Iran is one of the world’s major oil producers and since the discovery and extraction of petroleum in the South West of the country in 1908, the environment of this vast area has encountered serious petroleum pollution. Besides this direct pollution, the occasionally pipeline leaks, transportation accidents, storage tank ruptures and refining petroleum is further intensifying the pollution of this area. Most of these compounds are considered as carcinogenic, mutagenic and potent immunotoxicants and classified as priority environmental pollutant by the US Environmental Protection Agency (1-4). Constituents of this contaminant have long-term effects on ecosystems and increase the presence of toxicants in organisms towards the top of the food chain. Destroyed ecosystems and health risks have emphasized on the necessity for clean up and remedies for soil and water, which have prompted many researchers to develop environmentclean up strategies, such as bioremediation (5, 6). Crude petroleum and its associated products contain many kinds of organic compounds, dominated by aliphatic and aromatic hydrocarbons. Biodegradation of hydrocarbon contaminated soils, using the potentials of microorganisms to degrade or detoxify these contaminants, has been demonstrated as cost-effective and an environmentally sound remedy (2). Petroleum-degrading microorganisms have enzyme systems and metabolic capabilities, enablethem to withstand adverse environmental conditions. Bacteria abundance, their fast growth and wide spectrum of molecules, have allowed them to be considered as a main bioremediation tool. Up to now, many bacterial strains, such as Pseudomonas, Alcanivorax, Acinetobacter, Rhodococcus, and Bacillus (7-9) have been isolated from soil and water, and were effectively used for the bioremediation of petroleum contaminants.
It is well established that many microbial catabolic pathways are involved in the degradation of petroleum (8). For example, the alk pathway in Rhodococcussp. Q15, which degrades C8 to C32 n-alkanes (10); Acinetobacter sp. M-1, which can degrade n-alkanes ranging from C13 to C44 (11); the OCT plasmid in Pseudomonas putida GPo1, which uses n-alkanes (12) and naphthalene (nah gene); or the xylE gene of P. putida involved in the meta-cleavage of catechol in aromatic compounds (13) have been documented in the literature. Molecular identification of the genes responsible for biodegradation processes using Polymerase Chain Reaction (PCR) and hybridization analyses may allow monitoring oil biodegradation at the DNA level (14). For an efficient bioremediation of contaminated sites, the first step is the isolation and characterization of indigenous strains that are adapted to the particular environmental conditions, such as temperature, water availability, oxygen and nutrients (7, 15). Also, long-term presence of petroleum contaminants in soil can develop microorganisms that are able to use organic pollutants as the sole carbon and energy source (6, 8, 10). Rhizosphere, the area of soil under the direct influence of roots, could be an appropriate microsite for in situ degradation of petroleum contaminated soils. Microbial density, diversity and activity in the rhizosphere soil are greater and more effective for bioremediation than non rhizosphere soils, likely due to the growth stimulation by the exudation of chemical compounds from the roots (16). It has been shown that microbial populations in the rhizosphere may enhance a plant’s adaptation to petroleum hydrocarbons through detoxifying contaminated soils as a result of direct mineralization of these organic contaminants (17). Wild oat (Avena fatua L.) is a member of grass family (Poaceae), with a fibrous root system which grows and completes its life cycle in the petroleum-contaminated soils of South West of Iran. It is hypothesized that wild oat might be able to benefit specific rhizobacterial communities in its highly branched root system, with a good potential for petroleum degradation. We focused on: (I) isolating rhizosphere-inhabiting indigenous oil-degrading bacteria in wild oat grown in petroleum-polluted areas; (II) characterizing isolates based on 16sRNA gene sequence. (III) monitoring catabolic pathways, and finally (IV) evaluating the efficiency of oil biodegradability in vitro by microbial isolates with chemical methods.
Present study aimed to make appropriate consortium for rhizosphere inoculation in newly contaminated sites feared to be further polluted in near future.
3. Materials and Methods
3.1. Soil Sampling and Analyzes
Wild oat rhizosphere-surrounding soil samples were gathered in persistently antecedently contaminated sites in Khuzestan (arid climate with less than annual 250 mm rainfall; Max.: 48°C and Min.: 4°C). Additionally, soil showed hyperthermic regime. Seven root samples were collected per plant, seven soil samples were gathered from contaminated uncovered areas. Soil-adhered roots were shaken 4 to 5 times, soil remained adhered to the roots was considered as rhizosphere soil (18). Non contaminated soil samples were used as control. Soil samples were sieved with a 2 mm mesh,stored at -20°C. Certain soil characteristics were measured such as pH, electricity conductivity (EC), and total petroleum hydrocarbon (TPH). 10 g of each contaminated soil sample was mixed with 10 g anhydrous Na2SO4, extracted with dichloromethane for 12 h using a Soxhlet apparatus (1), finally, the extract was evaporated and remaining weighed to quantify TPH.
3.2. Isolation and Characterization of Bacteria
One gram of each soil sample (roots, adhering rhizosphere) was added to the minimal Bushnell-Haas medium (BH) which contains followings (g/L): MgSO4 0.2, CaCl2 0.02, KH2PO4 1.0, K2HPO4 1.0, NH4NO3 1.0 and FeCl3 0.05 (19), supplemented with 2% to 5% (w/v) crude petroleum (API gravity: 31, Tehran Refinery) as a carbon source and selective substrate. Cultures were incubated at 30°C on a rotary shaker at 180 rpm for 7 days, and 20 μL of the resulting culture was sub-cultured in fresh BH medium. After five subsequent subcultures, cultures were purified on diesel agar and nutrient agar (20). The identification of isolates was performed based on 16S rRNA gene sequencing. The isolated strains were grown in nutrient broth and harvested from overnight cultures. DNA extraction was performed following the CTAB method (21). A 1500 bp region of 16S rDNA from each strain was amplified with 27F and 1492R degenerated primers (20). The PCR mixtures (50 μL) contained 1.5 mM MgCl2, 200 mM concentration of each deoxynucleoside triphosphate (dNTP), 50 pmol of each primer, 2.5 U of Taq DNA polymerase and 10 ng of DNA template. PCR amplification was performed using a PeQ STAR thermocycler (Biotechnologie GmbH, Germany). Forty cycles were performed at the following temperatures and times: 94ºC for 5 min, 94ºC for 30 sec, 56ºC for 45 sec, 72ºC for 1 min and 72ºC for 10 min. The amplicons were analyzed by electrophoresis in a 1% (w/v) agarose gel and purified using the Agarose Gel DNA Extraction Kit (Roche, Germany), then ligated into the pTZ57RT vector (K1214, Fermentas, Germany). The constructed plasmids were subsequently introduced into Escherichia coli DH5α competent cells and spread onto X-gal plates. After 17 h incubation at 37°C, white colonies were picked, and the DNA was amplified by colony PCR. Positive colonies were grown in Luria Bertani (LB) liquid culture, and plasmids extracted using the Plasmid Extraction Kit (Roche), and then sequenced for at least three times. 16S rRNA sequencing data of each isolate was compared to the sequences of reference strains available in the Gen Bank (NCBI) databases using the BLASTn search tool (http://blast.ncbi.nlm.nih.gov) to determine the most similar sequences in the Gen Bank database. The sequences from this study were deposited to Gen Bank under the accession numbers of GU586300 to GU586322. Sequences of strains with the maximumhomology were selected to identify the isolates. A phylogenetic tree was constructed based on the UPGMA of Mega4 software version 4 (www.megasoftware.net).
3.3. Detection of Catabolic Genes by PCR and Hybridization Analysis
Primer sets include alkM (Alkane monooxygenase, F: 5’-CGIGIIGCIACICCTGAAGATCCAGC-3’ and R: 5’-ITTATTITTCCAICTATGCTCTGG-3’, (22), alkB (Alkane hydroxylase, F: 5’-TGGCCGGCTACTCCGATGATCGGAATCTGG-3’ and R: 5’-CGCGTGGTGATCCGAGTGCCGCTGAAGGTG-3’, (15), xylEa (Catechol 2,3-oxygenase, F: 5’-TCGAGTTGCTGGGCCTGATCG-3’ and R: 5’-CCCGCAGAACACTTCGTTGCG-3’, (23), xylE (F: 5’-GCGAGATAGAAGCGCTCTTG-3’ and R: 5’-GTATTGATACCTGGGAGGAAG-3’, (15), C23O (Catechol 2,3-oxygenase, F: 5’-CAAGGCCCACGACGTGGCNTT-3’ and R: 5’-CGGTTACCGGACGGGTCGAAGAAGT-3’, (24), NiadA (Pyrenedioxygenase, F: 5’-ATCTTCGGGCGCGCCTGGGTGTTTCTCGG-3’ and R: 5’-AATTGTCGGCGGCTGTCTTCCAGTTCGC-3’, (22), to dC1 (Toluene dioxygenase, F: 5’-GCGAGATAGAAGCGCTCTTG-3’, R: 5’-GTATTGATACCTGGGAGGAAG-3’ (15) and nahAC (Naphthalene dioxygenase, F: 5’-AAGCACCTGATTCATGGCGATGA-3’ and R: 5’-GAACTCAGCCCAGTTGGAGCTGCTG-3’, (4) were selected to determine the presence of certain catabolic genes that encode enzymes involved in hydrocarbon degradation pathways. DNA was extracted from isolated strains as described above and screened by PCR. The PCR fragments were cloned as mentioned above, and then sequenced. Each sequenced PCR product that showed similarity to the target catabolic gene was used for the synthesis of the probe. Hybridization reaction was performed to detect catabolic genes in other bacterial isolates. DNA of 23 isolates was denatured and spotted onto a Hybond nylon membrane (Roche). The membranes were hybridized with DNA probes specific for the genes alkB, alkM, and xylE using high-stringency prehybridization, hybridization, and washing conditions at 65°C. The probes were labeled with the digoxigenin (DIG) nonradioactive nucleic acid labeling and detection system, using the DIG DNA Labeling and Detection Kit (Roche), according to the manufacturer’s instructions.
3.4. Analysis of Hydrocarbon Degradation in Liquid Culture
Bushnell-Haas liquid culture with 2.5% (w/v) crude oil as carbon source was inoculated by the bacterial consortium at 25°C on a rotary shaker at 150 rpm for 10 days. Control flasks devoid of inoculum, were incubated under the same conditions. Residual TPH in liquid culture was extracted with dichloromethane, dried and purified with silica gel to remove polar compounds such as microbial matter and products (25). The GC-MS analysis of extracted TPH was performed using a Fisons instruments gas chromatograph 8000 (Italy) with a CP-Sil 5CB column (30 m × 0.25 mm, 0.25μm film thickness), connected to a mass detector (TRIO 1000) operating at an ionization energy of 70 ev, and using helium as the carrier gas with a split ratio of 1:20. The oven temperature was programmed at 50°C for 5 min and then increased to 280°C at 5°C per min, and held at this temperature for 20 min. The compounds were identified by comparison of their mass spectra with the mass-spectral libraries.
4.1. Isolation and Characterization of Bacteria
The soil samples taken from the polluted locations were very viscous and dark brown to black. BS9 and RS1 soil samples had the highest (33.1 ± 0.55%) and lowest (4.5 ± 0.29%) TPH concentrations, respectively (Table 1). Generally, TPH concentrations in the plant-free soils were greater than that in the rhizospheric soils. However, the number of different oil-degrading colonies showed a reverse trend, being greater in the rhizosphere soil samples than in the plant-free soils. This may indicate the availability of root exudates of wild oat to the bacteria inhibiting the rhizosphere, and thereby enhancing bacterial composition and diversity (Table 1). Similar results have been previously reported by Liste and Prutz (18). It appears that increasing TPH concentrations, decreased soil pH at polluted locations. In general, soil samples showed relatively high EC values (up to 4.93 dSm-1 in 1:2.5 soil extract, Table 1), probably due to the geographical location and the high water table. Totally, 150 bacterial colonies, capable of growing in presence of 10% (w/v) crude oil as the sole carbon source were isolated from the wild oat rhizosphere using serial enrichment method. The areas where the soil samples were collected usually exposed to an extreme temperature (above 50°C) during midsummer; thus, all isolated colonies were considered as thermo tolerant (data not shown). The twenty three fastest growing bacterial isolates were selected to characterize their 16S rRNA gene sequences, and deposited in GenBank database under accession numbers of GU586300 to GU586322.
Figure 1 shows a maximum likelihood tree constructed based on 16S rRNA gene sequences. The twenty three sequences were affiliated, with 98-100% identity, to twelve different genera including Acinetobacter, Pseudomonas, Enterobacter, Cronobacter, Stenotrophomonas, Achromobacter, Ochrobactrum, Paenibacillus, Bacillus, Microbacterium, Curtobacterium and Sphingobacterium.
Among the twenty three isolates, sixteen sequences were affiliated to Class Proteobacteria, four sequences to Class Firmicutes, two sequences to Class Actinobacteria, and one sequence to the Bacteroidetes class. In the Class Proteobacteria, thirteen sequences were affiliated to the gamma subclass, two sequences to the beta and one to the alphaone. Gamma Proteobacteria dominated rhizosphere-inhabiting oil-degrading populations; Bacteroidetes, Actinobacteria and Firmicutes were found at lower amounts. 65% of the effective isolates contained Gram-negative bacteria.
4.2. Detection of Catabolic Genes
Presence of catabolic genes was investigated with PCR tests and DNA-DNA hybridization. Results are shown in Figure 1. The alkB gene belonged to Pseudomonas sp. (GU586315) showed 89% nucleotide sequence identity with P. oleovorans alkB gene, and alkB-derived amino acid sequences indicated 94% similarity with alkB belonged to P. oleovorans TF4-1L. Amplified fragments of alkM from Acinetobactersp. (GU586302) showed 82% identity withalkM belonged to Acinetobacter sp. strain 69-V; amino acid sequence of corresponding gene was 90% similar to that of alkM. The xylEa nucleotide sequence of Pseudomonas sp. (GU586316) revealed 99% nucleotide sequence and 99% amino acid identities with those of P. putida, while the nucleotide sequence of xylE from Acinetobacter sp. (GU586302) showed 98% nucleotide sequence and amino acid identities with those of P. putida.
Hybridization probes were PCR products of alkB (Pseudomonas sp.), alk M (Acinetobacter sp.) and xylEa (Pseudomonas sp.). Specific hybridization experiments were performed to detect aforementioned catabolic genes in the case of unsuccessful amplifications as well as confirmation of positive isolates. No significant cross-hybridization was observed.
4.3. Microbial Degradation of Petroleum Hydrocarbon in Liquid Culture
GC-MS profiles of residual crude petroleum in inoculated and control media are depicted in Figure 2. Three isolates including Acinetobacter sp. (GU586302), Acinetobacter sp. (GU586303) and Stenotrophomonas sp. (GU586311) were selected as a consortium for the biodegradation assays due totheir fast growth in the presence of hydrocarbons (Table 2), reflecting corresponding catabolic pathways (Figure 1). GC-MS analysis of residual hydrocarbons in treated and untreated cultures showed decreased TPH after 10 days of incubation in the presence of bacterial consortium. N-alkane fractions removed in terms of in carbon number has been shown in Figure 3. The carbon number in alkane chains ranged from C7 to C24. GC-MS profile of oil residual components in liquid culture was compared to the control and showed that all components were highly reduced in inoculated culture; however, the consortium degraded C7-C24 n-alkanes in different quantities (Figure 3). Degraded aromatic compounds (PAHs) included C7 (toluene), C8 (ethylbenzene, m-, p- and o-xylene), C9, C10 (naphthalene), C12 and C18 (chrysene or benzo-anthracene). In cultures inoculated with the bacterial consortium, crude oil amounts generally decreased about 40% after 10 days of incubation (Table 2). With the exception of C9 and C10, GC-MS analyses revealed that removal rate of n-alkane components, i.e. C7-24 was more than 80% in the consortium culture. C10 was the least degraded component in maximum removal rate of approximately 2%. The overall removal rates of oil hydrocarbons (aliphatic, aromatic, naphthenic, olefin) by the AS and ASPP consortia were 40.14% and 43.7% after 10 days, respectively (Table 2).
Iran is the world’s fifth crude oil producer (www.wikipedia.org). Environmental pollution is a severe challenge which oil producing countries face due to crude oil exploitation, refining, transportation, storage as well as accidents. Khuzestan is Iranian oil producing center of excellence where soil has been heavily contaminated over the past 80 years, threatening local ecosystems such as wetlands and rivers. Numerous efforts have been made to isolate indigenous oil-degrading bacteria in oil-contaminated soil and waters in Mideast oil rich countries such as Kuwait and Bahrain (19, 26, 27). In the present research, previously isolated rhizosphere-inhabiting bacterial strains in wild oat plants growing in contaminated sites were screened and introduced through a series of oil-enriched cultures as the sole carbon and energy sources based on their oil degrading efficacies and capabilities. The strains belonged to different genera previously reported as oil degrading species (8, 9, 20, 28, 29). Studies suggested that hydrocarbon contamination enriches oil degrading bacteria (6, 10). Proteobacteria and gamma Proteobacteria are commonly found in microbial communities exposed to hydrocarbon pollution (30, 31). We also found class Proteobacteria and gamma Proteobacteria highly prevalent belonged to r-strategists group, such as Pseudomonas (22). Kaplan and Kitts (32) suggested that Gram-negative bacteria dominate oil-contaminated areas. The rhizosphere is a dynamic environment where the roots, soil and microorganisms interact. Branched root systems provide organic substrate for microbial communities and consequently might accelerate degradation of soil organic matter (SOM) and organic pollutants (33), a process leading to the soil priming effect (25, 34). Beneficial root effects and long-term petroleum contamination can develop a very specific highly potent microbial community capable of detoxifying pollutants. Major isolates belonged to Acinetobacter and Pseudomonas, while others were only found in rhizosphere-bound soil samples including Curtobacterium, Paenibacillus, Sphingobacterium and Microbacteriumsp., Stenotrophomonas rhizophila. Haichar et al. (33) reported that Sphingomonadales was specifically associated with monocotyledons such as wheat and maize, whereas Enterobacter-like bacteria were considered as generalists colonizing both mono and dicotyledons. We found that Sphingobacterium was associated with the wild oat roots (A. fatua) and could use petroleum hydrocarbon. Microbacterium sp. was found by Supaphol et al. (35) in tropical soil samples contaminated with petroleum, we recorded the species in the crude oil-contaminated wild oat rhizosphere in Iran for the first time. In Iran, most bioremediation works have focused on degradation rate of contaminant in oil-polluted soil and sediments, and we are the first to report molecular mechanisms probably involved in hydrocarbon degradation. Identifying catabolic genes in hydrocarbon-degrading bacteria is a common approach to evaluate hydrocarbon-contaminated sites for bioremediation processes (36). The specific probes of alkB, alkM, xylE genes, catabolic genes involved in hydrocarbon degradation, were used to determine the potential of bacterial isolates. Dot-blot hybridization experiments revealed the presence of these catabolic genes in several bacterial strains. Frequent occurrence of specific hybridization between sequenced DNA belonged to the isolates and their corresponding probes reflected high sequence identity between some isolates and corresponding regions of the target catabolic genes. The xylE gene involved in the aromatic hydrocarbon degradation pathway (xylE, BTEX degradation), was found in 10 of the 23 bacterial isolates (13). In addition to the Pseudomonas strains, the xylE gene was detected in other Proteobacteria isolates, such as Acinetobacter, Achromobacter, Enterobacter and Stenotrophomonas. In fact, hybridization and PCR amplification confirmed the presence of xylE in half of the isolates. Aromatic hydrocarbons are more resistant against biodegradation than aliphatic compounds, they often cause serious problems during bioremediation (3, 8). TOL catabolic plasmid carries the gene (13); thus, these bacteria could obtain the gene through horizontal transfer rather than independent evolution of their degradation capabilities (6, 18, 23). Root exudates (e.g. phenolics) selectively enriched bacterial flora, regulates the expression of PAHs catabolic genes in certain plant species (16). For instance, dioxygenase-expressing bacteria were the most abundant ones in contaminated rhizosphere in mustard, oat, and cress attributed to phenolic root exudates (16). Old persistent crude oil contamination as a most important environmental factor has driven bacterial population selection, followed by horizontal gene transfer between microbial communities diverged them to survive. Partial sequences analysis of 16S rRNA gene revealed a level of diversity mainly associated with two bacterial divisions; Proteobacteria and Firmicutes (Figure 1). Certain hydrocarbon-degrading strains, i.e. Stenotrophomonas, Achromobacter, Pseudomonas, Enterobacter and Acinetobacter possess both aromatic and aliphatic catabolic pathways suggesting that a single strain is capable of degrading both aromatic and aliphatic hydrocarbons. Two degradation pathways were previously reported in a single microorganism (4, 8). Some isolates carried none of the studied genes; however, they grew in the presence of crude oil implying the existence of other biodegradation pathways in an indigenous population. Certain bacterial species occasionally provide degrading pathways when acting synergistically (37); diverse root exudates promote the assembly of these synergistic communities (16). Although all isolates grew in mineral media with crude petroleum as the carbon source, but 5 of the 23 isolates showed the most significant growth rates. These 5 isolates were identified as different species of Stenotrophomonas, Pseudomonas and Acinetobacter (Table 2). In fact, these strains grew in media with high quantities (up to 10%) of crude petroleum effectively, confirming that these strains are capable of using n-alkanes. GC-MS experiment revealed hydrocarbons with carbon numbers as high as C24 suggesting that these strains possibly use longer chain n-alkanes. Acinetobacter sp. strain M-1 has reportedly degraded a variety of n-alkanes, including very long chain n-alkanes (or paraffin wax) with carbon chain lengths up to C44 that are under a solid state (38). A combination of Acinetobacter sp. and P. putida has been found to degrade Arabian light crude oil containing40% saturates and 21% aromatics (8). In the present study, crude petroleum at 25 g/L (2.5%) was degraded up to 40% during 10 days in the presence of bacterial consortia. Stenotrophomonas spp. and Stenotrophomonas maltophilia have been reported to use alkanes (39) and polycyclic aromatic hydrocarbons (29). The results also confirmed the presence of alkB and xylE genes in Stenotrophomonas species bearing ecological role in mineral cycle in nature, promoting plant growth via biological control of fungal diseases, and degrading a wide range of pollutants. Thus, it might be used potentially for bioremediation and phytoremediation (40).Using aromatic compounds by Enterobacteria has also been reported previously (41). Pseudomonas and Enterobacter strains, PAH-degrading microorganisms, have been isolated in highly oil-polluted soil, along with an Enterobacter strain capable of degrading naphthalene (42). Genetic study of PAH degradation demonstrated the presence of a highly efficient pathway for the degradation of naphthalene and phenanthrene, in Pseudomonas (29). Also, it has been reported that these species act as plant growth promoting rhizobacteria enhancing its tolerance to contaminants (43). During the incubation period, emulsification of crude petroleum was observed in the culture broth inoculated with consortia, suggesting that the production of extracellular bio surfactants may be one of the underlying mechanisms implemented by the isolates for using crude oil. Actually, production of bio surfactants is one of the ways that microorganisms take up hydrophobic substrates (2, 9). It has been frequently reported that the Acinetobacter spp. Produce biosurfactants/bioemulsifiers indicating the presence of a hydrophobic exterior enabling cellular contact with hydrocarbons (2). Production of biosurfactants has also been reported in other isolates, such as Pseudomonas aeruginosa, Acinetobacter spp., Acinetobacter radioresistens, Acinetobacter calcoaceticus and Bacillus polymyxa (8, 44). Presence of alkanes in control cultures revealed by GC-MS suggested that n-alkanes (C7-24) were removed up to 80% in consortium culture except for C9 and C10. The least degraded component was C10, which showed a maximum removal of 2% approximately. The overall removal of petroleum hydrocarbons (aliphatic, aromatic, naphthenic, olefin) by the AS and ASPP consortia was 40.14% and 43.7% after 10 days, respectively (Table 2). This study showed that longer chain petroleum hydrocarbons were degraded more effectively. Our results also revealed that C8 to C19-hydrocarbon chains dominated the crude petroleum samples, indicating that it is important to develop a more efficient consortium procedure to degrade shorter chain hydrocarbons. Studied strains used both long and shorter chain alkanes (Figure 2). As a substrate for bacterial enrichment, using crude oil has often led to isolation of microorganisms metabolizing n-alkanes (6). GC-MS analyses demonstrated that n-alkanes (C12-C24) were preferentially degraded by the consortium, when compared to PAHs present in the crude petroleum. Jain et al., (9) reported that most of n-alkanes can be significantly biodegraded within the first 10 days of incubation. The present work showed that the consumption of the substrates is faster and more efficient by mixed cultures in comparison to pure ones (Table 2). Thus, to achieve efficient crude petroleum biodegradation, a rather large consortium of the effective species would be needed. However, the higher growth rates of Stenotrophomonas, Pseudomonas and Acinetobacter, might be related to higher breakdown and using petroleum hydrocarbons compared to other isolated strains. We isolated a wild oat rhizosphere-inhabiting oil-degrading bacterial spectrum. Some of the studied species have been named as plant growth promoting rhizobacteria which might help plants to grow and withstand in severely polluted soil. Therefore, our findings highlighted the role of such isolates to be considered in further studies. The results of microbial isolation and identification corroborated literature data about the presence of aerobic bacteria in petroleum-contaminated rhizosphere in wild oat support the hypothesis that these bacteria might play a role in oil biodegradation processes in situ. Phytoremediation with such a spectrum of oil degraders would be the preferable choice of treatment at a demonstrative scale. Crude petroleum is a complex mixture of hydrophobic components, assemblies of mixed populations with overall broad enzymatic capacities are required for increasing the rate and extent of TPH biodegradation. Bio and phytoremediation techniques success relies on inoculating the right environment with the right microbial consortia. Therefore, an in-depth understanding of processes that occur, the responsible microorganisms and molecular mechanisms involved in the degradation process would provide more chance to tailor techniques to site-specific remediation. Long-term exposure to widespread crude oil pollution in southwestern of Iran facilitates the selection and co-evolution of bacteria and plants. The findings of this study affirmed the adaptability of microbes of the rhizosphere and their great potential to be exploited for cleaning up hydrocarbon contaminated sites, either through inoculation of specifically isolated microbial communities or targeted stimulation of the selected species in situ in the presence of wild oat. Isolation, identification of bacteria and detection of some catabolic pathways from wild oat rhizobacteria is the first study of this kind in Iran and thus might be considered as an important step towards the development of phytoremediation strategies for sites contaminated with these pollutants.
This work was supported by the Grant Number 285 from the National Institute of Genetic Engineering and Biotechnology (NIGEB).
The work presented here was carried out in collaboration between all authors. SMS and SR defined the research theme. SR and SMS designed methods and experiments, carried out the laboratory experiments, analyzed the data, interpreted the results and wrote the paper. JR, BR and FR co-designed experiments, discussed analyses, interpretation, and presentation. All authors have contributed to, seen and approved the manuscript.
1. EPA U. Method 3540C non-volatile and semi-volatile organic compounds: EPA; 1986.
2. Stroud JL, Paton GI, Semple KT. Microbe-aliphatic hydrocarbon interactions in soil: implications for biodegradation and bioremediation. J Appl Microbiol. 2007;102(5):1239-53.
3. Mishra S, Jyot J, Kuhad RC, Lal B. Evaluation of inoculum addition to stimulate in situ bioremediation of oily-sludge-contaminated soil. Appl Environ Microbiol. 2001;67(4):1675-81.
4. Churchill SA, Harper JP, Churchill PF. Isolation and characterization of a Mycobacterium species capable of degrading three- and four-ring aromatic and aliphatic hydrocarbons. Appl Environ Microbiol. 1999;65(2):549-52.
5. Margesin R, Schinner F. Bioremediation (natural attenuation and biostimulation) of diesel-oil-contaminated soil in an alpine glacier skiing area. Appl Environ Microbiol. 2001;67(7):3127-33.
6. Kuiper I, Lagendijk EL, Bloemberg GV, Lugtenberg BJ. Rhizoremediation: a beneficial plant-microbe interaction. Mol Plant Microbe Interact. 2004;17(1):6-15.
7. Atlas RM. Microbial degradation of petroleum hydrocarbons: an environmental perspective. Microbiol Rev. 1981;45(1):180-209.
8. Van Hamme JD, Singh A, Ward OP. Recent advances in petroleum microbiology. Microbiol Mol Biol Rev. 2003;67(4):503-49.
9. Jain PK, Gupta VK, Pathak H, Lowry M, Jaroli DP. Characterization of 2T engine oil degrading indigenous bacteria, isolated from high altitude (Mussoorie), India. World J Microbiol Biotechnol. 2010;26(8):1419-26.
10. Whyte LG, Schultz A, Beilen JB, Luz AP, Pellizari V, Labbe D, et al. Prevalence of alkane monooxygenase genes in Arctic and Antarctic hydrocarbon-contaminated and pristine soils. FEMS Microbiol Ecol. 2002;41(2):141-50.
11. Throne-Holst M, Markussen S, Winnberg A, Ellingsen TE, Kotlar HK, Zotchev SB. Utilization of n-alkanes by a newly isolated strain of Acinetobacter venetianus: the role of two AlkB-type alkane hydroxylases. Appl Microbiol Biotechnol. 2006;72(2):353-60.
12. Van Beilen JB, Panke S, Lucchini S, Franchini AG, Rothlisberger M, Witholt B. Analysis of the Pseudomonas putida alkanedegradation gene cluster and flanking insertion sequences: evolution and regulation of the alk genes. Microbiology. 2001;147:1621-30.
13. Jussila MM, Zhao J, Suominen L, Lindstrom K. TOL plasmid transfer during bacterial conjugation in vitro and rhizoremediation of oil compounds in vivo. Environ Pollut. 2007;146(2):510-24.
14. Piskonen R, Nyyssonen M, Itavaara M. Evaluating the biodegradation of aromatic hydrocarbons by monitoring of several functional genes. Biodegradation. 2008;19(6):883-95.
15. Ringelberg DB, Talley JW, Perkins EJ, Tucker SG, Luthy RG, Bouwer EJ, et al. Succession of phenotypic, genotypic, and metabolic community characteristics during in vitro bioslurry treatment of polycyclic aromatic hydrocarbon-contaminated sediments. Appl Environ Microbiol. 2001;67(4):1542-50.
16. Daane LL, Harjono I, Zylstra GJ, Haggblom MM. Isolation and characterization of polycyclic aromatic hydrocarbon-degrading bacteria associated with the rhizosphere of salt marsh plants. Appl Environ Microbiol. 2001;67(6):2683-91.
17. Siciliano SD, Germida JJ. Mechanisms of phytoremediation: biochemical and ecological interactions between plants and bacteria. Envir Rev. 1998;6(1):65-79.
18. Liste HH, Prutz I. Plant performance, dioxygenase-expressing rhizosphere bacteria, and biodegradation of weathered hydrocarbons in contaminated soil. Chemosphere. 2006;62(9):1411-20.
19. Yateem A, Al-Sharrah T, Bin-Haji A. Investigation of microbes in the rhizosphere of selected trees for the rhizoremediation of hydrocarbon-contaminated soils. Int J Phytoremediation. 2008;10:311-24.
20. Arvanitis N, Katsifas EA, Chalkou KI, Meintanis C, Karagouni AD. A refinery sludge deposition site: presence of nahH and alkJ genes and crude oil biodegradation ability of bacterial isolates. Biotechnol Lett. 2008;30(12):2105-10.
21. Wilson K. Preparation of genomic DNA from bacteria. In: Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Struhl K, editors. Current Protocols in Molecular Biology. New York: John Wiley & Sons; 1994. p. 241-5.
22. Margesin R, Labbe D, Schinner F, Greer CW, Whyte LG. Characterization of hydrocarbon-degrading microbial populations in contaminated and pristine Alpine soils. Appl Environ Microbiol. 2003;69(6):3085-92.
23. Jussila MM, Jurgens G, Lindstrom K, Suominen L. Genetic diversity of culturable bacteria in oil-contaminated rhizosphere of Galega orientalis. Environ Pollut. 2006;139(2):244-57.
24. Junca H, Pieper DH. Amplified functional DNA restriction analysis to determine catechol 2,3-dioxygenase gene diversity in soil bacteria. J Microbiol Methods. 2003;55(3):697-708.
25. Kuzyakova Y, Friedelb JK, Stahr K. Review of mechanisms and contaminated soil. Chemosphere. 2000;62:1485-95.
26. Obuekwe CO, Al-Jadi ZK, Al-Saleh ES. Hydrocarbon degradation in relation to cell-surface hydrophobicity among bacterial hydrocarbon degraders from petroleum-contaminated Kuwait desert environment. Int Biodeterior Biodegrad. 2009;63(3):273-79.
27. Dashti N, Khanafer M, El-Nemr I, Sorkhoh N, Ali N, Radwan S. The potential of oil-utilizing bacterial consortia associated with legume root nodules for cleaning oily soils. Chemosphere. 2009;74(10):1354-9.
28. Mohamed ME, Al-Dousary M, Hamzah RY, Fuchs G. Isolation and characterization of indigenous thermophilic bacteria active in natural attenuation of bio-hazardous petrochemical pollutants. Int Biodeterior Biodegrad. 2006;58(3–4):213-23.
29. Molina MC, Gonzalez N, Bautista LF, Sanz R, Simarro R, Sanchez I, et al. Isolation and genetic identification of PAH degrading bacteria from a microbial consortium. Biodegradation. 2009;20(6):789-800.
30. McKew BA, Coulon F, Yakimov MM, Denaro R, Genovese M, Smith CJ, et al. Efficacy of intervention strategies for bioremediation of crude oil in marine systems and effects on indigenous hydrocarbonoclastic bacteria. Environ Microbiol. 2007;9(6):1562-71.
31. Frenzel M, James P, Burton S, Rowland SJ, Lappin-Scott HM. Towards bioremediation of toxic unresolved complex mixtures of hydrocarbons: identification of bacteria capable of rapid degradation of alkyltetralins. J Soils Sediments. 2009;9(2):129-36.
32. Kaplan CW, Kitts CL. Bacterial succession in a petroleum land treatment unit. Appl Environ Microbiol. 2004;70(3):1777-86.
33. Haichar FZ, Marol C, Berge O, Rangel-Castro JI, Prosser JI, Balesdent J, et al. Plant host habitat and root exudates shape soil bacterial community structure. ISME J. 2008;2(12):1221-30.
34. Blagodatskaya Е, Kuzyakov Y. Mechanisms of real and apparent priming effects and their dependence on soil microbial biomass and community structure: critical review. Biol Fertil Soils. 2008;45(2):115-31.
35. Supaphol S, Panichsakpatana S, Trakulnaleamsai S, Tungkananuruk N, Roughjanajirapa P, O’Donnell AG. The selection of mixed microbial inocula in environmental biotechnology: example using petroleum contaminated tropical soils. J Microbiol Methods. 2006;65(3):432-41.
36. Márquez-Rocha FJ, Olmos-Soto J, Concepción Rosano-Hernández M, Muriel-García M. Determination of the hydrocarbon-degrading metabolic capabilities of tropical bacterial isolates. Int Biodeterior Biodegrad. 2005;55(1):17-23.
37. Lappin HM, Greaves MP, Slater JH. Degradation of the herbicide mecoprop [2-(2-methyl-4-chlorophenoxy)propionic Acid] by a synergistic microbial community. Appl Environ Microbiol. 1985;49(2):429-33.
38. Tani A, Ishige T, Sakai Y, Kato N. Gene structures and regulation of the alkane hydroxylase complex in Acinetobacter sp. strain M-1. J Bacteriol. 2001;183(5):1819-23.
39. Tesar M, Reichenauer TG, Sessitsch A. Bacterial rhizosphere populations of black poplar and herbal plants to be used for phytoremediation of diesel fuel. Soil Biol Biochem. 2002;34(12):1883-92.
40. Ryan RP, Monchy S, Cardinale M, Taghavi S, Crossman L, Avison MB, et al. The versatility and adaptation of bacteria from the genus Stenotrophomonas. Nat Rev Microbiol. 2009;7(7):514-25.
41. Gupta A, Singh R, Khare SK, Gupta MN. A solvent tolerant isolate of Enterobacter aerogenes. Bioresour Technol. 2006;97(1):99-103.
42. Toledo FL, Calvo C, Rodelas B, Gonzalez-Lopez J. Selection and identification of bacteria isolated from waste crude oil with polycyclic aromatic hydrocarbons removal capacities. Syst Appl Microbiol. 2006;29(3):244-52.
43. Zhuang X, Chen J, Shim H, Bai Z. New advances in plant growth-promoting rhizobacteria for bioremediation. Environ Int. 2007;33(3):406-13.
44. Das K, Mukherjee AK. Characterization of biochemical properties and biological activities of biosurfactants produced by Pseudomonas aeruginosa mucoid and non-mucoid strains isolated from hydrocarbon-contaminated soil samples. Appl Microbiol Biotechnol. 2005;69(2):192-9.