.16]-[C.sub.32] among which the hydrocarbon components ([C.sub.20]-[C.sub.25]) were abundant. While little or no biodegradation of aromatic hydrocarbons was observed, the extensive biodegradation of aliphatic hydrocarbons occurred after 429 days (at the end of the experiment): shorter chain hydrocarbons [C.sub.16] to [C.sub.21] were highly degraded, whereas longer chain hydrocarbons [C.sub.22] to [C.sub.32] were moderately degraded.





While little increase in cell number density was observed over a 429-day bioremediation in the Eh range of 179 ~ 362, the decrease of the solution pH was as well observed (Table 1). This study indicates that the biodegradation of heavy oil has occurred under aerobic condition. Such a condition is considerably more amenable to bacterial and fungal growth (Table 1). However, Eweis et al. [3] reported that in well-aerated soils both bacteria and fungi will be important, but in poorly aerated soils the bacteria alone are responsible for the biological and chemical changes taking place. Thus, treatment without shaking has a significant effect on the biodegradation rate of heavy oil in this study. A low biodegradation of hydrocarbons occurred particularly as shaking stopped at a period of days of 105 to 429 due to poorly aerated aqueous phase, thus resulting in little fungal role under acidic condition, as confirmed by low fungal cell number densities. Therefore, the cell number of fungi appeared to be lower than that of bacteria throughout the experiment.

Moreover, the present study also confirms the significant role of both bacteria and fungi, as natural inocula, in remediating the Nakhodka oil spill under uncontrolled parameters, in particular pH. Over the 429-day bioremediation, there was a benevolent microbial competition of bacteria and fungi in degrading heavy oil which was dependent upon the pH of the solution. The number of bacteria or fungi and the predominant species present are a function of the specific environment (e.g., pH) (Table 1). The bacteria play the largest role in the bioremediation processes under neutral-alkaline condition. In contrast, when the solution pH decreased to be acidic, the fungi play a role in the degradation of heavy oil. The high decrease of the pH in the solution appears clearly as a result of aerobic biodegradation of hydrocarbons that released organic acids, resulting in the formation of hydrogen ions as a final product [3]. The addition of protons to the solution yielded the pH decrease of the ambient solution (Table 1). The decrease in pH might have encouraged the predominance of a competitor (i.e., more acid-tolerant microorganisms), therefore causing drastic shifts in the relative numbers of different species in the population of the solution. These, in turn, might allow fungi to thrive and become predominant (Table 1), since most fungi prefer an acid environment and have minimum pH values between 1 and 3 with an optimum pH near 5 [10]. It is also possible that competition among species for nutritional requirements (related to the variety of sources of carbon and nitrogen available and to the organic growth factors present in the aqueous solution) might have influenced the predominance of microorganisms as well (Table 1), since the biodegradation of hydrocarbons produces some metabolites [11], which are in good agreement with our present result (Figs. 4-5). However, both bacteria and fungi appeared to be unable to tolerate the high concentration of heavy oil (~ 150 g/l), as shown by their low cell number densities (Table 1).

The heavy oil used in this study has very low water solubility, thus resulting in very low bioavailability for microbial degradation. Indeed, the microbial activity on heavy oil could not increased by refeeding with yeast extract after 36 days (Table 1). However, a visual observation showed that heavy oil was emulsified rapidly after adding 1 g/l yeast extract to the solution. Subsequently, the micelles of oil formed in the solution at day 64, followed by the fungal growth (Table 1). Up to 105 days of incubation, the heavy oil, as shown to be brown-black color floating on the surface water, remained constantly and appeared to be entrapped on oil-fungal cell complexes on the surface water to be a solid material. In this case, fungi acted to take oil for their growth by making soil plates (figure not shown). Furthermore, heavy oil appeared to be invisible after 239 days of incubation. In addition, the yeast extract is required for bacterial strain Pseudomonas aeruginosa used in this study as co-substrate (a source of cellular nitrogen) to produce high extracellular emulsifying activity in utilizing the heavy oil as carbon source [6, 7], so that biodegradation of heavy oil (in particular aliphatic hydrocarbons) sustained (Fig. 3). The addition of yeast extract resulted in the apparent aqueous solubility of heavy oil that was visually observed in liquid medium (figure not shown) and the production of extracellular emulsifying agent (e.g., EPS) that might be associated with hydrocarbon utilization [12, 13] by the bacterial cells, as shown by TEM images (Fig. 1). Since the low bioavailability of heavy oil is a major rate-limiting factor in its degradation by bacteria [14, 15], the bacterial extracellular products might have resulted in increased bioavailability of heavy oil, thus facilitating hydrocarbon uptake by bacteria and sustaining biodegradation of heavy oil.

The GC analyses showed that the major products of the oxidation were both nonpolar and polar metabolites (Figs. 3-5). The oxidized oil metabolites also have a higher aqueous solubility, resulting in the increased bioavailability of these compounds so that they can be easily mineralized by microorganisms. In general, the biodegradation of oil components usually occurs in the following order: alkanes, branched alkanes, the aromatic compounds and finally cycloalkanes [e.g., 16], where their biodegradation rate depends on their concentration and the time of exposure. The lack of degradation of aromatic compounds by the indigenous microorganisms here may be attributed to inhibition by aromatic hydrocarbon [17]. The characterization of our bacterial strain used in this study has shown that the strain is able to grow well on aliphatic hydrocarbons but not on aromatic hydrocarbons [4, 5]. In this study, either bacteria or fungi have the low ability to degrade aromatic hydrocarbons, as shown in Figure 4. But they are capable of degrading n-alkanes (Fig. 3). However, the presence of both bacteria and fungi as indigenous mixed microbial consortia suggests that they may contribute to the degradation of heavy oil from the Nakhodka oil spill. Thus, the present study provides a significant evidence for bioremediation of oil spill (as complex hydrocarbon mixture) by indigenous microbial consortia, since studies on the utilization of complex hydrocarbon mixture by microbial consortia are few [17, 18, 19, 20].


This study has demonstrated that the long-term biodegradation of hydrocarbon mixtures (aliphatic and aromatic hydrocarbons) by indigenous mixed microbial consortia can involve the succession of different microorganisms with respect to the pH of the aqueous phase. In addition, the successful oil bioremediation is very dependent on the benevolent and favorable interactions between indigenous microorganisms (herein bacteria and fungi) in contaminated sites to fully biodegrade all hydrocarbon components of oil spill.


We would like to thank Dr. Yousuke Degawa (Kanagawa Prefectural Museum of Natural History, Japan) for helping with the fungal identification, Prof. Dr. Takashi Hasegawa (Kanazawa University) for helpful discussions, and all the students of the Tazaki laboratory and the Hasegawa laboratory of Kanazawa University for their cooperation and assistance. We also thank the anonymous reviewers for their constructive comments. SKC would like to express her thanks for the scholarship provided by the MONBUKAGAKUSHO during her study at Kanazawa University (Japan) and for the IMC16/IFM scholarship provided by the Kazato Research Foundation (JEOL Group). This study was funded by a grant from the Japanese Ministry of Education, Culture, Sports, Science and Technology to KT.


[1] Grady, Jr. C.P.L. and Lim, H.C., 1980, Biological Wastewater Treatment: Theory and Applications, Marcel Dekker, New York, USA, pp. 197-228.

[2] Tazaki, K., (editor), 2003, Heavy Oil Spilled from Russian Tanker “Nakhodka” in 1997: Towards Eco-responsibility Earth Sense, 21st Century COE Kanazawa University, Kanazawa University Press, Kanazawa, Japan.

[3] Ewies, J.B., Ergas, S.J., Chang, D.P.Y. and Schroeder, E.D., 1998, Bioremediation principles, McGraw-Hill Inc., Singapore, pp. 66-98.

[4] Chaerun, S.K., Tazaki, K., Asada, R., and Kogure, K., 2004, “Bioremediation of coastal areas 5 years after the Nakhodka oil spill in the Sea of Japan: isolation and characterization of hydrocarbon-degrading bacteria,” Environment International, 30, pp. 911-922.

[5] Chaerun, S.K., Tazaki, K., Asada, R., and Kogure, K., 2004, “Alkane-degrading bacteria and heavy metals from the Nakhodka oil spill-polluted seashores in the Sea of Japan after five years of bioremediation,” The Science Reports of Kanazawa University, 49, pp. 25- 46.

[6] Chaerun, S.K., Tazaki, K., Asada, R., and Kogure, K., 2005, “Interaction between clay minerals and hydrocarbon-utilizing indigenous microorganisms in high concentration of heavy oil: Implications for bioremediation,” Clay Minerals, 40, pp. 105-114.

[7] Chaerun, S.K., Tazaki, K., and Asada, R., 2003, “Double function of bentonite and kaolinite as adsorbents and microbial growth-support media for degradation of crude oil,” Heavy Oil Spilled from Russian Tanker Nakhodka in 1997: Towards Eco-responsibility Earth Sense, K. Tazaki ed., 21st Century COE Kanazawa University, Kanazawa University Press, Kanazawa, Japan, pp. 253-277.

[8] Chaerun, S.K., and Tazaki, K., 2003, “Hydrocarbon-degrading bacteria in the heavy oil polluted soil and seawater after 5 years bioremediation,” Water and Soil Environments: Microorganisms Play An Important Role, K. Tazaki, ed., 21st Century COE Kanazawa University, Kanazawa University Press, Kanazawa, Japan, pp. 187- 204.

[9] Suzuki, T., Shibata, M., Tanaka, K., Tsuchida, K., and Toda, T., 1995, “A new drying method: low vacuum SEM freeze drying and its application to plankton observation,” Contribution to the Bulletin of Planktonic Society of Japan, 42, pp. 53-62.

[10] Gaudy Jr., A.F., and Gaudy, E.T., 1981, Microbiology for Environmental scientists and Engineers, McGraw-Hill, New York, USA, pp. 175-206.

[11] Widdel, F., 1988, “Microbiology and ecology of sulfate- and sulfur-reducing bacteria,” Biology of Anaerobic Microorganisms, A.J.B. Zehnder, ed., Wiley, New York, USA, pp. 469-585.

[12] Berg, G., Seech, A.G., Lee, H., and Trevors J.T., 1990, “Identification and characterization of a soil bacterium with extracellular emulsifying activity,” Journal of Environmental Science and Health, A25 (7), pp. 753-764.

[13] Cooper, D.G., 1986, “Biosurfactant,” Microbiological Sciences, 3, pp. 145-149.

[14] Stucki, G., and Alexander, M., 1987, “Role of dissolution rate and solubility in biodegradation of aromatic compounds,” Appl. Environ. Microbiol., 53, pp. 292-297.

[15] Volkering, F., Breure, A.M., Sterkenburg, A., and van Andel, J.G., 1992, “Microbial degradation of polycyclic aromatic hydrocarbons: effect of substrate availability on bacterial growth kinetics,” Appl. Microbiol. Biotechnol., 36, pp. 548-552. 30 S.K. Chaerun, Ryuji Asada and Kazue Tazaki

[16] Sugiura, K., Ishihara, M., Schimauchi, T., Harayama, S., 1997, “Physicochemical properties and biodegradability of crude oil,” Environ. Sci. Tech., 31, pp. 45-51.

[17] Richard, J.Y., and Vogel, T.M., 1999, “Characterization of a soil bacterial consortium capable of degrading diesel fuel,” International Biodeterioration & Biodegradation, 44, pp. 93-100.

[18] Arvin, E., Jensen, B.K., and Gunderssen, A.T., 1989, “Substrate interactions during aerobic biodegradation of benzene,” Appl. Environ. Microbiol., 55, pp. 3221-3225.

[19] Song, H.G., Wang, X., and Bartha, R., 1990, “Bioremediation potential of terrestrial fuel spills,” Appl. Environ. Microbiol., 56, pp. 652-656.

[20] Alvarez, P.J.J., and Vogel, T.M., 1991, “Substrate interactions of benzene, toluene and para-xylene during microbial degradation by pure cultures and mixed culture aquifer slurries,” Appl. Environ. Microbiol., 57, pp. 2981-2985.

Siti Khodijah Chaerun (a) *, Ryuji Asada (b) and Kazue Tazaki (b)

(a) School of Life Sciences and Technology, Bandung Institute of Technology Ganesa 10, Bandung 40132, Indonesia

(b) Department of Earth Sciences, Faculty of Science, Kanazawa University Kakuma, Kanazawa, Ishikawa 920-1192, Japan

Table 1: Physico-chemical and biological parameters during the

biodegradation of heavy oil.

Parameters 0 day 36 days

pH 7.8 [+ or -] 0.0 7.7 [+ or -] 0.2

Eh 179 [+ or -] 1.5 –

Microbial growth (x [10.sup.5]

CFU/ml) 53 [+ or -] 10 65 [+ or -] 5.7

Predominant microorganism Bacteria bacteria

Percentage of bacteria 100% 99%

Percentage of fungi 0% 1.5%

Parameters 64 days 105 days

pH 3.7 [+ or -] 0.2 3.4 [+ or -] 0.1

Eh – –

Microbial growth (x [10.sup.5]

CFU/ml) 25 [+ or -] 7.3 9 [+ or -] 2.2

Predominant microorganism fungi fungi

Percentage of bacteria 0.5% 0.5%

Percentage of fungi 99.5% 99.5%

Parameters 239 days 429 days

pH 2.0 [+ or -] 0.3 1.9 [+ or -] 0.1

Eh 248 [+ or -] 6.0 362 [+ or -] 2.0

Microbial growth (x [10.sup.5]

CFU/ml) 8 [+ or -] 3.1 4 [+ or -] 1.0

Predominant microorganism fungi fungi

Percentage of bacteria 0% 0%

Percentage of fungi 100% 100%

(Mean [+ or -] SD, n = 3); -: not measured

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