- Email: [email protected]
Enrichment and isolation of crude oil degrading bacteria from some mussels collected from the Persian Gulf
Marine Pollution Bulletin 101 (2015) 85–91
Contents lists available at ScienceDirect
Marine Pollution Bulletin journal homepage: www.elsevier.com/locate/marpolbul
Enrichment and isolation of crude oil degrading bacteria from some mussels collected from the Persian Gulf Zeynab Bayat, Mehdi Hassanshahian ⁎, Majid Askari Hesni Department of Biology, Faculty of Sciences, Shahid Bahonar University of Kerman, Kerman, Iran
a r t i c l e
i n f o
Article history: Received 12 June 2015 Received in revised form 5 November 2015 Accepted 6 November 2015 Available online 12 November 2015 Keywords: Crude oil Marine environment Mussel Persian Gulf Pollution
a b s t r a c t To date, little is known about existing relationships between mussels and bacteria in hydrocarbon-contaminated marine environments. The aim of this study is to find crude oil degrading bacteria in some mussels at the Persian Gulf. Twenty eight crude oil degrading bacteria were isolated from three mussels species collected from oil contaminated area at Persian Gulf. According to high growth and degradation of crude oil four strains were selected between 28 isolated strains for more study. Determination the nucleotide sequence of the gene encoding for 16S rRNA show that these isolated strains belong to: Shewanella algae isolate BHA1, Micrococcus luteus isolate BHA7, Pseudoalteromonas sp. isolate BHA8 and Shewanella haliotis isolate BHA35. The residual crude oil in culture medium was analysis by Gas Chromatography (GC). The results confirmed that these strains can degrade: 47.24%, 66.08%, 27.13% and 69.17% of crude oil respectively. These strains had high emulsification activity and biosurfactant production. Also, the effects of some factors on crude oil degradation by isolated strains were studied. The results show that the optimum concentration of crude oil was 2.5% and the best degradation take place at 12% of salinity. This research is the first reports on characterization of crude oil degrading bacteria from mussels at Persian Gulf and by using of these bacteria in the field the effect of oil pollution can be reduce on this marine environment. © 2015 Elsevier Ltd. All rights reserved.
1. Introduction Many techniques are utilized to mitigate or cleanse petroleum pollution in the environment. During exploration, production, refining, transport and storage of petroleum and petroleum products, some accidental spill may be released into the sea waters (Dave and Ghaly, 2011; Ghanavati et al., 2008). Crude oil is toxic, mutagenic and carcinogenic this pollutant concedes a serious damage to marine life (Todd et al., 2010). Bioremediation is the best method for elimination of oil spills. Bioremediation of oil contamination compared with physicochemical treatment is more effective and lower cost price (Saxena et al., 2013). Bacteria are important for the biodegradation of petroleum hydrocarbons and many hydrocarbon-degrading bacteria have been isolated from different environments (Kaczorek et al., 2008; Udeh et al., 2013; Hassanshahian et al., 2012a; Radwan et al., 2002; Khan et al., 2006). Biodegradation rates can be controlled by concentration and composition of hydrocarbons, nutrients, oxygen, salt concentrations, moisture and temperature (Sathishkumar et al., 2008; Hassanshahian et al., 2010). Also, biosurfactants increase the solubility of hydrocarbons to bacteria and enhanced degradation of these toxic compounds (Hassanshahian et al., 2012a). Marine organisms can take up contaminants from bottom
⁎ Corresponding author. E-mail address: [email protected] (M. Hassanshahian).
http://dx.doi.org/10.1016/j.marpolbul.2015.11.021 0025-326X/© 2015 Elsevier Ltd. All rights reserved.
sediments, suspended particulate material and food sources. Filterfeeding bivalves such as mussels, oysters, and clams are very important components in the process of bioremediation of the marine environment (Stabili et al., 2005). They are capable to filter and accumulate a large number of fine particulate matters, including phytoplankton, zooplankton, microorganism and other particulate organic debris. Furthermore, the organic components in the suspended matters can be trapped and applied by filter-feeding bivalves (Manganaro et al., 2009; Zhou et al., 2014; Hassanshahian et al., 2013). Little work has been done on the distribution and physiology of autochthonous hydrocarbondegrading microbes inhabiting marine organisms (Cappello et al., 2012a; Hassanshahian et al., 2012b). Some researchers confirmed that crude oil degrading bacteria were existing in marine organisms. For example, Radwan et al. (2005) establish that some crude oil degrading bacteria such as Pseudomonas, Bacillus and Acinetobacter were associated with cyanobacterial mat at Arabian Gulf (Radwan et al., 2005; Radwan et al., 2002). Also, Al-Mailem et al. (2010) study crude oil degrading bacteria that symbiosis with filamentous cyanobacteria and their results show that some bacteria such as Halomonas aquamarina, Marinobacter hydrocarbonoclasticus, Marinobacter sp.; Dietzia maris and Alcanivorax can degrade crude oil and present in cyanobacteria (Al-Mailem et al., 2010; Sorkhoh et al., 1990; Al-Awadhi et al., 2003). In all research that studied the relationship between bacteria and marine animals, it is confirmed that the density of bacteria in the marine animal samples were higher compared to surrounding water. For
86
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
Fig. 1. Enumeration of heterotrophic and hydrocarbon-degrading bacteria by CFU and MPN methods. All data present in this figure are the average of three replicates of enumeration in the same dilution.
example, mussels can filter the seawater and concentrate the marine bacteria that cause reducing the bacterial concentration in the seawater (Cavallo et al., 2009). To date, little is known about existing relationships between mussels and bacteria in hydrocarbon-contaminated marine environments. The aim of this study is to find crude oil degrading bacteria in some mussels at the Persian Gulf. Also identification of these strains and the degradation capacity of these isolates is another purpose of this research.
related to: Crassostrea gigas, Chama asperella and Barbatia tenella genus. Collected samples were transported on ice to the laboratory. Mussels shell was removed by sterile knife and gill and other tissue of mussel samples was washed with sterile seawater. In fact, the tissue was homogenized in buffer. Finally, macerate of mussels was used for subsequent studies.
2. Materials and methods
Crude oil degrading bacteria were isolated in ONR7a medium supplemented with 1% (v/v) of crude oil (Iranian light crude oil) as sole carbon source and energy. ONR7a contained (per liter of distilled water) 40 g of NaCl, 11.18 g of MgCl2·6H2O, 3.98 g of Na2SO4, 1.46 g of CaC12·2H2O, 1.3 g of TAPS0 {3-[N tris(hydroxymethyl) methylamino]-2 hydroxypropanesulfonic acid}, 0.72 g of KCl, 0.27 g of NH4Cl, 89 mg of Na2HP04·7H20, 83 mg of NaBr, 31 mg of NaHCO3, 27 mg of H3BO3, 24 mg of SrCl2·6H20, 2.6 mg of NaF and 2 mg of FeCl2·4H20. For solid media, Bacterial Agar (15 g/l) was added to the solution (Hasanshahian and Emtiazi, 2008). Tissue macerate of mussel (5 ml), condensed seawater (5 ml) and portion of sediments (10 g) were added to Erlenmeyer flasks containing
2.1. Sampling Mussels, seawater and sediment samples were collected from two oil contaminated sites at Persian Gulf. These two stations were located in the Qeshm island (AZ: Zakeri Harbor; 36°15, N; 34°15, E) and (PZ: Park Ziton 37°30, N; 49°15, E). Mussels were collected from depth range 5–15 m. Also, samples of seawater from above the mussels' bed in a sterile bottle, and samples of sediment from below were collected. The mussels were identified according to the standard key (Huber, 2010). The results of identification confirmed that these three mussels
2.2. Isolation and selection of crude-oil degrading bacteria
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
87
Fig. 2. Phylogenetic tree of 16S rDNA sequences of the BHA1, BHA7, BHA8 and BHA35 isolates obtained from Persian Gulf. The tree was constructed using sequences of comparable region of the 16S rDNA gene sequences available in public databases. Neighbor-joining analysis using 1000 bootstrap replicates was used to infer tree topology. The bar represents 0.02% sequence divergence.
100 ml of medium and the flasks were incubated for 7 days at 30 °C on rotary shaker (180 rpm, INFORS AG). Then 5 ml were transported to fresh medium. After a series of four further subcultures, inoculums from the flask were streaked out, and phenotypically different colonies
on ONR7a agar were purified. Phenotypically different colonies obtained from the plates were transferred to fresh medium with and without crude oil to eliminate autotrophic and agar utilizing bacteria. The procedure was repeated, and only isolates exhibiting pronounced growth on
Table 1 Growth rate, crude oil removal, emulsification activity and BATH assay for isolated strains. Isolate
BHA1 BHA7 BHA8 BHA35
Growth rate (OD600 nm)
Percentage of oil removal (Spectrometry)
Percentage of oil removal (GC method)
Emulsification activity (E24%)
Cell surface hydrophobicity (BATH %)
Average
SD⁎
Average
SD
Average
SD
Average
SD
Average
SD
0.76 0.81 0.91 0.84
0.124 0.163 0.137 0.081
58.4 55.87 19.74 40
1.71 1.22 1.12 1.61
47.24 66.08 27.13 69.17
1.65 0.85 1.31 0.81
79 25 71 77
0.72 0.42 1.1 0.72
12.9 0 0 36.3
0.56 0 0 1.4
⁎ Standard Deviation (SD).
88
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
Table 2 The effect of different concentrations of NaCl on growth of selected bacterial strains. NaCl concentration (%)
0
20
40
60
80
100
120
140
160
180
200
++++ ++++ − +++
++++ ++++ ++++ ++++
++++ ++++ ++++ ++++
+++ +++ +++ +++
+++ +++ +++ +++
++ ++ ++ ++
+ ++ + ++
− + − −
− + − −
− + − −
− + − −
strain Shewanella algae BHA1 Micrococcus luteus BHA7 Pseudoalteromonas sp. BHA8 Shewanella haliotis BHA35
crude oil were stored in stock media with glycerol at −20 °C for further characterization (Hasanshahian and Emtiazi, 2008).
motility, starch hydrolysis, indole, H2S production, catalase and oxidase, oxidation/fermentation, reduction of nitrate, growth and acidification of carbohydrates tests were performed (Holt et al., 1998).
2.3. Enumeration of bacteria 2.3.1. Quantification the number of cultivable heterotrophic and hydrocarbon degrading bacteria Measurements of bacterial abundance within the mussel and surrounding environment were performed by colony-forming units (CFU) and Most Probable Number (MPN) procedures (Cappello et al., 2012b; Fuchsluger et al., 2010). Heterotrophic bacteria in the mussel, seawater and sediment were estimated by spreading 100 μl of 10-fold diluents flask on plates of Marine Agar medium (MA) and incubating at 30 °C for 3 days. Also, crude oil degrading bacteria in the mussels, seawater and sediment were estimated by spreading 100 μl of 10-fold diluents flask on plates of ONR7a agar medium with crude oil and incubating at 30 °C for 7 days. The bacteria quantification results were expressed as CFU·g− 1 (Cappello et al., 2012b; Hassanshahian et al., 2014a). 2.3.2. Most probable number (MPN) of heterotrophic and hydrocarbon degrading bacteria The MPN method for hydrocarbon-degrading bacteria was done in sterile 24-well microplates using sample aliquots with corresponding dilutions and cultivation media. 1700 μl ONR7a medium was provided in each well of the microplates then dilution series of samples (usually tenfold dilution until 10−1–10−3) were prepared in ONR7a medium. Wells were inoculated with 100 μl of sample diluents. Following sample inoculation, 100 μl of sterile Iranian light crude oil was applied at the centre of each well. Plates were incubated at 30 °C for 21 days. After incubation, visual evaluation of microbial growth was done. On basis, the number of positive test tubes and the counts of microorganisms were statistically evaluated (Fuchsluger et al., 2010). The MPN method for heterotrophic bacteria was done in sterile 24-well microplates. Each well of the microplates was provided with 1700 μl Marine Broth (MB) and 100 μl dilution 10−3–10−4 of sample, and then plates were incubated at 30 °C for 14 days. All samples from each station were used for MPN count. MPN counts were analyzed with the computer program MPN calculator version 4.2 (Fuchsluger et al., 2010; Hassanshahian, 2014d).
2.4.2. Molecular identification Analysis of 16S rRNA was performed to taxonomically characterize the isolated strains. Total DNA of bacterial strains was extracted with the CTAB method. PCR amplification of 16S rRNA genes was performed using the general bacteria primer 27F (5-AGAGTTTGATCCTGGCTCAG-3) and universal reverse primer 1492R (5-TACGYTACCTTGTTACGACTT-3) (Cappello et al., 2012b). The amplification reaction was carried out in a total volume of 25 μl consisting, 2 mM MgCl2 (1 μl), 10× PCR reaction buffer (200 mM Tris; 500 mM KCl) (2.5 μl), 2 mM each dNTP (2 μl), 0.15 mM each primer (1 μl), 1 U (0.5 μl) Taq DNA polymerase (Qiagen, Hilden, Germany) and 2 μl of template DNA (50 pmol). Total volume was brought up to 15 μl using sterile milliQ water. PCR program consisted of 35 cycles was performed in a thermal cycler GeneAmp 5700 (PE Applied Biosystem, Foster City, CA, USA). The temperature profile for PCR was kept, 94 °C for 5 min, 94 °C for 1 min, 54 °C for 1 min, 72 °C for 1 min, 30 cycles; then 72 °C for 10 min and finally storage at 4 °C. The 16S rRNA amplified was sequenced with Big Dye terminator V3.1 cycle sequencing kit on an automated capillary sequencer (model 3100 Avant Genetic Analyzer, Applied Biosystems). Similarity rank from the Ribosomal Database Project RDP) and FASTA Nucleotide Database Query were used to determine partial 16S rRNA sequences to estimate the degree of similarity to other 16S rRNA gene sequences. Analysis and phylogenetic affiliates of sequences was performed as following the protocol described by Yakimov et al. (2007). Phylogenetic tree was drawn by MEGA5 software with neighbor joining method. 2.5. Measure of bacterial growth and crude oil degradation Bacterial isolates were grown at 30 °C for 15 days on rotator shaker (180 rpm). The growth of the isolates was routinely assessed indirectly by measuring the turbidity (OD600 nm) using UV–Visible spectrophotometer (Shimadzu UV-160, Japan). The crude oil removal assay was carried out by dissolving the residual crude oil in the medium in dichloromethane (DCM) and reading the optical density of the oil extract against blank (distill water) at wavelength of 420 nm (Rahman et al., 2004).
2.4. Identification of the isolates 2.6. Gas chromatography (GC) of residual crude oil 2.4.1. Biochemical characterization The following characteristics were determined according to the “Bergey's Manual of Determinative Bacteriology: the Gram stain, Table 3 The effect of individual and mixed bacterial consortia on growth and crude oil degradation by selected bacterial strains. BHA1
Growth rate (OD600 nm) Oil degradation (%) ⁎ Standard Deviation.
BHA35
BHA1 + BHA35
Average
SD⁎
Average
SD
Average
SD
0.90 67.31
0.04 0.816
1.17 73.45
0.138 1.63
1.78 88.69
0.204 1.86
Crude oil degradation was accuracy estimated by GC-FID. The residual crude oil was extracted in each samples. The extraction protocol was as follow: the same volume of DCM was added to each flask and residual crude oil was extracted then, treated with anhydrous sodium sulfate (Na2SO4) to remove residual water. Extracts were concentrated by separating funnel (Hassanshahian et al., 2014c). Analyses were done by GC-FID (Varian 3800 model, USA) equipped with a SE-54 capillary column (25 m × 0.32 mm × 0.1 μm) and flame ionization detector (FID). Helium was used as the carrier gas (30 ml/min). The oven was programmed as follows: 100 °C (1 min) then increased to 300 °C (2 min) at a rate of 30 °C min−1. The samples were quantified according to previously described protocols (Hassanshahian et al., 2014).
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
89
Fig. 3. The effect of crude oil concentrations on removal of crude oil by isolated strains (A), the effect of incubation time on crude oil degradation by isolated strains (B).
2.7. Measure of emulsification activity and bacterial adherence to hydrocarbons (BATH) The emulsification activity (E24) was determined by the addition of hexadecane to the same volume of cell free culture broth. After mixing with vortex for 2 min and leaving to stand for 24 h, the E24 index is given as percentage of height of emulsified layer (in millimeters) divided by total height of the liquid column (in millimeters). Measurement of the bacterial adhesion to hydrocarbon was performed as described by Pruthi and Cameotra (1997).
inoculated with bacterial culture and incubated at 30 °C and 180 rpm for 15 days (Kumar et al., 2007). To study the effect of incubation time on the growth and degradation of crude oil by the selected bacterial strains samples were incubated for 15 and 30 days. The individual and mixed bacterial consortia from overnight culture were transferred to ONR7a medium with 1% (v/v) of crude oil at 30 °C and 180 rpm. Bacterial growth and crude oil degradation measurement were carried out respectively at 600 nm and 420 nm. The experiments were conducted in triplicates and the average values were considered (Sathishkumar et al., 2008). 3. Results
2.8. Effect of different factors on crude oil degradation by isolated strains Different factors were examined for optimization of crude oil degradation. These factors were concentrations of crude oil, NaCl concentration, incubation time and mixed bacterial consortia. The effect of different concentrations of crude oil (1%, 2.5%, 4% and 5.5%) on the growth and degradation of crude oil by the selected bacterial strains was studied using ONR7a medium at 30 °C and 180 rpm for 15 days (Khan et al., 2006). The influence of salinity tolerance on the growth and degradation of crude oil by the selected bacterial strains was carried out using ONR7a medium with 1% (v/v) of crude oil and 0–20% (w/v) concentration of NaCl. The flasks were
3.1. Quantity of heterotrophic and crude oil degrading bacteria in collected samples (CFU and MPN) The quantity of heterotrophic and crude oil degrading bacteria was determined in all collected samples by two enumeration methods: CFU and MPN. The results were shown in Fig. 1. As shown in this figure, the numbers of heterotrophic bacteria in AZ 1 and PZ 1 were the highest (respectively 1.64 × 106, 1.08 × 106 CFU·ml− 1) compared to other samples. AZ 1 and PZ 1 samples have the highest abundance of crude oil degrading bacteria (respectively 5 × 103, 2 × 103 CFU·ml−1). Also, the MPN value for samples AZ 1 and PZ 1 (respectively 6.9 × 103,
90
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
8 × 103) has the highest number of crude oil degrading bacteria compared to other samples and the highest quantity (MPN) of heterotrophic bacteria related to AZ 1 and AZ 3 samples (respectively 4.6 × 106, 3.7 × 106). Also, Fig. 1 illustrate that the MPN values in all samples were higher than CFU values. 3.2. Isolation and identification of bacteria Twenty eight crude oil degrading bacteria were isolated after enrichment cultures that incubated at 30 °C for two weeks. Thirteen isolated strains that show high growth rate on crude oil were selected for further study. These strains were first identified by classical biochemical tests. Four strains identified molecularly by amplification and sequencing the 16S rRNA gene sequencing and comparing them to the database of known 16S rRNA sequences. The results of the identification procedure showed that four isolated bacteria belong to: Shewanella algae isolate BHA1 (obtained from C. gigas mussel), Micrococcus luteus isolate BHA7 (obtained from C. asperella mussel), Pseudoalteromonas sp. isolate BHA8 (obtained from B. tenella mussel) and Shewanella haliotis isolate BHA35 (obtained from C. gigas mussel). All sequences of four bacteria were submitted to the Genetic Sequence Database at the National Center for Biotechnology Information (NCBI). The GenBank IDs of these strains in NCBI are LK391612 (for BHA1), LK391613 (for BHA7), LK391614 (for BHA8) and LK391623 (for BHA35). The phylogenic trees of these four isolated strains were illustrated in Fig. 2. This figure shows that strains BHA35 and BHA1 have high similarity with other sequences that are present in Gene bank database, although the similarity of other strains is less compared to these two strains. 3.3. Growth rate, crude-oil removal, cell surface hydrophobicity and emulsification activity of prevalent isolated strains All bacterial strains were grown in 1% crude oil for 15 days in a shaker incubator. After 15 days, the levels of microbial growth and crude oil biodegradation were monitored using spectrometry and GC-FID methods. As reported in Table 1, the strain BHA35 exhibits highest level (69.17%) and the strain BHA8 exhibits the lowest levels (27.13%) of crude-oil degradation between all isolated strains. The GC-FID chromatograms for these strains in comparison to blank confirm that peaks of crude oil compounds were decreased dramatically by these strains. Emulsification activity and bacterial adhesion to hydrocarbon (BATH) were analyzed for each strain separately. Data obtained from these tests were presented in Table 1. Strain BHA1 has the highest values of emulsification activity (E24: 79%) and the high values of cell surface hydrophobicity relate to strain BHA35 (BATH: 36.3%). 3.4. Effect of different factors on degradation of crude oil by prevalent isolated strains Results obtained from the effect of different concentration of NaCl on crude oil degradation by the tested strains are represented in Table 2. As shown in this table, the optimum concentration of NaCl for highest degradation of crude oil by these isolates was 2–4%. However, these strains can tolerate high concentrations of salt up to 12% (w/v). The effects of mixed bacterial consortia in comparison to individual isolates were presented in Table 3. The data in this table confirmed that the capacity of oil degradation by the mixed bacterial consortia was better than individual bacteria. Effect of different concentrations of crude oil on degradation was determined. The results were presented in Fig. 3. As shown in this figure, the highest degradation of crude oil takes place at 2.5% concentration however crude oil degradation decreased when concentrations of crude oil were increased. The effect of incubation time on crude oil degradation was shown in Fig. 3. It has been observed that the oil degradation at 30 days was higher than samples incubated for 15 days only.
4. Discussion Biodegradation of crude oil by bacteria has been recently reported at soil and aquatic ecosystems. The ability of hydrocarbon degrading bacteria has been recognized in the past decade but little is known about symbiotic bacterial populations in marine animals. Thomas et al. (2014) isolated 72 oil-degrading bacteria at waters, sediments, permeable beach sands, coastal wet lands, and marine life in the Gulf of Mexico. These researchers found that the majority of oil-degrading bacteria corresponded to Pseudomonas. Some crude oil degrading bacteria was isolated from Persian Gulf by different researcher (Hassanshahian et al., 2014b; Tebyanian et al., 2013). For example, Radwan et al. (2005) isolate Pseudomonas, Bacillus and Acinetobacter as crude oil degrader from this Gulf. Hassanshahian et al. (2012a) describe 12 crude oil degrading bacteria from Persian Gulf. In the current study, we have isolated 28 hydrocarbon-degrading bacteria symbiotic with mussels (C. gigas, C. asperella and B. tenella) and surrounding environments. Then we have compared quantity of microbial communities in all samples. The enumeration result of crude oil degrading bacteria and heterotrophic bacteria in this research confirmed that the quantity of bacteria was higher in the mussel samples compared to the surrounding environment. Some researchers also report similar results. Kueh and Chan (1985) suggested that differences exist between microbial communities of the surrounding seawater and mussels (C. gigas). In other study, Cavallo et al. (2009) reported that heterotrophic bacterial densities and diversity were higher in the mussel samples (Mytilus galloprovincialis) than in the surrounding seawater. In fact, mussel samples accumulated bacteria by filter feeding. Also, Silverman et al. (1996) report filtration activity by zebra mussel, this ability causes a change in the microbial communities and biodiversity of marine bacteria. The biodiversity of bacterial strain that isolated in this study was less compared to other researchers that work in the Persian Gulf (Radwan et al., 2005). This low diversity of crude oil degrading bacteria in this research can be interpreted as our sampling zone was limited, also the sampling sites was chronically polluted with crude oil and the level of crude oil pollution in the sampling sites was very high. It is possible that high crude oil contamination will result in the disappearance of bacteria in the collected mussel, seawater and sediment samples or crude oil contamination will cause decrease in the diversity of bacteria. The ability of the isolated strains to degrade crude oil was measured in this research. The results confirmed that the best strain for crude oil degradation was S. haliotis isolate BHA35 (69.17%). For the first time crude oil degrading bacteria of the genus Shewanella were isolated from mussels. The scale of crude oil degradation by microbial population depends on physical and chemical factors. The results of this research propose optimum condition for crude oil degradation as: 2.5% concentration of crude oil, 2–4% (w/v) concentration of NaCl, 30 days incubation time and mixed bacterial consortia. Fusey and Oudot (1984) reported that when the concentrations of crude oil were increased hydrocarbon biodegradation decreased. This pattern can be related to oxygen or nutrients limitation in higher concentrations of crude oil. Hasanshahian and Emtiazi (2008) demonstrated that maximum crude oil removal levels have been carried out at 1% concentration. Zahed et al. (2010) reported that most of the crude oil removal occurred at 100 mg/L and lower concentrations of crude oil are more efficient for crude oil removal. In fact with increasing concentration of crude oil and NaCl the hydrocarbon biodegradation decreased (Margesin and Schinner, 2001). This study also agrees with research of Kumar et al. (2007) which concluded that a negative relationship exists between salinity and hydrocarbon biodegradation. Mnif et al. (2009) observed that strain Halomonas sp. C2SS100 is able to degrade crude oil in the presence of 100 g/l NaCl. In this study the isolated strains tolerate high concentrations of salt (12% w/v) thus suitable for bioremediation of oil pollution in marine environments with high salinity such as Persian
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
Gulf. Sathishkumar et al. (2008) suggested that crude oil degradation by microbial consortia is more effective in comparison to individual bacteria. They interpreted this result as in the microbial consortia there were different enzymes for various compounds of crude oil then the degradation was faster and has high performance. 5. Conclusions The results of this study confirmed the presence of crude oil degrading bacteria with mussels that collected from the Persian Gulf. These crude oil degrading bacteria have high biodegradation activity and have high tolerance to NaCl and crude oil. In addition, higher incubation time, mixed bacterial consortia and emulsifier production by strains improved the efficiency of crude oil degradation. Acknowledgment This work was supported financially by Shahid Bahonar University of Kerman. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.marpolbul.2015.11.021. References Al-Awadhi, H., Al-Hasan, R.H., Sorkhoh, N.A., Radwan, S.S., 2003. Establishing oildegrading biofilms on gravel particles and glass plates. Int. Biodeterior. Biodegrad. 51 (3), 181–189. Al-Mailem, D.A., Sorkhoh, N.A., Salamah, S., Eliyas, M., Radwan, S.S., 2010. Oilbioremediation potential of Arabian Gulf mud flats rich in diazotrophic hydrocarbon-utilizing bacteria. Int. Biodeterior. Biodegrad. 64 (3), 218–225. Cappello, S., Genovese, M., Della Torre, C., Crisari, A., Hassanshahian, M., Santisi, S., Calogero, R., Yakimov, M.M., 2012a. Effect of bioemulsificant exopolysaccharide EPS2003 on microbial community dynamics during assays of oil spill bioremediation: a microcosm study. Mar. Pollut. Bull. 64 (12), 2820–2828. Cappello, S., Russo, D., Santisi, S., Calogero, R., 2012b. Presence of hydrocarbon-degrading bacteria in the gills of mussel Mytilus galloprovincialis in a contaminated environment: a mesoscale simulation study. Chem. Ecol. 28 (3), 239–252. Cavallo, R.A., Acquaviva, M.I., Stabili, L., 2009. Culturable heterotrophic bacteria in sea water and Mytilus galloprovincialis from a Mediterranean area (Northern Ionian Sea — Italy). Environ. Monit. Assess. 149, 465–475. Dave, D., Ghaly, A.E., 2011. Remediation technologies for marine oil spills: a critical review and comparative analysis. Am. J. Environ. Sci. 7 (5), 423–440. Fuchsluger, C., Preims, M., Fritz, I., 2010. Automated measurement and quantification of heterotrophic bacteria in water samples based on the MPN method. J. Ind. Microbiol. Biotechnol. 38, 241–247. Fusey, P., Oudot, J., 1984. Relative influence of physical removal and biodegradation in the depuration of petroleum-contaminated seashore sediments. Mar. Pollut. Bull. 15, 136–141. Ghanavati, H., Emtiazi, G., Hassanshahian, M., 2008. Synergism effects of phenol degrading yeast and ammonia oxidizing bacteria for nitrification in coke wastewater of Esfahan steel company. Waste. Manage. Res. 26 (2), 203–208. Hasanshahian, M., Emtiazi, G., 2008. Investigation of alkane biodegradation using the microtiter plate method and correlation between biofilm formation, biosurfactant production and crude oil biodegradation. Int. Biodeterior. Biodegrad. 62, 170–178. Hassanshahian, M., Emtiazi, G., Kermanshahi, R., Cappello, S., 2010. Comparison of oil degrading microbial communities in sediments from the Persian Gulf and Caspian Sea. Soil. Sediment. Contaminat. 19 (3), 277–291. Hassanshahian, M., Emtiazi, G., Cappello, S., 2012a. Isolation and characterization of crude-oil-degrading bacteria from the Persian Gulf and the Caspian Sea. Mar. Pollut. Bull. 64, 7–12. Hassanshahian, M., Tebyanian, H., Cappello, S., 2012b. Isolation and characterization of two crude-oil degrading yeast strains, Yarrowia lipolytica PG-20 and PG-32 from Persian Gulf. Mar. Pollut. Bull. 64, 1386–1391. Hassanshahian, M., Ahmadinejad, M., Tebyanian, H., Kariminik, A., 2013. Isolation and characterization of alkane degrading bacteria from petroleum reservoir waste water in Iran (Kerman and Tehran provenances). Mar. Pollut. Bull. 73, 300–305.
91
Hassanshahian, M., Zeynalipour, M.S., Hosseinzadeh Musa, F., 2014a. Isolation and characterization of crude oil degrading bacteria from the Persian Gulf (Khorramshahr provenance). Mar. Pollut. Bull. 82, 39–44. Hassanshahian, M., Emtiazi, G., Caruso, G., Cappello, S., 2014b. Bioremediation (bioaugmentation/biostimulation) trials of oil polluted seawater: a mesocosm simulation study. Mar. Environ. Res. 95, 28–38. Hassanshahian, M., Yakimov, M.M., Denaro, R., Genovese, M., Cappello, S., 2014c. Using Real-Time PCR to assess changes in the crude oil degrading microbial community in contaminated seawater mesocosms. Inte. Biodeterior. Biodegrad. 93, 241–248. Hassanshahian, M., 2014d. Isolation and characterization of biosurfactant producing bacteria from Persian Gulf (Bushehr provenance). Mar. Pollut. Bull. 86, 361–366. Holt, S.G., Kriey, N.R., Sneath, P.H.A., Staley, J.T., Williams, S.T., 1998. Bergy's Manual of Determinative for Bacteriology. Williams and Wilkins, New York. Huber, M., 2010. Compendium of Bivalves, A Full-Color Guide to 3300 of the World's Marine Bivalves, a Status on Bivalvia After 250 Years of Research. Conch Books, Hackenheim, Germany (904 pp.). Kaczorek, E., Chrzanowski, L., Pijanowska, A., Olszanowski, A., 2008. Yeast and bacteria cell hydrophobicity and hydrocarbon biodegradation in the presence of natural surfactants: rhamnolipides and saponins. Bioresour. Technol. 99 (10), 4285–4291. Khan, K., Naeem, M., Asif, M., 2006. Extraction and characterization of oil degrading bacteria. J. Appl. Sci. 6 (10), 2302–2306. Kueh, C.S., Chan, K.Y., 1985. Bacteria in bivalve shellfish with special reference to the oyster. J. Appl. Bacteriol. 59, 41–47. Kumar, M., Leon, V., Manterano, A.D.S., Ilzins, O.A., 2007. A halotolerant and thermotolerant Bacillus sp. degrades hydrocarbons and produces tensio-active emulsifying agent. World J. Microbiol. Biotechnol. 23, 211–220. Manganaro, A., Pulicano, G., Reale, A., Sanfilippo, M., Sara, G., 2009. Filtration pressure by bivalves affects the trophic conditions in Mediterranean shallow ecosystems. Chem. Ecol. 25, 467–478. Margesin, R., Schinner, F., 2001. Bioremediation (natural attenuation and biostimulation) of diesel-oil-contaminated soil in an alpine glacier skiing area. Appl. Environ. Microbiol. 67, 3127–3133. Mnif, S., Chamkha, M., Sayadi, S., 2009. Isolation and characterization of Halomonas sp. Strain C2SS100, a hydrocarbon-degrading bacterium under hypersaline conditions. J. Appl. Microbiol. 107, 785–794. Pruthi, V., Cameotra, S.S., 1997. Rapid identification of biosurfactant-producing bacterial strains using a cell surface hydrophobicity technique. Biotechnol. Tech. 11, 671–674. Radwan, S., Al-Hasan, R.H., Al-Dabbous, S., Salamah, S., 2002. Bioremediation of oily sea water by bacteria immobilized in biofilms coating macroalgae. Int. Biodeterior. Biodegrad. 50 (1), 55–59. Radwan, S.S., Al-Hasan, R.H., Salamah, A., Khanafer, M., 2005. Oil-consuming microbial consortia floating in the Arabian Gulf. Int. Biodeterior. Biodegrad. 56, 28–33. Rahman, K.S.M., Thahira-Rahman, J., Lakshmanaperumalsamy, P., Banat, I.M., 2004. Towards efficient crude oil degradation by a mixed bacterial consortium. Bioresour. Technol. 85, 257–261. Sathishkumar, M., Binupriya, A., Baik, S.H., Yun, R.E., 2008. Biodegradation of crude oil by individual bacterial strains and a mixed bacterial consortium isolated from hydrocarbon contaminated areas. Clean 36 (1), 92–96. Saxena, R., Gothawal, R., Khan, M.N., 2013. Prospective in-silco approach in bioremediation of petroleum hydrocarbon: success so far. Octa J. Environ. Res. 1 (2), 137–149. Silverman, H., Lynn, J.W., Achebergee, C.R., Dietz, T.H., 1996. Gill structure in zebra mussels: bacterial-sized particle filtration. Am. Zool. 36, 373–384. Sorkhoh, N.A., Ghannoum, M.A., Ibrahim, A.S., Stretton, R.J., Radwan, S.S., 1990. Crude oil and hydrocarbon-degrading strains of Rhodococcus rhodochrous isolated from soil and marine environments in Kuwait. Environ. Pollut. 65 (1), 1–17. Stabili, L., Acquaviva, M.I., Cavallo, R.A., 2005. Mytilus galloprovincialis filter feeding on the bacterial community in a Mediterranean coastal area (Northern Ionian Sea, Italy). Water Res. 39, 469–477. Tebyanian, H., Hassanshahian, M., Kariminik, A., 2013. hexadecane-degradation by Teskumurella and Stenotrophomonas strains iso lated from hydrocarbon contaminated soils. Jundishapur. J. Microbiol. 26 (7), e9182. Thomas, J.C., Wafula, D., Chauhan, A., Green, S.J., Gragg, R., Jagoe, C., 2014. A survey of deepwater horizon (DWH) oil-degrading bacteria from the Eastern oyster biome and its surrounding environment. Front. Microbiol. 5, 149. Todd, P.A., Ong, X., Chou, L.M., 2010. Impacts of pollution on marine life in Southeast Asia. Biodivers. Conserv. 19 (4), 1063–1082. Udeh, N.U., Nwaogazie, I.L., Momoh, Y., 2013. Bioremediation of a crude oil contaminated soil using water hyacinth (Eichhornia crassipes). Adv. Appl. Sci. Res. 4 (2), 362–369. Yakimov, M.M., Timmis, K.N., Golyshin, P.N., 2007. Obligate oil-degrading marine bacteria. Curr. Opin. Biotechnol. 18 (3), 257–266. Zahed, M.A., Hamidi, A.A., Isa, M.H., Mohajeri, L., 2010. Effect of initial oil concentration and dispersant on crude oil biodegradation in contaminated seawater. Bull. Environ. Contam. Toxicol. 84, 438–442. Zhou, Y., Zhang, S., Liu, Y., Yang, H., 2014. Biologically induced deposition of fine suspended particles by filter-feeding bivalves in land-based industrial marine aquaculture wastewater. PLoS One 9 (9).
Contents lists available at ScienceDirect
Marine Pollution Bulletin journal homepage: www.elsevier.com/locate/marpolbul
Enrichment and isolation of crude oil degrading bacteria from some mussels collected from the Persian Gulf Zeynab Bayat, Mehdi Hassanshahian ⁎, Majid Askari Hesni Department of Biology, Faculty of Sciences, Shahid Bahonar University of Kerman, Kerman, Iran
a r t i c l e
i n f o
Article history: Received 12 June 2015 Received in revised form 5 November 2015 Accepted 6 November 2015 Available online 12 November 2015 Keywords: Crude oil Marine environment Mussel Persian Gulf Pollution
a b s t r a c t To date, little is known about existing relationships between mussels and bacteria in hydrocarbon-contaminated marine environments. The aim of this study is to find crude oil degrading bacteria in some mussels at the Persian Gulf. Twenty eight crude oil degrading bacteria were isolated from three mussels species collected from oil contaminated area at Persian Gulf. According to high growth and degradation of crude oil four strains were selected between 28 isolated strains for more study. Determination the nucleotide sequence of the gene encoding for 16S rRNA show that these isolated strains belong to: Shewanella algae isolate BHA1, Micrococcus luteus isolate BHA7, Pseudoalteromonas sp. isolate BHA8 and Shewanella haliotis isolate BHA35. The residual crude oil in culture medium was analysis by Gas Chromatography (GC). The results confirmed that these strains can degrade: 47.24%, 66.08%, 27.13% and 69.17% of crude oil respectively. These strains had high emulsification activity and biosurfactant production. Also, the effects of some factors on crude oil degradation by isolated strains were studied. The results show that the optimum concentration of crude oil was 2.5% and the best degradation take place at 12% of salinity. This research is the first reports on characterization of crude oil degrading bacteria from mussels at Persian Gulf and by using of these bacteria in the field the effect of oil pollution can be reduce on this marine environment. © 2015 Elsevier Ltd. All rights reserved.
1. Introduction Many techniques are utilized to mitigate or cleanse petroleum pollution in the environment. During exploration, production, refining, transport and storage of petroleum and petroleum products, some accidental spill may be released into the sea waters (Dave and Ghaly, 2011; Ghanavati et al., 2008). Crude oil is toxic, mutagenic and carcinogenic this pollutant concedes a serious damage to marine life (Todd et al., 2010). Bioremediation is the best method for elimination of oil spills. Bioremediation of oil contamination compared with physicochemical treatment is more effective and lower cost price (Saxena et al., 2013). Bacteria are important for the biodegradation of petroleum hydrocarbons and many hydrocarbon-degrading bacteria have been isolated from different environments (Kaczorek et al., 2008; Udeh et al., 2013; Hassanshahian et al., 2012a; Radwan et al., 2002; Khan et al., 2006). Biodegradation rates can be controlled by concentration and composition of hydrocarbons, nutrients, oxygen, salt concentrations, moisture and temperature (Sathishkumar et al., 2008; Hassanshahian et al., 2010). Also, biosurfactants increase the solubility of hydrocarbons to bacteria and enhanced degradation of these toxic compounds (Hassanshahian et al., 2012a). Marine organisms can take up contaminants from bottom
⁎ Corresponding author. E-mail address: [email protected] (M. Hassanshahian).
http://dx.doi.org/10.1016/j.marpolbul.2015.11.021 0025-326X/© 2015 Elsevier Ltd. All rights reserved.
sediments, suspended particulate material and food sources. Filterfeeding bivalves such as mussels, oysters, and clams are very important components in the process of bioremediation of the marine environment (Stabili et al., 2005). They are capable to filter and accumulate a large number of fine particulate matters, including phytoplankton, zooplankton, microorganism and other particulate organic debris. Furthermore, the organic components in the suspended matters can be trapped and applied by filter-feeding bivalves (Manganaro et al., 2009; Zhou et al., 2014; Hassanshahian et al., 2013). Little work has been done on the distribution and physiology of autochthonous hydrocarbondegrading microbes inhabiting marine organisms (Cappello et al., 2012a; Hassanshahian et al., 2012b). Some researchers confirmed that crude oil degrading bacteria were existing in marine organisms. For example, Radwan et al. (2005) establish that some crude oil degrading bacteria such as Pseudomonas, Bacillus and Acinetobacter were associated with cyanobacterial mat at Arabian Gulf (Radwan et al., 2005; Radwan et al., 2002). Also, Al-Mailem et al. (2010) study crude oil degrading bacteria that symbiosis with filamentous cyanobacteria and their results show that some bacteria such as Halomonas aquamarina, Marinobacter hydrocarbonoclasticus, Marinobacter sp.; Dietzia maris and Alcanivorax can degrade crude oil and present in cyanobacteria (Al-Mailem et al., 2010; Sorkhoh et al., 1990; Al-Awadhi et al., 2003). In all research that studied the relationship between bacteria and marine animals, it is confirmed that the density of bacteria in the marine animal samples were higher compared to surrounding water. For
86
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
Fig. 1. Enumeration of heterotrophic and hydrocarbon-degrading bacteria by CFU and MPN methods. All data present in this figure are the average of three replicates of enumeration in the same dilution.
example, mussels can filter the seawater and concentrate the marine bacteria that cause reducing the bacterial concentration in the seawater (Cavallo et al., 2009). To date, little is known about existing relationships between mussels and bacteria in hydrocarbon-contaminated marine environments. The aim of this study is to find crude oil degrading bacteria in some mussels at the Persian Gulf. Also identification of these strains and the degradation capacity of these isolates is another purpose of this research.
related to: Crassostrea gigas, Chama asperella and Barbatia tenella genus. Collected samples were transported on ice to the laboratory. Mussels shell was removed by sterile knife and gill and other tissue of mussel samples was washed with sterile seawater. In fact, the tissue was homogenized in buffer. Finally, macerate of mussels was used for subsequent studies.
2. Materials and methods
Crude oil degrading bacteria were isolated in ONR7a medium supplemented with 1% (v/v) of crude oil (Iranian light crude oil) as sole carbon source and energy. ONR7a contained (per liter of distilled water) 40 g of NaCl, 11.18 g of MgCl2·6H2O, 3.98 g of Na2SO4, 1.46 g of CaC12·2H2O, 1.3 g of TAPS0 {3-[N tris(hydroxymethyl) methylamino]-2 hydroxypropanesulfonic acid}, 0.72 g of KCl, 0.27 g of NH4Cl, 89 mg of Na2HP04·7H20, 83 mg of NaBr, 31 mg of NaHCO3, 27 mg of H3BO3, 24 mg of SrCl2·6H20, 2.6 mg of NaF and 2 mg of FeCl2·4H20. For solid media, Bacterial Agar (15 g/l) was added to the solution (Hasanshahian and Emtiazi, 2008). Tissue macerate of mussel (5 ml), condensed seawater (5 ml) and portion of sediments (10 g) were added to Erlenmeyer flasks containing
2.1. Sampling Mussels, seawater and sediment samples were collected from two oil contaminated sites at Persian Gulf. These two stations were located in the Qeshm island (AZ: Zakeri Harbor; 36°15, N; 34°15, E) and (PZ: Park Ziton 37°30, N; 49°15, E). Mussels were collected from depth range 5–15 m. Also, samples of seawater from above the mussels' bed in a sterile bottle, and samples of sediment from below were collected. The mussels were identified according to the standard key (Huber, 2010). The results of identification confirmed that these three mussels
2.2. Isolation and selection of crude-oil degrading bacteria
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
87
Fig. 2. Phylogenetic tree of 16S rDNA sequences of the BHA1, BHA7, BHA8 and BHA35 isolates obtained from Persian Gulf. The tree was constructed using sequences of comparable region of the 16S rDNA gene sequences available in public databases. Neighbor-joining analysis using 1000 bootstrap replicates was used to infer tree topology. The bar represents 0.02% sequence divergence.
100 ml of medium and the flasks were incubated for 7 days at 30 °C on rotary shaker (180 rpm, INFORS AG). Then 5 ml were transported to fresh medium. After a series of four further subcultures, inoculums from the flask were streaked out, and phenotypically different colonies
on ONR7a agar were purified. Phenotypically different colonies obtained from the plates were transferred to fresh medium with and without crude oil to eliminate autotrophic and agar utilizing bacteria. The procedure was repeated, and only isolates exhibiting pronounced growth on
Table 1 Growth rate, crude oil removal, emulsification activity and BATH assay for isolated strains. Isolate
BHA1 BHA7 BHA8 BHA35
Growth rate (OD600 nm)
Percentage of oil removal (Spectrometry)
Percentage of oil removal (GC method)
Emulsification activity (E24%)
Cell surface hydrophobicity (BATH %)
Average
SD⁎
Average
SD
Average
SD
Average
SD
Average
SD
0.76 0.81 0.91 0.84
0.124 0.163 0.137 0.081
58.4 55.87 19.74 40
1.71 1.22 1.12 1.61
47.24 66.08 27.13 69.17
1.65 0.85 1.31 0.81
79 25 71 77
0.72 0.42 1.1 0.72
12.9 0 0 36.3
0.56 0 0 1.4
⁎ Standard Deviation (SD).
88
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
Table 2 The effect of different concentrations of NaCl on growth of selected bacterial strains. NaCl concentration (%)
0
20
40
60
80
100
120
140
160
180
200
++++ ++++ − +++
++++ ++++ ++++ ++++
++++ ++++ ++++ ++++
+++ +++ +++ +++
+++ +++ +++ +++
++ ++ ++ ++
+ ++ + ++
− + − −
− + − −
− + − −
− + − −
strain Shewanella algae BHA1 Micrococcus luteus BHA7 Pseudoalteromonas sp. BHA8 Shewanella haliotis BHA35
crude oil were stored in stock media with glycerol at −20 °C for further characterization (Hasanshahian and Emtiazi, 2008).
motility, starch hydrolysis, indole, H2S production, catalase and oxidase, oxidation/fermentation, reduction of nitrate, growth and acidification of carbohydrates tests were performed (Holt et al., 1998).
2.3. Enumeration of bacteria 2.3.1. Quantification the number of cultivable heterotrophic and hydrocarbon degrading bacteria Measurements of bacterial abundance within the mussel and surrounding environment were performed by colony-forming units (CFU) and Most Probable Number (MPN) procedures (Cappello et al., 2012b; Fuchsluger et al., 2010). Heterotrophic bacteria in the mussel, seawater and sediment were estimated by spreading 100 μl of 10-fold diluents flask on plates of Marine Agar medium (MA) and incubating at 30 °C for 3 days. Also, crude oil degrading bacteria in the mussels, seawater and sediment were estimated by spreading 100 μl of 10-fold diluents flask on plates of ONR7a agar medium with crude oil and incubating at 30 °C for 7 days. The bacteria quantification results were expressed as CFU·g− 1 (Cappello et al., 2012b; Hassanshahian et al., 2014a). 2.3.2. Most probable number (MPN) of heterotrophic and hydrocarbon degrading bacteria The MPN method for hydrocarbon-degrading bacteria was done in sterile 24-well microplates using sample aliquots with corresponding dilutions and cultivation media. 1700 μl ONR7a medium was provided in each well of the microplates then dilution series of samples (usually tenfold dilution until 10−1–10−3) were prepared in ONR7a medium. Wells were inoculated with 100 μl of sample diluents. Following sample inoculation, 100 μl of sterile Iranian light crude oil was applied at the centre of each well. Plates were incubated at 30 °C for 21 days. After incubation, visual evaluation of microbial growth was done. On basis, the number of positive test tubes and the counts of microorganisms were statistically evaluated (Fuchsluger et al., 2010). The MPN method for heterotrophic bacteria was done in sterile 24-well microplates. Each well of the microplates was provided with 1700 μl Marine Broth (MB) and 100 μl dilution 10−3–10−4 of sample, and then plates were incubated at 30 °C for 14 days. All samples from each station were used for MPN count. MPN counts were analyzed with the computer program MPN calculator version 4.2 (Fuchsluger et al., 2010; Hassanshahian, 2014d).
2.4.2. Molecular identification Analysis of 16S rRNA was performed to taxonomically characterize the isolated strains. Total DNA of bacterial strains was extracted with the CTAB method. PCR amplification of 16S rRNA genes was performed using the general bacteria primer 27F (5-AGAGTTTGATCCTGGCTCAG-3) and universal reverse primer 1492R (5-TACGYTACCTTGTTACGACTT-3) (Cappello et al., 2012b). The amplification reaction was carried out in a total volume of 25 μl consisting, 2 mM MgCl2 (1 μl), 10× PCR reaction buffer (200 mM Tris; 500 mM KCl) (2.5 μl), 2 mM each dNTP (2 μl), 0.15 mM each primer (1 μl), 1 U (0.5 μl) Taq DNA polymerase (Qiagen, Hilden, Germany) and 2 μl of template DNA (50 pmol). Total volume was brought up to 15 μl using sterile milliQ water. PCR program consisted of 35 cycles was performed in a thermal cycler GeneAmp 5700 (PE Applied Biosystem, Foster City, CA, USA). The temperature profile for PCR was kept, 94 °C for 5 min, 94 °C for 1 min, 54 °C for 1 min, 72 °C for 1 min, 30 cycles; then 72 °C for 10 min and finally storage at 4 °C. The 16S rRNA amplified was sequenced with Big Dye terminator V3.1 cycle sequencing kit on an automated capillary sequencer (model 3100 Avant Genetic Analyzer, Applied Biosystems). Similarity rank from the Ribosomal Database Project RDP) and FASTA Nucleotide Database Query were used to determine partial 16S rRNA sequences to estimate the degree of similarity to other 16S rRNA gene sequences. Analysis and phylogenetic affiliates of sequences was performed as following the protocol described by Yakimov et al. (2007). Phylogenetic tree was drawn by MEGA5 software with neighbor joining method. 2.5. Measure of bacterial growth and crude oil degradation Bacterial isolates were grown at 30 °C for 15 days on rotator shaker (180 rpm). The growth of the isolates was routinely assessed indirectly by measuring the turbidity (OD600 nm) using UV–Visible spectrophotometer (Shimadzu UV-160, Japan). The crude oil removal assay was carried out by dissolving the residual crude oil in the medium in dichloromethane (DCM) and reading the optical density of the oil extract against blank (distill water) at wavelength of 420 nm (Rahman et al., 2004).
2.4. Identification of the isolates 2.6. Gas chromatography (GC) of residual crude oil 2.4.1. Biochemical characterization The following characteristics were determined according to the “Bergey's Manual of Determinative Bacteriology: the Gram stain, Table 3 The effect of individual and mixed bacterial consortia on growth and crude oil degradation by selected bacterial strains. BHA1
Growth rate (OD600 nm) Oil degradation (%) ⁎ Standard Deviation.
BHA35
BHA1 + BHA35
Average
SD⁎
Average
SD
Average
SD
0.90 67.31
0.04 0.816
1.17 73.45
0.138 1.63
1.78 88.69
0.204 1.86
Crude oil degradation was accuracy estimated by GC-FID. The residual crude oil was extracted in each samples. The extraction protocol was as follow: the same volume of DCM was added to each flask and residual crude oil was extracted then, treated with anhydrous sodium sulfate (Na2SO4) to remove residual water. Extracts were concentrated by separating funnel (Hassanshahian et al., 2014c). Analyses were done by GC-FID (Varian 3800 model, USA) equipped with a SE-54 capillary column (25 m × 0.32 mm × 0.1 μm) and flame ionization detector (FID). Helium was used as the carrier gas (30 ml/min). The oven was programmed as follows: 100 °C (1 min) then increased to 300 °C (2 min) at a rate of 30 °C min−1. The samples were quantified according to previously described protocols (Hassanshahian et al., 2014).
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
89
Fig. 3. The effect of crude oil concentrations on removal of crude oil by isolated strains (A), the effect of incubation time on crude oil degradation by isolated strains (B).
2.7. Measure of emulsification activity and bacterial adherence to hydrocarbons (BATH) The emulsification activity (E24) was determined by the addition of hexadecane to the same volume of cell free culture broth. After mixing with vortex for 2 min and leaving to stand for 24 h, the E24 index is given as percentage of height of emulsified layer (in millimeters) divided by total height of the liquid column (in millimeters). Measurement of the bacterial adhesion to hydrocarbon was performed as described by Pruthi and Cameotra (1997).
inoculated with bacterial culture and incubated at 30 °C and 180 rpm for 15 days (Kumar et al., 2007). To study the effect of incubation time on the growth and degradation of crude oil by the selected bacterial strains samples were incubated for 15 and 30 days. The individual and mixed bacterial consortia from overnight culture were transferred to ONR7a medium with 1% (v/v) of crude oil at 30 °C and 180 rpm. Bacterial growth and crude oil degradation measurement were carried out respectively at 600 nm and 420 nm. The experiments were conducted in triplicates and the average values were considered (Sathishkumar et al., 2008). 3. Results
2.8. Effect of different factors on crude oil degradation by isolated strains Different factors were examined for optimization of crude oil degradation. These factors were concentrations of crude oil, NaCl concentration, incubation time and mixed bacterial consortia. The effect of different concentrations of crude oil (1%, 2.5%, 4% and 5.5%) on the growth and degradation of crude oil by the selected bacterial strains was studied using ONR7a medium at 30 °C and 180 rpm for 15 days (Khan et al., 2006). The influence of salinity tolerance on the growth and degradation of crude oil by the selected bacterial strains was carried out using ONR7a medium with 1% (v/v) of crude oil and 0–20% (w/v) concentration of NaCl. The flasks were
3.1. Quantity of heterotrophic and crude oil degrading bacteria in collected samples (CFU and MPN) The quantity of heterotrophic and crude oil degrading bacteria was determined in all collected samples by two enumeration methods: CFU and MPN. The results were shown in Fig. 1. As shown in this figure, the numbers of heterotrophic bacteria in AZ 1 and PZ 1 were the highest (respectively 1.64 × 106, 1.08 × 106 CFU·ml− 1) compared to other samples. AZ 1 and PZ 1 samples have the highest abundance of crude oil degrading bacteria (respectively 5 × 103, 2 × 103 CFU·ml−1). Also, the MPN value for samples AZ 1 and PZ 1 (respectively 6.9 × 103,
90
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
8 × 103) has the highest number of crude oil degrading bacteria compared to other samples and the highest quantity (MPN) of heterotrophic bacteria related to AZ 1 and AZ 3 samples (respectively 4.6 × 106, 3.7 × 106). Also, Fig. 1 illustrate that the MPN values in all samples were higher than CFU values. 3.2. Isolation and identification of bacteria Twenty eight crude oil degrading bacteria were isolated after enrichment cultures that incubated at 30 °C for two weeks. Thirteen isolated strains that show high growth rate on crude oil were selected for further study. These strains were first identified by classical biochemical tests. Four strains identified molecularly by amplification and sequencing the 16S rRNA gene sequencing and comparing them to the database of known 16S rRNA sequences. The results of the identification procedure showed that four isolated bacteria belong to: Shewanella algae isolate BHA1 (obtained from C. gigas mussel), Micrococcus luteus isolate BHA7 (obtained from C. asperella mussel), Pseudoalteromonas sp. isolate BHA8 (obtained from B. tenella mussel) and Shewanella haliotis isolate BHA35 (obtained from C. gigas mussel). All sequences of four bacteria were submitted to the Genetic Sequence Database at the National Center for Biotechnology Information (NCBI). The GenBank IDs of these strains in NCBI are LK391612 (for BHA1), LK391613 (for BHA7), LK391614 (for BHA8) and LK391623 (for BHA35). The phylogenic trees of these four isolated strains were illustrated in Fig. 2. This figure shows that strains BHA35 and BHA1 have high similarity with other sequences that are present in Gene bank database, although the similarity of other strains is less compared to these two strains. 3.3. Growth rate, crude-oil removal, cell surface hydrophobicity and emulsification activity of prevalent isolated strains All bacterial strains were grown in 1% crude oil for 15 days in a shaker incubator. After 15 days, the levels of microbial growth and crude oil biodegradation were monitored using spectrometry and GC-FID methods. As reported in Table 1, the strain BHA35 exhibits highest level (69.17%) and the strain BHA8 exhibits the lowest levels (27.13%) of crude-oil degradation between all isolated strains. The GC-FID chromatograms for these strains in comparison to blank confirm that peaks of crude oil compounds were decreased dramatically by these strains. Emulsification activity and bacterial adhesion to hydrocarbon (BATH) were analyzed for each strain separately. Data obtained from these tests were presented in Table 1. Strain BHA1 has the highest values of emulsification activity (E24: 79%) and the high values of cell surface hydrophobicity relate to strain BHA35 (BATH: 36.3%). 3.4. Effect of different factors on degradation of crude oil by prevalent isolated strains Results obtained from the effect of different concentration of NaCl on crude oil degradation by the tested strains are represented in Table 2. As shown in this table, the optimum concentration of NaCl for highest degradation of crude oil by these isolates was 2–4%. However, these strains can tolerate high concentrations of salt up to 12% (w/v). The effects of mixed bacterial consortia in comparison to individual isolates were presented in Table 3. The data in this table confirmed that the capacity of oil degradation by the mixed bacterial consortia was better than individual bacteria. Effect of different concentrations of crude oil on degradation was determined. The results were presented in Fig. 3. As shown in this figure, the highest degradation of crude oil takes place at 2.5% concentration however crude oil degradation decreased when concentrations of crude oil were increased. The effect of incubation time on crude oil degradation was shown in Fig. 3. It has been observed that the oil degradation at 30 days was higher than samples incubated for 15 days only.
4. Discussion Biodegradation of crude oil by bacteria has been recently reported at soil and aquatic ecosystems. The ability of hydrocarbon degrading bacteria has been recognized in the past decade but little is known about symbiotic bacterial populations in marine animals. Thomas et al. (2014) isolated 72 oil-degrading bacteria at waters, sediments, permeable beach sands, coastal wet lands, and marine life in the Gulf of Mexico. These researchers found that the majority of oil-degrading bacteria corresponded to Pseudomonas. Some crude oil degrading bacteria was isolated from Persian Gulf by different researcher (Hassanshahian et al., 2014b; Tebyanian et al., 2013). For example, Radwan et al. (2005) isolate Pseudomonas, Bacillus and Acinetobacter as crude oil degrader from this Gulf. Hassanshahian et al. (2012a) describe 12 crude oil degrading bacteria from Persian Gulf. In the current study, we have isolated 28 hydrocarbon-degrading bacteria symbiotic with mussels (C. gigas, C. asperella and B. tenella) and surrounding environments. Then we have compared quantity of microbial communities in all samples. The enumeration result of crude oil degrading bacteria and heterotrophic bacteria in this research confirmed that the quantity of bacteria was higher in the mussel samples compared to the surrounding environment. Some researchers also report similar results. Kueh and Chan (1985) suggested that differences exist between microbial communities of the surrounding seawater and mussels (C. gigas). In other study, Cavallo et al. (2009) reported that heterotrophic bacterial densities and diversity were higher in the mussel samples (Mytilus galloprovincialis) than in the surrounding seawater. In fact, mussel samples accumulated bacteria by filter feeding. Also, Silverman et al. (1996) report filtration activity by zebra mussel, this ability causes a change in the microbial communities and biodiversity of marine bacteria. The biodiversity of bacterial strain that isolated in this study was less compared to other researchers that work in the Persian Gulf (Radwan et al., 2005). This low diversity of crude oil degrading bacteria in this research can be interpreted as our sampling zone was limited, also the sampling sites was chronically polluted with crude oil and the level of crude oil pollution in the sampling sites was very high. It is possible that high crude oil contamination will result in the disappearance of bacteria in the collected mussel, seawater and sediment samples or crude oil contamination will cause decrease in the diversity of bacteria. The ability of the isolated strains to degrade crude oil was measured in this research. The results confirmed that the best strain for crude oil degradation was S. haliotis isolate BHA35 (69.17%). For the first time crude oil degrading bacteria of the genus Shewanella were isolated from mussels. The scale of crude oil degradation by microbial population depends on physical and chemical factors. The results of this research propose optimum condition for crude oil degradation as: 2.5% concentration of crude oil, 2–4% (w/v) concentration of NaCl, 30 days incubation time and mixed bacterial consortia. Fusey and Oudot (1984) reported that when the concentrations of crude oil were increased hydrocarbon biodegradation decreased. This pattern can be related to oxygen or nutrients limitation in higher concentrations of crude oil. Hasanshahian and Emtiazi (2008) demonstrated that maximum crude oil removal levels have been carried out at 1% concentration. Zahed et al. (2010) reported that most of the crude oil removal occurred at 100 mg/L and lower concentrations of crude oil are more efficient for crude oil removal. In fact with increasing concentration of crude oil and NaCl the hydrocarbon biodegradation decreased (Margesin and Schinner, 2001). This study also agrees with research of Kumar et al. (2007) which concluded that a negative relationship exists between salinity and hydrocarbon biodegradation. Mnif et al. (2009) observed that strain Halomonas sp. C2SS100 is able to degrade crude oil in the presence of 100 g/l NaCl. In this study the isolated strains tolerate high concentrations of salt (12% w/v) thus suitable for bioremediation of oil pollution in marine environments with high salinity such as Persian
Z. Bayat et al. / Marine Pollution Bulletin 101 (2015) 85–91
Gulf. Sathishkumar et al. (2008) suggested that crude oil degradation by microbial consortia is more effective in comparison to individual bacteria. They interpreted this result as in the microbial consortia there were different enzymes for various compounds of crude oil then the degradation was faster and has high performance. 5. Conclusions The results of this study confirmed the presence of crude oil degrading bacteria with mussels that collected from the Persian Gulf. These crude oil degrading bacteria have high biodegradation activity and have high tolerance to NaCl and crude oil. In addition, higher incubation time, mixed bacterial consortia and emulsifier production by strains improved the efficiency of crude oil degradation. Acknowledgment This work was supported financially by Shahid Bahonar University of Kerman. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.marpolbul.2015.11.021. References Al-Awadhi, H., Al-Hasan, R.H., Sorkhoh, N.A., Radwan, S.S., 2003. Establishing oildegrading biofilms on gravel particles and glass plates. Int. Biodeterior. Biodegrad. 51 (3), 181–189. Al-Mailem, D.A., Sorkhoh, N.A., Salamah, S., Eliyas, M., Radwan, S.S., 2010. Oilbioremediation potential of Arabian Gulf mud flats rich in diazotrophic hydrocarbon-utilizing bacteria. Int. Biodeterior. Biodegrad. 64 (3), 218–225. Cappello, S., Genovese, M., Della Torre, C., Crisari, A., Hassanshahian, M., Santisi, S., Calogero, R., Yakimov, M.M., 2012a. Effect of bioemulsificant exopolysaccharide EPS2003 on microbial community dynamics during assays of oil spill bioremediation: a microcosm study. Mar. Pollut. Bull. 64 (12), 2820–2828. Cappello, S., Russo, D., Santisi, S., Calogero, R., 2012b. Presence of hydrocarbon-degrading bacteria in the gills of mussel Mytilus galloprovincialis in a contaminated environment: a mesoscale simulation study. Chem. Ecol. 28 (3), 239–252. Cavallo, R.A., Acquaviva, M.I., Stabili, L., 2009. Culturable heterotrophic bacteria in sea water and Mytilus galloprovincialis from a Mediterranean area (Northern Ionian Sea — Italy). Environ. Monit. Assess. 149, 465–475. Dave, D., Ghaly, A.E., 2011. Remediation technologies for marine oil spills: a critical review and comparative analysis. Am. J. Environ. Sci. 7 (5), 423–440. Fuchsluger, C., Preims, M., Fritz, I., 2010. Automated measurement and quantification of heterotrophic bacteria in water samples based on the MPN method. J. Ind. Microbiol. Biotechnol. 38, 241–247. Fusey, P., Oudot, J., 1984. Relative influence of physical removal and biodegradation in the depuration of petroleum-contaminated seashore sediments. Mar. Pollut. Bull. 15, 136–141. Ghanavati, H., Emtiazi, G., Hassanshahian, M., 2008. Synergism effects of phenol degrading yeast and ammonia oxidizing bacteria for nitrification in coke wastewater of Esfahan steel company. Waste. Manage. Res. 26 (2), 203–208. Hasanshahian, M., Emtiazi, G., 2008. Investigation of alkane biodegradation using the microtiter plate method and correlation between biofilm formation, biosurfactant production and crude oil biodegradation. Int. Biodeterior. Biodegrad. 62, 170–178. Hassanshahian, M., Emtiazi, G., Kermanshahi, R., Cappello, S., 2010. Comparison of oil degrading microbial communities in sediments from the Persian Gulf and Caspian Sea. Soil. Sediment. Contaminat. 19 (3), 277–291. Hassanshahian, M., Emtiazi, G., Cappello, S., 2012a. Isolation and characterization of crude-oil-degrading bacteria from the Persian Gulf and the Caspian Sea. Mar. Pollut. Bull. 64, 7–12. Hassanshahian, M., Tebyanian, H., Cappello, S., 2012b. Isolation and characterization of two crude-oil degrading yeast strains, Yarrowia lipolytica PG-20 and PG-32 from Persian Gulf. Mar. Pollut. Bull. 64, 1386–1391. Hassanshahian, M., Ahmadinejad, M., Tebyanian, H., Kariminik, A., 2013. Isolation and characterization of alkane degrading bacteria from petroleum reservoir waste water in Iran (Kerman and Tehran provenances). Mar. Pollut. Bull. 73, 300–305.
91
Hassanshahian, M., Zeynalipour, M.S., Hosseinzadeh Musa, F., 2014a. Isolation and characterization of crude oil degrading bacteria from the Persian Gulf (Khorramshahr provenance). Mar. Pollut. Bull. 82, 39–44. Hassanshahian, M., Emtiazi, G., Caruso, G., Cappello, S., 2014b. Bioremediation (bioaugmentation/biostimulation) trials of oil polluted seawater: a mesocosm simulation study. Mar. Environ. Res. 95, 28–38. Hassanshahian, M., Yakimov, M.M., Denaro, R., Genovese, M., Cappello, S., 2014c. Using Real-Time PCR to assess changes in the crude oil degrading microbial community in contaminated seawater mesocosms. Inte. Biodeterior. Biodegrad. 93, 241–248. Hassanshahian, M., 2014d. Isolation and characterization of biosurfactant producing bacteria from Persian Gulf (Bushehr provenance). Mar. Pollut. Bull. 86, 361–366. Holt, S.G., Kriey, N.R., Sneath, P.H.A., Staley, J.T., Williams, S.T., 1998. Bergy's Manual of Determinative for Bacteriology. Williams and Wilkins, New York. Huber, M., 2010. Compendium of Bivalves, A Full-Color Guide to 3300 of the World's Marine Bivalves, a Status on Bivalvia After 250 Years of Research. Conch Books, Hackenheim, Germany (904 pp.). Kaczorek, E., Chrzanowski, L., Pijanowska, A., Olszanowski, A., 2008. Yeast and bacteria cell hydrophobicity and hydrocarbon biodegradation in the presence of natural surfactants: rhamnolipides and saponins. Bioresour. Technol. 99 (10), 4285–4291. Khan, K., Naeem, M., Asif, M., 2006. Extraction and characterization of oil degrading bacteria. J. Appl. Sci. 6 (10), 2302–2306. Kueh, C.S., Chan, K.Y., 1985. Bacteria in bivalve shellfish with special reference to the oyster. J. Appl. Bacteriol. 59, 41–47. Kumar, M., Leon, V., Manterano, A.D.S., Ilzins, O.A., 2007. A halotolerant and thermotolerant Bacillus sp. degrades hydrocarbons and produces tensio-active emulsifying agent. World J. Microbiol. Biotechnol. 23, 211–220. Manganaro, A., Pulicano, G., Reale, A., Sanfilippo, M., Sara, G., 2009. Filtration pressure by bivalves affects the trophic conditions in Mediterranean shallow ecosystems. Chem. Ecol. 25, 467–478. Margesin, R., Schinner, F., 2001. Bioremediation (natural attenuation and biostimulation) of diesel-oil-contaminated soil in an alpine glacier skiing area. Appl. Environ. Microbiol. 67, 3127–3133. Mnif, S., Chamkha, M., Sayadi, S., 2009. Isolation and characterization of Halomonas sp. Strain C2SS100, a hydrocarbon-degrading bacterium under hypersaline conditions. J. Appl. Microbiol. 107, 785–794. Pruthi, V., Cameotra, S.S., 1997. Rapid identification of biosurfactant-producing bacterial strains using a cell surface hydrophobicity technique. Biotechnol. Tech. 11, 671–674. Radwan, S., Al-Hasan, R.H., Al-Dabbous, S., Salamah, S., 2002. Bioremediation of oily sea water by bacteria immobilized in biofilms coating macroalgae. Int. Biodeterior. Biodegrad. 50 (1), 55–59. Radwan, S.S., Al-Hasan, R.H., Salamah, A., Khanafer, M., 2005. Oil-consuming microbial consortia floating in the Arabian Gulf. Int. Biodeterior. Biodegrad. 56, 28–33. Rahman, K.S.M., Thahira-Rahman, J., Lakshmanaperumalsamy, P., Banat, I.M., 2004. Towards efficient crude oil degradation by a mixed bacterial consortium. Bioresour. Technol. 85, 257–261. Sathishkumar, M., Binupriya, A., Baik, S.H., Yun, R.E., 2008. Biodegradation of crude oil by individual bacterial strains and a mixed bacterial consortium isolated from hydrocarbon contaminated areas. Clean 36 (1), 92–96. Saxena, R., Gothawal, R., Khan, M.N., 2013. Prospective in-silco approach in bioremediation of petroleum hydrocarbon: success so far. Octa J. Environ. Res. 1 (2), 137–149. Silverman, H., Lynn, J.W., Achebergee, C.R., Dietz, T.H., 1996. Gill structure in zebra mussels: bacterial-sized particle filtration. Am. Zool. 36, 373–384. Sorkhoh, N.A., Ghannoum, M.A., Ibrahim, A.S., Stretton, R.J., Radwan, S.S., 1990. Crude oil and hydrocarbon-degrading strains of Rhodococcus rhodochrous isolated from soil and marine environments in Kuwait. Environ. Pollut. 65 (1), 1–17. Stabili, L., Acquaviva, M.I., Cavallo, R.A., 2005. Mytilus galloprovincialis filter feeding on the bacterial community in a Mediterranean coastal area (Northern Ionian Sea, Italy). Water Res. 39, 469–477. Tebyanian, H., Hassanshahian, M., Kariminik, A., 2013. hexadecane-degradation by Teskumurella and Stenotrophomonas strains iso lated from hydrocarbon contaminated soils. Jundishapur. J. Microbiol. 26 (7), e9182. Thomas, J.C., Wafula, D., Chauhan, A., Green, S.J., Gragg, R., Jagoe, C., 2014. A survey of deepwater horizon (DWH) oil-degrading bacteria from the Eastern oyster biome and its surrounding environment. Front. Microbiol. 5, 149. Todd, P.A., Ong, X., Chou, L.M., 2010. Impacts of pollution on marine life in Southeast Asia. Biodivers. Conserv. 19 (4), 1063–1082. Udeh, N.U., Nwaogazie, I.L., Momoh, Y., 2013. Bioremediation of a crude oil contaminated soil using water hyacinth (Eichhornia crassipes). Adv. Appl. Sci. Res. 4 (2), 362–369. Yakimov, M.M., Timmis, K.N., Golyshin, P.N., 2007. Obligate oil-degrading marine bacteria. Curr. Opin. Biotechnol. 18 (3), 257–266. Zahed, M.A., Hamidi, A.A., Isa, M.H., Mohajeri, L., 2010. Effect of initial oil concentration and dispersant on crude oil biodegradation in contaminated seawater. Bull. Environ. Contam. Toxicol. 84, 438–442. Zhou, Y., Zhang, S., Liu, Y., Yang, H., 2014. Biologically induced deposition of fine suspended particles by filter-feeding bivalves in land-based industrial marine aquaculture wastewater. PLoS One 9 (9).