Population dynamics, isolation and characterization of potential engine oil degrading indigenous bacteria in contaminated sites in Taif, KSA.
The elimination of a wide range of pollutants and wastes from the environment is an absolute requirement to promote a sustainable development of our society with low environmental impact. Thanks to their metabolic potential and wide biodiversity micro-organisms play a fundamental role in making biodegradation processes more efficient, and in transforming recalcitrant macromolecules into chemical substances more amenable to further metabolization. Oil pollution is a great hazard to soil and aquatic environments. Crude oil is a complex mixture of many compounds such as alkanes, aromatics, resins and asphaltenes.
To remedy these problems, millions of dollars is spent annually around the globe. Currently, the main prevailing method of coping with the problem is the use of special chemicals (Jorda, 1966). Nevertheless, starting from the early 80's, application of the microbial methods to fight the deposit formations have found grounds and improved by time. The progress in this field is such that in some occasions not only they may compete with the chemical methods, but also they may be the only choice due to the technical and/or economical reasons (Partidas et al., 1999).
Bioremediation is the application of microbial processes to convert environmental contaminants into harmless substances (for instance, C[H.sub.4], C[O.sub.2], [H.sub.2]O). It has emerged as a good technique for environmental treatment regarding organic compounds, such as petroleum hydrocarbons, due to its flexibility and adaptability in different sites (Ryan et al., 1991). The advantages of the study of bioremediation in crude oil contaminated soil treatment is its cost-effectiveness when compared to some physicochemical techniques.
Paraffinic hydrocarbons are organic compounds commonly found in crude oil, consisting of various forms and combinations of aliphatic hydrocarbons, aromatic hydrocarbons, napthenes, and asphaltenes. Environmental pollution with petroleum and petroleum products (complex mixture of hydrocarbons) has been recognized as one of the most serious current problems especially when associated with accidental spills on large-scale. It is used to lubricate the parts of automobile engine in order to keep everything running smoothly (Hagwell et al., 1992). Used oil was defined by the US Environmental Protection Agency (40CFR Pan 270) as oil that has been refined from crude oil or any synthetic oil; this has been used and as a result of such use is contaminated by chemical impurities which contribute to chronic hazards including mutagencity and carcinogenicity as well as environmental hazard with global ramifications (Blodgette, 2001).
Bioremediation has become an alternative way to remedy oil polluted sites, where the addition of specific microorganism (bacteria, cyanobacteria, algae, fungi, protozoa) or enhancement of microorganism already present, can improve biodegradation efficiency (Hagwell et al., 1992). These microorganisms can degrade a wide range of target constituents present in oil sludge (Barathi and Vanudevan, 2001; Mishra et al., 2001). A large number of Pseudomonas strains capable of degrading polycyclic aromatic hydrocarbons have been isolated from soil (Johnson et al., 1996; Kiyohara et al., 1994). Other petroleum hydrocarbon degraders include Yokenella spp., Alcaligenes spp., Roseomanas spp., Sreanotrophomanas spp., Acinetobacter spp., Flavobacter spp., cyanobacterium spp., capnocytophage spp., Moraxella spp. and Bacillus spp. (Antai, 1990). Other microorganism such as fungi is also capable of degrading the hydrocarbons in engine oil to a certain extent. However, they take longer time (Udeani et al., 2009).
Whatever the mechanisms involved, a point which is still only partially understood at the present time is the kinetics of long-chain alkane biodegradation. Studies focused on the kinetics of growth of yeasts on n-alkanes were conducted in the seventies and models were proposed to account for the complex kinetic patterns observed (Gutierrez and Erickson, 1977; Verkooyen and Rietma, 1980).
The vast range of substrates and metabolites present in hydrocarbon impacted soils surely provides an environment for the development of a quite complex microbial community (Butier and Mason, 1997). Microbial populations that consist of strains that belong to various genera have been detected in petroleum-contaminated soil or water (Sorkhoh et al., 1995, Chikere et al., 2009). This strongly suggests that each strain or genera have their roles in the hydrocarbon transformation processes. More recently, microbial remediation was found to be an available alternative method over the conventional methods. Microbial treatment can control contamination of soils with used or fresh petroleum products by reducing the length of the paraffin and oil molecules and by producing by-products that act as surfactants and paraffin and oil solvents. However, information on numbers and local species of microorganisms as well as their efficiency in degradation in Saudi Arabia is scarce.
In this paper, it is attempted first to survey total indigenous bacteria and fungi present in contaminated sites and to isolate and identify bacteria capable of degradation of used petroleum oil residuals. The capability of isolated bacteria to degrade mineral oil as a representative of hydrocarbon residuals was evaluated. In addition, morphological, physiological, biochemical characteristics tests and Api kit profiling were employed for the identification of the petroleum oil biodegrading-bacterial strains.
Materials and Methods
Locations were different gas stations and mechanic workshops in Taif area, KSA at altitudes ranging from 1500 to 2,000 m above sea level. The places were industrial areas in Taif state.
Soil sampling and processing
Samples were collected from multiple areas within the gas station or mechanic workshops that had heavy spillage of fresh or used engine oil, benzene, diesel and lubrication oil and mixed to produce composite samples. The locations had no grasses growing on them and soil samples were collected at three different locations at each workshop. One sample of 100g from each location was collected during Fall 2009 and Spring 2010. Samples were taken down to 10 cm depth, after discarding the upper 3 cm of the soil surface. Each soil sample was crushed, thoroughly mixed then sieved through a 2 mm pore size sieve to get rid of large debris. The sieved soil was then used for the enumeration and isolation purposes. Samples were placed in polyethylene bags. Closed tightly and stored at 4[+ or -]1[degrees]C. The soils were characterized by hardened surfaces and blackish in color. Moisture content and pH were measured.
pH and moisture content determination
Moisture contents were estimated by weighting 10.0g contaminated soil and put it in oven at 105[degrees]C to a constant weight from the following equation: % of moisture content = B - C/ C - A x 100 where A= weight of empty can, B= weight of can and sample before drying and C= weight of can and sample after drying.
The pH is the negative logarithm of hydrogen ion in one litter where a solution of 1: 2.5 of sample and water after calibration of the pH-meter with pH buffer 4 and 7 for reading.
Indigenous microorganism's enumeration
Samples were enumerated by making tenfold serial dilution of the soil samples using physiological saline. From the diluted sample, using a dropper pipette, 1 ml of each dilution was dropped onto Petri dish then the plate count agar and Sabouraud media were poured. Duplicates of plates were used for each dilution. Plates were incubated for 48-72 h at 30[degrees]C in an incubator. Each inoculum of microorganism developed into a discreet colony. All plates yielding 30-300 colonies were counted. The number of viable microorganisms in the sample was calculated from the number of colonies formed, the volume of inoculum used by dropper pipette and the dilution factor expressed in colony forming unit (CFU) (Krieg, 1984).
Isolation of bacteria
The microorganisms used in this study were obtained by the enrichment culture technique from contaminated soils which had been contaminated with hydrocarbon for a long time at mechanic workshop and gas stations. Samples were obtained from different locations at Taif area. The culture media used were Bushnell-Haas (1941) mineral media which is an enrichment medium for isolation of bacterial degrading organism and nutrient agar. Bushnell-Haas broth medium containing (g/L): 0.2 g MgS[O.sub.4].7[H.sub.2]O; 1.0 g [K.sub.2]HP[O.sub.4]; 1.0 g K[H.sub.2]P[O.sub.4]; 0.05 g Fe[Cl.sub.3]; 1.0 g N[H.sub.4]N[O.sub.3]; 0.02 g Ca[Cl.sub.2]; pH to 7-7.2 and sterilize at 121C[degrees] for 15 min. Inoculation was done using Bushnell-Haas enrichment medium of 10 ml in 250 mL flasks into which ten grams of the contaminated soil was added and incubated at 30[degrees]C for 4 weeks. Samples that were turbid were sub-cultured into nutrient agar using LB broth, as diluents to observe the morphological characteristics of the isolates. Colonies showing a good growth and characters were picked and streaked on new Bushnell-Haas minimal agar plates. A rapidly growing, visually distinct colony and a separate, morphologically unique isolate were selected for further analysis and purified by repeated plating.
Bacterial identification and tests
The morphological characteristics of the isolates were identified by gram stain and biochemical reactions. The biochemical reactions include glucose fermentation, oxidase test, catalase production reaction, cell motility; egg yolk reaction and reaction in tryptose soya broth were performed. The isolates were purified, identified and named based on morphological, physiological and biochemical characteristics presented in Bergey's Manual of Determinative Bacteriology (Krieg and Holt, 1994) and the APi Kit profiling (bioMerieux, France, 2009).
Culture growth conditions
Medium used was Luria-Bertani (LB) broth (Trypton, 10g; yeast extract, 5g; NaCl, 5g; distilled water, 1000 ml) to grow mother cultures in 125 ml Erlenmeyer flasks with 20 ml of medium and incubated with shaking (150 rpm) at 37[degrees]C overnight. The mother cultures of the isolates were cultivated in a minimum salt medium containing 1 mL of mineral oil as a sole carbon and energy source. Cultures were cultivated in 250 ml Erlenmeyer flasks with 100 ml of medium and incubated with shaking (150 rpm) at 37[degrees]C.
Optical density and biomass measurement
The turbidity of the cultures was determined by measuring the Optical Density (OD) at a wavelength of 595 nm in 2 ml cuvettes using a spectrophotometer (Biophotometer plus, Eppendorf). The net dry weight for the biomass was determined simultaneously. A 1 mL of culture was centrifuged at 1500 rpm for 10 min, washed twice with distilled water, poured into a pre-weighed container, dried overnight at 90 [degrees]C to constant weight and cooled for reweighing. The linear relation between [OD.sub.595] and dry mass was obtained.
Mineral oil concentrations and bacterial growth
Growth of the isolated bacterial strains on different concentrations of mineral oil 1, 3, 5 % (v/v) was evaluated by measuring culture optical density (OD) at 595 nm.
Growth rate measurement
The growth rates of cultures in exponential phase were determined from linear regressions of log10 absorbency vs. time, calculating a least squares fit of data from the exponential growth phase, and determining the slope of this line. The instantaneous growth rate ([micro]) was determined from the slope of this line x ln10; [micro] had the dimensions /h (Koch, 1984).
Gas chromatograph measurements
The samples were measured at the Regional Center for Mycology and Biotechnology (RCMB), Al-Azhar University, Cairo, Egypt. The samples were extracted by adding 100 [micro]l of hexane and gentle mixing by continuous inversion for 10 min. The top phase containing the organic solvent with the extracted oil was removed with a Pasteur pipette and transferred to a new set of tubes, sealed and stored at -20[degrees]C until analysis. The analyses were carried out in triplicate in a TechComp D-7900 ver. 1.30 Gas Chromatograph equipped with a flame ionization detector using a 30 m x 0.32 mm, 0.25 [micro]m internal diameter, polar DB1 fused silica capillary column (Supelco). The carrier gas was nitrogen at a flow rate of 20 ml/min. The temperature of the injector was 230[degrees]C and that of the flame ionization detector was maintained at 320[degrees]C. The oven temperature after sample injection (2[micro]l) was 1 min at 50[degrees]C, increasing to 200[degrees]C at 15[degrees]C/min and held at this temperature for 2 min then raised to 280[degrees]C with an increasing rate of 15[degrees]C/min and held for 2 min.
Results and Discussion
After collection of soil samples, they were placed in polyethylene bags and the moisture content and pH degree of each sample were determined (Table 1).
Soil moisture content ranged between 0.2 and 4.6 %, and pH ranged between 2.3 and 7.6. Soil moisture percentage and pH degree affect numbers and types of bacteria. The optimum pH for biodegradation of hydrocarbons is around pH 6-8 (Mentzer and Ebere, 1996). Biodegradation of crude petroleum in acid soil (pH 4.5) could be doubled by liming to pH 7.4. Soil that is hydrated with 50-80 % of the maximum water-holding capacity has the greatest microbial activity (Mentzer and Ebere, 1996). Below that level, osmotic and matrix forces limit the availability of water to microbes; above that level, the reduction of air space and oxygen decrease microbial activity.
Total viable count
Total viable bacteria and fungi count was determined (Table, 2). Total bacteria number ranged between 2.1x[10.sup.2] - 4.7x[10.sup.7] CFU/g dry soil. Total fungi count ranged between 0.3x[10.sup.2] - 5.9x[10.sup.3] CFU/g dray soil. Ten compound soil samples were collected from 10 different mechanic workshops and gas stations. The results show a prevalence rate of 10.0% yield of bacteria degrading agents.
Potential hydrocarbon degrader bacteria were isolated from soil samples that have been exposed to petroleum oil spills in mechanic workshops and gas stations. Saadoun (2002) reported that bacterial population of polluted soils showed counts ranging between 9.5 x [10.sup.5] and 237.5 x [10.sup.5] CFU/g soil with 2 different colony types of bacterial strains which have been recovered on the agar plates. Results indicated that longer aged contamination exhibited a greater number of microorganisms. Udeani et al. (2008) evaluated the bacterial diversity of soil environment contaminated with used engine oil of mechanic workshops in Nigeria and found it had bacteria densities of 1.25 x [10.sup.4] to 6.25 x [10.sup.5] from the soil samples collected from each site.
Phenotypic examination of the recovered bacteria isolate revealed that they belong mainly to the genera of Enterobacter, Acinetobacter, Bacillus, Corynebacterium, Micrococcus, Serretia, Pseudomonas, Erwinia, Actinomyces sp. and Saccharomyces. Isolates were identified on the basis of their cultural and biochemical characteristics according to Bergey's Manual of Determinative Bacteriology (9th edition) as well as APi kit profiling. Six isolates were selected to examine their growth rate and biomass on mineral oil and physical appearances were used as indication for the ability of these isolates to grow on mineral oil. Four isolates were selected to examine their potential in degradation of mineral oil 1 % (v/v), and rate of degradation was determined. The isolates were named as AF1-GT, AF8-GT, AF10-MT, AF11-GT, AF104-MH, and AF203-MH. Based on their morphological, biochemical characterization isolates were identified as the following: AF1-GT1 to be Erwinia species, AF8-GT and AF10-MT to be Bacillus species, AF11-GT to be Pseudomonas aerogenosa, AF203-MH to be Saccharomyces cerevieace and AF104-MH to be filamentous type of Actinomyces species.
Saadoun (2002) isolated Pseudomonas, Enterobacter and Acinetobacter from contaminated soils. Udeani et al. (2008) showed the isolation of Bacillus Stearothermophilus (8.3%) and Cyanobacteria (1.7%) from mechanic workshops at the sites sampled. The number of viable bacterial growth of B. Stearothermophilus and Cyanobacteria were enumerated and expressed in colony forming units (CFU).
Lazar et al. (1999) stated that microorganisms involved with the microbial treatment of crude oil, are generally live, naturally occurring, and are mainly facultative anaerobic, non-pathogenic, contain no sulphate-reducing bacteria or slime-forming bacteria and are environmentally safe. All our bacterial strains, however, catalase positive and were aerobes except for AF8-GT and AF10-MT which were facultative anaerobe.
[FIGURE 1 OMITTED]
Saadoun (2002) isolated Pseudomonas, Enterobacter and Acinetobacter from contaminated soils. Turbidity, dry weight and physical appearance were used as an indication for the ability of these bacteria to grow on diesel. Action of three different Pseudomonas species, Acinetobacter lowffi, Enterobacter cloacae and Rhodococcus erythropol on 0.1 % (v/v) diesel was followed at 1, 2, 6 and 12 h. Pseudomonas putida and P. mallei and Enterobacter cloacae indicated a positive reaction; however, Pseudomonas maltophilia and Acinetobacter lowffi showed no effect. Udeani et al. (2008) showed the isolation of Bacillus Stearothermophilus (8.3%) and Cyanobacteria (1.7%) from mechanic workshops at the sites sampled. The number of viable bacterial growth of B. Stearothermophilus and Cyanobacteria were enumerated and expressed in colony forming units (CFU).
Lazar et al. (1999), Pokethitiyook et al. (2002), and Sadeghazad and Ghaemi (2003) reported that, Pseudomonas and Bacilli species were the most effective microorganisms in the biodegradation of heavy hydrocarbons. Lazar et al. (1999) also, stated that microorganisms involved with the microbial treatment of crude oil, are generally live, naturally occurring, and are mainly facultative anaerobic, nonpathogenic, contain no sulphate-reducing bacteria or slime-forming bacteria and are environmentally safe. All our bacterial strains, however, catalase positive and were aerobes except for AF8-GT and AF10-MT which were facultative anaerobe.
The specific growth rates of the different isolates on mineral oil (Fig. 2 and Fig. 3) showed AF8-GT and AF10-MT to be the fastest growing in mineral salts medium containing 1.0 % (v/v) mineral oil. Most growth occurred in the initial 12 h for isolate AF10-MT resulting in approximately 0.001 g/ml (Fig. 2) biomass production. AF8-GT has a lag phase of approximately 6 h and peak growth at 10 h reaching 0.001 g/ml (optical density of 1 at 595 nm) (Fig. 3). Maximum specific growth rates ([[micro].sub.max]) for AF8-GT and AF10-MT were 0.065[h.sup.-1] and 0.055 [h.sup.-1], respectively (Table 4).
Individual microorganisms can metabolize only a limited range of hydrocarbon substrates, so assemblages of mixed populations with overall broad enzymatic capacities are required to bring the rate and extent of petroleum biodegradation further (Ghazali et. al. 2004). Microbial populations that consist of strains that belong to various genera have been detected in petroleum-contaminated soil or water (Sorkhoh et al., 1995). This strongly suggests that each strain or genera have their roles in the hydrocarbon transformation processes.
The vast range of substrates and metabolites present in hydrocarbon impacted soils surely provides an environment for the development of a quite complex microbial community (Butier and Mason, 1997). It is used to lubricate the parts of automobile engine in order to keep everything running smoothly (Hagwell et al., 1992).
In motor mechanics workshops there is a constant change in the soil microorganism as a result of deliberate spillage of used engine oil. These alter the biomass and ecology of the soil such that both microbial communities and grasses can no longer grow on the soil spots. The color and texture of the soil are affected; this leads to different microbial flora establishment in an attempt to remedy the petroleum product spillage (Bartha and Atlas, 1977). The total heterotrophic bacteria count range from 1.25 x 104 - 6.25 x 105 in this work. Butier and Mason (1997) indicated that there is an increase in heterotrophic bacteria population in the presence of dispersant agent. Also, Antai (1990) reported two major response to crude oil in which there is an increase in biomass. Although, this disagrees with the work done by Lizarraga-Partida et al. (1982) who observed that petroleum hydrocarbon has little or no effect on the total number of heterotrophic bacteria.
Biomass and optical density
Optical density and biomass were determined simultaneously. The linear relation between O[D.sub.595] and dry mass was obtained during growth on 1% of mineral oil as shown in Fig. 2-7.
The specific growth rate and net dry weight of the different isolates were determined and illustrated in figures 2-7. These figures indicate the effluence of the specific growth rate and biomass precipitation on a period of bacterial cultivation in mineral salts medium containing 1 % (v/v) mineral oil. The highest specific growth rate was observed on isolate AF1-GT, AF11-MT with [[micro].sub.max] (0.110[h.sup.-1]) and (0.911 [h.sup.-1]), respectively. Four isolates AF1-GT, AF8-GT, AF10-MT, AF11-GT were later selected for further investigations, isolate AF8-GT, and AF10-MT have faster growth rate over the other isolates (Table 4).
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Degradation of mineral oil by the bacterial isolates
The standard cultures of the isolates AF1-GT (Erwinia sp.), AF8-GT (Bacillus sp.), AF10-MT (Bacillus subtilis), AF11-GT(Pseudomonas aergenosa) were examined for their capability to degrade mineral oil as sole carbon source and energy in mineral media for 24 hr. All isolates exhibited high efficiency of assimilating the mineral oil. Isolate AF11-GT
Isolation of alkane degrading microorganisms from oil contaminated soil has been reported by several researchers. Nazina et al. (2005) have obtained hydrocarbon oxidizing Geobacilli strains from formation waters of oil fields. Hydrocarbon degrading members of the family Bacillaceae were found to dominate the oil contaminated soil of Kuwait (Mohamed et al., 2006). Moreover, a thermophilic bacterial strain G. thermodenitrificans that shows selective degradation of long chain alkanes, similar to the degradation pattern of isolate AF11-GT, was also isolated (Wang et al., 2006).
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The effects of mineral oil concentration as a sole carbon source on different isolates were studied in Erylenmyer flasks 250 ml with different concentration 1, 3 and 5 % (v/v). Strain AF11-GT (Pseudomonas aerogenosa) showed good growth on three different concentrations of mineral oil. However, highest growth was observed on 3% mineral oil followed by 1% and 5% concentrations. This data suggests that the lower the concentration of mineral oil the higher was the utilization.
Etoumi (2007) observed a reduction in wax appearance temperature and heavy hydrocarbon fractions by biodegradation of paraffinic hydrocarbons using Pseudomonas and Actinomyces species. It was reported that the lower the concentration of hydrocarbons the higher was the utilization.
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In conclusion, the results obtained from the present study could help in understanding the biodegradation of mineral oil in contaminated sites, as well as to design efficient biocatalyst allowing transformation of oil fractions into valuable compounds. The isolation of pure strains from such a consortium has also been achieved, their mineral oil degradation ability was confirmed, and different effects of mineral oil on their degrading capacity have been shown. Preliminary identification of isolated strains has been achieved and further work continues on their molecular characterization.
More research is required to understand the fundamental mechanisms of enhancement and inhibition of the microbial degradation of high concentration of toxic compounds. However, these microorganisms could be used very effectively for in situ bioremediation in an environment which is highly contaminated with oil products. However, further research could be carried out on genetic manipulation of bacterial isolates for improvement and exploitation as bioremediation vehicles.
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Shahaby A.F. and A.E. EL Tarras
Biotechnology and Genetic Engineering Research Center (BGERC), College of Medicine, Taif University, KSA
Table (1): The soil moisture percentage and pH degrees. Sample Moisture % pH 1 0.2 7.33 2 2.2 7.44 3 0.6 7.40 4 1.4 2.3 5 0.4 6.5 5 0.4 7.3 7 1.0 7.3 8 4.6 6.8 9 1.4 7.6 10 0.6 7.45 Table (2): Total bacteria and Fungi count in industrial area at Taif, KSA. Sample Total fungal Total Bacterial count Count 1 1.1 x [10.sup.2] 2.9 x [10.sup.6] 2 4.3 x [10.sup.2] 3.5 x [10.sup.5] 3 0.3 x [10.sup.2] 2.7 x [10.sup.4] 4 ND 2.1 x [10.sup.2] 5 5.9 x [10.sup.3] 8.8 x [10.sup.6] 6 3.1 x [10.sup.2] 4.7 x [10.sup.6] 7 2.3 x [10.sup.2] 4.7 x [10.sup.7] 8 3.3 x [10.sup.3] 6.3 x [10.sup.5] 9 1.5 x [10.sup.2] 5.5 x [10.sup.4] 10 ND 3.3 x [10.sup.6] ND = Not determined Table (3): Morphological and biochemical characters of the obtained bacterial isolates. Proposed name Oxidase Catalase Motility Gram Erwinia sp. - + + - Bacillus sp. - + + + Bacillus + + + - subtilis Psuedomonas - + + + aerogenosa Actinomyces + + + + Saccharomyces + + - + cerevieace Proposed name Morphology Color Isolate Erwinia sp. Short rod Clear AF1-GT Bacillus sp. Long rod Creamy AF8-GT Spore Bacillus Long rod Heavy AF10-MT subtilis Spore Creamy Psuedomonas Short rod Clear AF11-GT aerogenosa greenish Actinomyces Long thin White AF104- Hyphe MH Saccharomyces Oval Heavy AF203- cerevieace Creamy MH Table 4: Growth rates of the selected isolates grown in minimal medium with mineral oil as carbon and energy source. Strains Growth rate ([micro]) Biomass yield (g cells/mL ([h.sup.-1]) mineral oil) AF1-GT 0.110 2.90 AF8-GT 0.065 2.74 AF10-MT 0.055 2.94 AF11-GT 0.911 2.96 AF104-MH 0.073 3.63 AF203-MH 0.064 3.55
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|Author:||A.F., Shahaby; Tarras, A.E. El|
|Publication:||International Journal of Petroleum Science and Technology|
|Date:||Jan 1, 2011|
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