Evaluation of Alcaligenes faecalis Degradation of Chrysene and Diesel Oil with Concomitant Production of Biosurfactant
Alcaligenes faecalis was evaluated for its potential to degrade varying concentrations of chrysene and diesel oil with concomitant biosurfactant production. Biodegradation was set up for 7 days utilizing the substrates as sole carbon and energy sources. Residual chrysene obtained after degradation of 30, 50 and 100 mg L-1, respectively was 17.4±1.5, 27.2±1.2 and 28.7±1.4 mg L-1 while total petroleum hydrocarbon remaining after degradation of 3, 5, 15 and 30% (v/v) diesel oil respectively was 2.58±0.5, 3.09±1.2, 21.65±5.4 and 63.92±8.1%. Microbial cells of A. faecalis and sterilized cell-free extract from diesel oil media showed emulsifying activities against kerosene, diesel oil, engine oil, hexadecane, dodecane, xylene and hexane whereas no emulsifying activity was observed of microbial cells and sterilized cell-free extract from chrysene media. Alcaligenes faecalis cells harvested from diesel oil media also showed haemolytic activity unlike the microbial cells from chrysene media. Growth of the isolate in chrysene and diesel oil media induced secretion of protein and carbohydrate into the media which were statistically significantly (p<0.05) different compared to controls. This study portrays the potential of Alcaligenes faecalis to degrade and grow on chrysene and diesel oil and induce extracellular protein and carbohydrate with concomitant production of biosurfactant for industrial purposes and in hydrocarbon bioremediation.
Environmental pollution by crude or refined petroleum products is a global
problem. Soil fertility problems arising from oil impact, leads to reduction
and deformation of agricultural produce (Nwachukwu et
al., 2001). Domestic water supplies may become contaminated by run offs
from agricultural soils polluted with crude oil. Petroleum hydrocarbon contamination
is also of great concern due to the toxicity and recalcitrance of many fuel
components and products such as polycyclic aromatic hydrocarbons (Saeed
and Al-Mutairi, 2000; Xue and Warshawsky, 2005;
Masih and Taneja, 2006). These thus warrant the need
to remediate petroleum hydrocarbon polluted environment and formulate bioremediation
protocols that would be safe and cost effective. Polycyclic Aromatic Hydrocarbons
(PAHs) include the high and low molecular weight PAHss such as chrysene, pyrene,
fluoranthene, benzanthracene, benzopyrene, phenanthrene, anthracene and fluorene
(Kanaly and Harayama, 2000).
Bioremediation of petroleum and PAHs contaminated environment is a promising
alternative remedial strategy (Gogoi et al., 2003;
Okoh and Trejo-Hernandez, 2006). Crude oil polluted
agricultural soil has been found to be bioremediated by application of a soil
bacterium and inorganic nutrient (Nwachukwu, 2001).
Dean-Ross (2005) conducted slurry bioreactors bioremediation
batch experiment on pristine sediment spiked with phenanthrene, anthracene,
pyrene and fluoranthene and observed that bioaugmentation with Rhodococcus
sp., resulted in enhanced bioremediation rates and yields.
One main factor that influences the extent of petroleum, aliphatic hydrocarbons
and PAHs biodegradation is their bioavailability; this is a priority research
objective in the bioremediation field (Makkar and Rockne,
2003). Thus, approaches to enhancing these compounds biodegradation often
attempt to increase their apparent solubility by treatments such as addition
of synthetic surfactants and/or release of biosurfactants (Barkay
et al., 1999; Makkar and Rockne, 2003; Al-Turki,
2009). Biosurfactants function to reduce surface tension and interfacial
tensions between individual molecules at the surface and interface, respectively
(Rahman et al., 2002; Lu
et al., 2007). They also exhibit emulsification properties forming
micelles whereby hydrocarbons become solubilized in the hydrophobic cores of
the micelles thus making them available for microbial degradation (Bento
et al., 2005; Zhang et al., 2005).
Many bacteria growing on alkanes produce biosurfactant to increase the bioavailability
of these poorly available substrates (Sepahy et al.,
2004; Bento et al., 2005). A similar strategy
was suggested for PAH degrading bacteria (Deziel et al.,
1996). However, in other studies, no correlation was found between biosurfactant
productions and PAH mineralization or dissolution rates (Volkering
et al., 1992; Willumsen and Karlson, 1997).
It has therefore become imperative to study the mechanisms of bioavailability of hydrocarbon pollutants to individual microorganisms. In addition, for petroleum bioremediation to be a viable alternative to non-biological remediation techniques, the search for microorganisms with degradation capabilities for petroleum products and components which are recalcitrant and poorly soluble would be a continual process. In this study, we report the potential of A. faecalis degradation of varying concentrations of chrysene and diesel oil with concomitant production of biosurfactant.
MATERIALS AND METHODS
This study was carried out from June 2006 to 2007 in Biochemistry Department, College of Medicine, University of Lagos, Nigeria.
The bacterium A. faecalis was previously isolated in our Laboratory
by chrysene enrichment from Nigerian PAHs polluted soil sites (Igwo-Ezikpe
et al., 2006). Chrysene enrichment is an isolation technique whereby
chrysene was used as the sole carbon and energy source in Mineral Salt Medium
(MSM) to isolate chrysene degraders. The isolate was maintained on nutrient
agar slants and stored at 4°C when not in use.
Chemicals and Media
All chemicals were of analytical grade. Mineral Salt Media (MSM) components
were supplied by (Sigma, Germany). Hydrocarbons used in this study include chrysene
(Sigma, Germany, 98% purity), ethyl acetate, dodecane, hexane, xylene and hexadecane
(BDH, England). Diesel oil, kerosene and engine oil were obtained from African
Petroleum (Nigeria). Nutrient agar for bacterial enumeration was purchased from
Sigma (Germany) and blood agar plates for haemolytic assay were prepared using
bacteriological agar (Oxoid, Germany) in our laboratory.
Preparation of Starter Culture
The MSM composed per liter (pH 7.2): NH4NO3, 4.0 g;
Na2HPO4, 2.0 g; KH2PO4, 0.53 g;
K2SO4, 0.17 g, MgSO4.7H2O, 0.10
g and trace elements solution (1 ml L-1) as described by Ilori
and Amund (2000). The MSM was sterilized by autoclaving at 121°C for
20 min. The isolate being a chrysene degrader was precultured in 3% diesel oil
prior to use to ensure adaptation to diesel oil. This was achieved by growing
bacterial cells in sterilized 50 mL MSM containing 3% diesel oil for 7 days
at 30±0.2°C. The cells were thereafter cold harvested from the medium
by centrifugation at 4,000x g, 4°C for 20 min and cultured on nutrient agar
plate 48 h. Cell were harvested from surface of nutrient agar, suspended in
10 mL phosphate buffer (50 mM, pH 7.2) in a test tube, centrifuged at 10,000x
g, 4°C for 10 min. The supernatant discarded while pelleted cells were resuspended
in the same buffer and washed twice. The pelleted cells were finally suspended
in phosphate buffer, optical density (OD600nm) adjusted to 0.4.
The bacterial isolate was evaluated for the ability to degrade varying concentrations
of chrysene (30, 50 and 100 mg L-1) and diesel oil (3, 5, 15 and
30% v/v) as sole carbon and energy source. The various concentrations of chrysene
were dissolved in Erlenmeyer flasks (250 mL) containing 5 mL ethyl acetate and
evaporated, 20 mL MSM added and foiled to prevent photolysis. Media were sterilized
by autoclaving at 121°C for 20 min. Starter cultures (10 mL) were thereafter
inoculated into the media and incubated aerobically at 30±0.2°C and
agitated at 150 rpm. This set up was designated Experimental (E). Two controls
(C1 and C2) were also included; C1 consisted of the same materials present in
E but without the carbon source while C2 contained all the materials in E with
no test isolate inoculated.
Biodegradation was assayed by determining the residual chrysene or Total Petroleum
Hydrocarbon (TPH) remaining and growth profile of the isolate. Residual chrysene
was estimated after extracting twice with equal volume of ethyl acetate and
analyzed using High Performance Liquid Chromatography (HPLC). The HPLC analyses
were performed with VYDAC RP C18 reverse phase column (250x0.4 mm). Separation
was achieved by gradient elution in acetonitrile: water (60, 50, 40, 30, 20,
10 and 0% water), temperature 25°C, with a flow rate of 0.8 mL min-1
and UV absorbance detector set at 254 nm. Estimation of diesel oil degradation
was only at the end of 7 days. Diesel oil was extracted from media by mixing
with equal volume of n-hexane and filtered through glasswool in a funnel. The
extraction procedure was repeated three times and the extracts were pooled.
The extracts were analyzed using a gas chromatography-mass spectrometry (Model:
Agilent Tec 7890A, USA) and HP5-MS column (30 mx0.25 μmx0.25 μm) using
helium as the carrier gas. The temperature was programmed to vary linearly from
40 to 270°C at the rate of 10°C min-1 and maintained for
20 min. Total Petroleum Hydrocarbon (TPH) was measured as the sum of all the
peak areas on the gas chromatogram to obtain quantitative data. The degree of
degradation (%) was calculated as described by Lee et
al. (2005). The area of TPH from the experimental media was divided
by the area of TPH from control C2 and multiplying by 100. Growth profile of
the isolate was determined by measuring the Total Viable Counts (TVC) and population
density of the isolates. The TVC was enumerated by spread-plate technique whereby
the inoculum from various media was cultured on nutrient agar for 48 h and microbial
cells counted. Optical density of the media was measured spectrophometerically
(OD600nm) to determine the population density of the inoculum.
Emulsifying Activity Measurement
The various experimental media and controls were made cell free after 7
days of incubation by centrifugation at 10,000x g, 4°C for 10 min and heat
sterilized. Cell-free extracts (2 mL) were vortex with n-hexadecane (2 mL) in
a test tube and left undisturbed for 24 h. The emulsification index (E24) was
calculated as the percentage of the height of emulsified layer (cm) divided
by total height of liquid column (cm) (Ilori and Amund,
2001; Moraes et al., 2002). Emulsifying activity
was also measured against different hydrophobic sources (kerosene, diesel oil,
engine oil, hexadecane, dodecane, xylene and hexane). This activity measurement
was repeated using the microbial cells.
This was undertaken to confirm the production of biosurfactant by A.
faecalis. The bacterial cells from MSM chrysene and diesel oil growth media
were plated onto blood agar and incubated at 37°C for 48 h (Bicca
et al., 1999).
Determination of Total Protein and Carbohydrate Content of Cell-Free Extracts
The total protein content of the cell-free extracts was assayed by the Bradford
method as described by Sepahy et al. (2004) using
bovine serum albumin as standard. The reaction mixture contained 0.1 mL of cell-free
extract to which 5.0 mL of coomassie blue dye reagent is added, incubated for
5 min and absorbance read at 595 nm. The total carbohydrate contents of cell-free
extracts were estimated by anthrone method using glucose as standard (Jayaraman,
1981). A volume of 0.5 mL of cell-free extract was mixed with 2.0 mL of
anthrone solution (2% anthrone reagent in conc H2SO4),
kept in boiling water bath for 10 min. Cooled to room temperature and absorbance
read at 620 nm against reagent blank. The total protein and carbohydrate content
in test samples and controls were extrapolated from standard calibration curves.
The experiments were set up in duplicate and analysis carried out in triplicates
of each set up. Results are expressed as Mean±SEM (Standard Error of
Mean). Statistically significant difference (p<0.05) was determined using
analysis of variance (ANOVA). All data were analyzed using Statistical Package
for the Social Science15.0 for windows (SPSS 15.0).
RESULTS AND DISCUSSION
Alcaligenes faecalis used in this study is a Gram negative rod previously
identified as chrysene degrading bacterium with potential to degrade petrochemical
products such as diesel oil, engine oil, crude oil and kerosene of Nigerian
origin (Igwo-Ezikpe et al., 2006). The isolate
was able to degrade varying concentrations of chrysene and diesel oil utilizing
them as sole carbon and energy source. Residual chrysene obtained after 7 days
degradation of 30, 50 and 100 mg L-1, respectively was 17.4±1.5,
27.2±1.2 and 28.7±1.4 mg L-1 while THC remaining after
7 days degradation of 3, 5, 15 and 30% (v/v) diesel oil respectively were 2.58±0.5,
3.09±1.2, 21.65±5.4 and 63.92±8.1% (Fig.
1). No significant (p<0.05) loss of the carbon sources was obtained in
C2 controls indicating that the isolate mediated the degradation process. This
corroborates earlier findings (Nwachukwu et al., 2001; Lee
et al., 2005).
Residual chrysene and (b) total petroleum hydrocarbon remaining after
A. faecalis degradation of varying concentration of chrysene and
profile of A. faecalis during degradation of varying concentration
(a) chrysene and (b) diesel oil
Growth profile of the isolate on the various tested concentrations of chrysene
and diesel oil was found to be concentration dependent (Fig. 2).
Alcaligenes faecalis grown on 30, 50 and 100 mg L-1 chrysene
yielded highest cell densities of 6.3x108, 9.8x108 and
9.2x105 cfu mL-1, respectively after 7 days of incubation.
||(a) Emulsifying activity of microbial cells and (b) cell-free
extract of A. faecalis grown on diesel oil
The organism was able to adapt and grew on the various concentrations of chrysene,
multiplying in cell density and increasing the turbidity of the media compared
to non-chrysene growth media. However, bacterial growth was best supported by
50 mg L-1 chrysene under the studied conditions. Previous studies
have shown microbial growth and degradation of different concentrations of PAHs
ranging from 5 to 100 mg L-1 (Yu et al.,
2005; Igwo-Ezikpe et al., 2007). Highest cell
densities of 3.2x1011, 3.2x1014, 2.5x 1012
and 9.9x105 cfu mL-1 were, respectively observed for 3,
5, 15 and 30% (v/v) diesel oil media. The isolate utilized various diesel oil
as the sole carbon and energy but as the concentration of diesel oil increased,
microbial cell generation decreased. However, 5% diesel oil best supported cell
mass generation. Possible explanation for increasing concentration of hydrocarbon-related
inhibition of microbial growth include decreased bioavailability of micelle-solubilized
hydrocarbon to support microbial growth or hydrocarbon interference with microbial
cell integrity (Sikkema et al., 1995). This may
explain why crude oil and its petrochemical products cause sterility of agricultural
soils and persist in the environment. The observed growth and degradation potential
of the isolate on various concentrations of chrysene and diesel oil may be as
a result of its previous exposure to the pollutants.
Microbial cells and cell-free extracts of the various diesel oil growth media
showed emulsifying activity against various hydrocarbons such as kerosene, diesel
oil, engine oil, hexadecane, dodecane and xylene (Fig. 3).
This may indicate the production of biosurfactant by the isolate when grown
on diesel oil. Earlier study has shown the emulsification of crude oil and other
hydrocarbons by biosurfactant producing bacteria (Ilori
and Amund, 2001; Rahman et al., 2002; Pornsunthorntawee
et al., 2008). The emulsifying activities were found to be dependent
on the test hydrocarbons and the source of the cell-free extract. Microbial
cells and cell-free extracts from 3 and 5% diesel oil showed better emulsifying
activities than those from 15 and 30% suggesting increased constrain of microbial
assess and poor solubilization of 15 and 30% diesel oil. Emulsifying activities
of microbial cells from the various diesel oil media were higher than that of
cell-free extracts. This may indicate that bioemulsifier of A. faecalis
synthesized using diesel oil as sole carbon source may be cell-bound and only
partially released into the liquid phase of the media. This corroborates the
finding of previous research where biosurfactant activity was mediated by microbial
cell mediated and/or released into growth media (Gutnick
et al., 2003; Bento et al., 2005).
In contrast, the microbial cells and cell-free extracts from chrysene media
did not show emulsifying activities against the various hydrocarbons tested.
This indicated that the organism may not have produced bioemulsifier using chrysene
as the sole carbon and energy source under the specified conditions of growth.
This is in accordance with earlier study which reported that production of biosurfactants
is largely dependent on the carbon source (Maneerat, 2005;
Abouseouda et al., 2008).
Haemolysis is one of the properties of biosurfactants (Tabatabaee
et al., 2005). Zones of clearance on blood agar plates as a result
of haemolysis have been used as a rapid method for screening microorganisms
for potential biosurfactant production (Lin, 1996). Cleared
zones on blood agar plates were observed by microbial cells cultivated from
diesel oil media (Fig. 4). This may further confirm biosurfactant
production of the isolate when grown on diesel oil. However, haemolytic activity
was also not observed of bacterial cells and cell-free extracts from chrysene
growth media (Fig. 4).
Furthermore, growth of A. faecalis in the various chrysene and diesel
oil growth media induced the secretion of extracellular protein and carbohydrate
(Fig. 5, 6). There was statistical significant
difference (p<0.05) in the total protein and carbohydrate content of chrysene
and diesel oil growth media compared to controls non-carbon media extracts.
Results showed that an increase in chrysene and diesel oil concentrations induced
relative increase in secretion of extracellular carbohydrate into the media.
This may be as a result of increased toxicity of the hydrocarbons on microbial
cell membrane leading to leaching of cell constituents and/or isolates response
to hydrophobic environment by synthesis and release of biosurfactant components.
Since, there was no carbohydrate in the media, its appearance and increase in
response to hydrocarbon concentration may be due to the nature of the biosurfactant
released by A. faecalis, this corroborates previous research (Bicca
et al., 1999). Previous studies have also shown hydrocarbon-degrading
bacteria to produce extracellular emulsifying agents, generally consisting polysaccharides
associated with proteins. Some examples included emulsan, the exopolysaccharide
from Acinetobacter, alasan produce by
||Alcaligenes faecalis harvested from chrysene and diesel
oil media growth on blood agar plate
||Total protein of cell-free extract from (a) chrysene and (b)
diesel oil media
||Total carbohydrate of cell-free extract from (a) chrysene
and (b) diesel oil media
Acinetobacter radioresistens KA53, rhamnolipid and glycolipid from Pseudomonas
sp. (Toren et al., 2001; Abdel-El-Haleem,
2003; Gutnick et al., 2003). The combination
of hydrophilic sugar main chain repeat units and the hydrophobic side groups
of biosurfactants lead to their amphipathic behavior and therefore, their ability
to form stable oil-in-water emulsions and reduce surface tension. In addition,
the biosynthesis and excretion of biosurfactants into medium are considered
to be cell mechanisms aimed at an adaptation of microorganism to using external
lipophilic compounds as carbon and energy sources (Barkay
et al., 1999; Maneerat, 2005). As such, the
response of the isolate in terms of possible biosurfactant production may be
as a result of external factor on the microbial genetic make-up leading to the
observed expression of macromolecules.
This study shows the potential of A. faecalis to adapt and degrade varying concentrations of chrysene and diesel oil, produce biosurfactant using diesel oil as carbon source and possible use in hydrocarbon bioremediation studies. Non-emulsification of the microbial cells and cell-free extracts from chrysene media may not totally rule out its biosurfactant production under this condition. Therefore, other parameters such as surface tension and adhesion may be investigated. We suggest that chrysene and diesel oil when used as sole carbon and energy source induced secretion of extracellular protein and carbohydrate into the growth medium.
We wish to thank Mr. Sunday O. Adenikan, Technologist, Department of Biochemistry, College of Medicine, University of Lagos. He assisted in the biochemical quantification of total protein and carbohydrate.
1: Abdel-El-Haleem, D., 2003. Acinetobacter: Environmental and biotechnology applications. Afr. J. Biotechnol., 2: 71-74.
2: Abouseouda, M., R. Maachi, A. Amranec, S. Boudergua and A. Nabia, 2008. Evaluation of different carbon and nitrogen sources in production of biosurfactant by Pseudomonas fluorescens. Desalination, 223: 143-151.
3: Al-Turki, A.I., 2009. Microbial polycyclic aromatic hydrocarbons degradation in soil. Res. J. Environ. Toxicol., 3: 1-8.
CrossRef | Direct Link |
4: Gutnick, D., H. Bach and Y. Berdichevsky, 2003. An exocellular protein from the oil-degrading microbe Acinetobacter venetianus RAG-1 enhances the emulsifying activity of the polymeric bioemulsifier emulsan. Applied Environ. Microbiol., 69: 2608-2615.
5: Barkay, T., S. Navon-Venezia, E. Ron and E. Rosenberg, 1999. Enhancement of solubilisation and biodegradation of polycyclic aromatic hydrocarbon by the bioemulsifier Alasan. Applied Environ. Microbiol., 65: 2697-2702.
6: Bento, F.M., F.A.D.O. Camargo, B.C. Okeke and W.T. Frankenberger, 2005. Diversity of biosurfactant producing microorganisms isolated from soils contaminated with diesel oil. Microbiol. Res., 160: 249-255.
7: Bicca, F.C., L.C. Fleck and M.A.Z. Ayub, 1999. Production of biosurfactant by hydrocarbon degrading Rhodococcus rubber and Rhodococcus erythropolis. Rev. Microbiol., 30: 231-236.
8: Dean-Ross, D., 2005. Biodegradation of selected PAH from sediment in bioslurry reactors. Bull. Environ. Contam. Toxicol., 74: 32-39.
9: Ziel, E., G. Paquette, R. Villemur, F. Lepine, J. Bisaillon, 1996. Biosurfactant production by a soil Pseudomonas strain growing on polycyclic aromatic hydrocarbons. Applied Environ. Microbiol., 62: 1908-1912.
PubMed | Direct Link |
10: Gogoi, B.K., N.N. Duttaa, P. Goswamia and K.T.R. Mohanb, 2003. A case study of bioremediation of petroleum-hydrocarbon contaminated soil at a crude oil spill site. Adv. Environ. Res., 7: 767-782.
11: Igwo-Ezikpe, M.N., O.G. Gbenle and M.O. Ilori, 2006. Growth study on chrysene degraders isolated from polycyclic aromatic hydrocarbon polluted soils in Nigeria. Afr. J. Biotechnol., 5: 823-828.
Direct Link |
12: Ilori, M.O.N. and D.I. Amund, 2001. Production of peptidoglycolipid bioemulsifier by Pseudomonas aeruginosa grown on hydrocarbons. Z. Naturforsch. C, 56: 547-552.
13: Ilori, M.O. and D.L. Amund, 2000. Degradation of anthracene by bacteria isolated from soil polluted tropical soils. Z. Naturforsch. C, 55: 890-897.
14: Jayaraman, J., 1981. Laboratory Manual in Biochemistry. Wiley Eastern Ltd., New Delhi, India, pp: 53
15: Kanaly, R.A. and S. Harayama, 2000. Biodegradation of high-molecular-weight polycyclic aromatic hydrocarbons by bacteria. J. Bacteriol., 182: 2059-2067.
Direct Link |
16: Lee, M., M.K. Kim, M. Kwon, B.D. Park, M.H. Kim, M. Goodfellow and S. Lee, 2005. Effect of the synthesized mycolic acid on the biodegradation of diesel oil by Gordonia nitida strain LE31. J. Biosci. Bioeng, 100: 429-436.
17: Lin, S.C., 1996. Biosurfactants: Recent advances. J. Chem. Tech. Biotechnol., 66: 109-120.
18: Lu, J.R., X.B. Zhao and M. Yaseen, 2007. Biomimetic amphiphiles: Biosurfactants. Curr. Opin. Colloid Interface Sci., 12: 60-67.
CrossRef | Direct Link |
19: Makkar, R.S. and K.J. Rockne, 2003. Comparison of synthetic surfactants and biosurfactants in enhancing biodegradation of polycyclic aromatic hydrocarbons. Environ. Toxicol. Chem., 22: 2280-2292.
20: Maneerat, S., 2005. Biosurfactants from marine microorganisms. Songklanakarin J. Sci. Technol., 27: 1263-1272.
Direct Link |
21: Masih, A. and A. Taneja, 2006. Polycyclic Aromatic Hydrocarbons (PAHs) concentrations and related carcinogenic potencies in soil at a semi-arid region of India. Chemosphere, 65: 449-456.
Direct Link |
22: Moraes, I.D., M. Benincass and R.M. Alegre, 2002. Production and characterization of rhamnolipids produced by a newly isolated strain of Pseudomonas aeruginosa. Braz. J. Food Technol., 5: 145-149.
Direct Link |
23: Nwachukwu, S.C.U., P. James and T.R. Gurney, 2001. Impact of crude oil on the germination and growth of cress seeds (Lepidium sp.) after bioremediation of agricultural soil polluted with crude petroleum using adapted Pseudomonas putida. J. Environ. Biol., 22: 29-36.
24: Nwachukwu, S.C.U., 2001. Bioremediation of sterile agricultural soils polluted with crude petroleum by application of the soil bacterium, Pseudomonas putida, with inorganic nutrient supplementations. Curr. Microbiol., 42: 231-236.
CrossRef | Direct Link |
25: Okoh, A.I. and M.R. Trejo-Hernandez, 2006. Remediation of petroleum hydrocarbon polluted systems: Exploiting the bioremediation strategies. Afr. J. Biotechnol., 5: 2520-2525.
Direct Link |
26: Pornsunthorntawee, O., P. Wongpanit, S. Chavadej, M. Abe and R. Rujiravanit, 2008. Structural and physicochemical characterization of crude biosurfactant produced by Pseudomonas aeruginosa SP4 isolated from petroleum-contaminated soil. Bioresour. Technol., 99: 1589-1595.
CrossRef | PubMed |
27: Rahman, K.S.M., T.J. Rahman, P. Lakshmanaperumalsamy, R. Marchant and I.M. Banat, 2003. The Potential of Bacterial Isolates for Emulsification with a Range of Hydrocarbons. Acta Biotechnol., 23: 335-345.
28: Saeed, T. and M. Al-Mutairi, 2000. Comparative composition of polycyclic aromatic hydrocarbons (PAHs) in the sea water-soluble fractions of different Kuwaiti crude oils. Adv. Environ. Res., 4: 141-145.
29: Sepahy, A.A., M.M. Assadi, V. Saggadian and A. Noohi, 2004. Production of biosurfactant from Iranian oil fields by isolated Bacilli. Int. J. Environ. Sci. Technol., 1: 287-293.
Direct Link |
30: Sikkema, J., J.A. de Bont and B. Poolman, 1995. Mechanisms of membrane toxicity of hydrocarbons. Microbiol. Mol. Biol. Rev., 59: 201-222.
PubMed | Direct Link |
31: Tabatabaee, A., M.A. Mazaheri, A.A. Noohi and V.A. Sajadian, 2005. Isolation of biosurfactant producing bacteria from oil reservoirs. Iran. J. Environ. Health Sci. Eng., 2: 6-12.
Direct Link |
32: Toren, A., S. Navon-Venezia, E.Z. Ron and E. Rosenberg, 2001. Emulsifying activities of purified alasan proteins from acinetobacter radioresistens ka53. Applied Environ. Microbiol., 67: 1102-1106.
33: Volkering, F., A.M. Breure, A. Sterkenburg and J.G. van Andel, 1992. Microbial degradation of polycyclic aromatic hydrocarbons: Effect of substrate availability on bacterial growth kinetics. Applied Microbiol. Biotechnol., 36: 548-552.
34: Willumsen, P.A. and U. Karlson, 1996. Screening of bacteria, isolated from PAH-contaminated soils, for production of biosurfactants and bioemulsifiers. Biodegradation, 7: 415-423.
CrossRef | Direct Link |
35: Xue, W. and D. Warshawsky, 2005. Metabolic activation of polycyclic and heterocyclic aromatic hydrocarbons and DNA damage: A review. Toxicol. Applied Pharmacol., 206: 73-93.
CrossRef | PubMed | Direct Link |
36: Yu, S.H., L. Ke, Y.S. Wong and N.F.Y. Tam, 2005. Degradation of polycyclic aromatic hydrocarbons by a bacterial consortium enriched from mangrove sediments. Environ. Int., 31: 149-154.
Direct Link |
37: Zhang, G., Y. Wu, X. Qian and Q. Meng, 2005. Biodegradation of crude oil by Pseudomonas aeruginosa in the presence of rhamnolipids. J. Zhejiang Univ. Sci. 6: 725-730.
Direct Link |
38: Igwo-Ezikpe, M.N., O.G. Gbenle and M.O. Ilori, 2007. Biodegradation of polycyclic aromatic hydrocarbons and petroleum products by Acinetobacter mallei. University of Lagos, Nigeria. http://www.unilag.edu.ng/publicationhtml/1978.html.