Characterization of a Polycyclic Aromatic Hydrocarbon-Degrading Microbial Consortium From a Petrochemical Sludge Landfarming Site

March 31, 2007

By Jacques, Rodrigo J S; Okeke, Benedict C; Bento, Fatima M; Peralba, Maria C R; Camargo, Flvio A O


Anthracene, phenanthrene, and pyrene are polycyclic aromatic hydrocarbon (PAHs) that display both mutagenic and carcinogenic properties. They are recalcitrant to microbial degradation in soil and water due to their complex molecular structure and low solubility in water. This study presents the characterization of an efficient PAH (anthracene, phenanthrene, and pyrene)degrading microbial consortium, isolated from a petrochemical sludge landfarming site. Soil samples collected at the landfarming area were used as inoculum in Warburg flasks containing soil spiked with 250 mg kg^sup -1^ of anthracene. The soil sample with the highest production of CO^sub 2^-C in 176 days was used in liquid mineral medium for further enrichment of anthracene degraders. The microbial consortium degraded 48%, 67%, and 22% of the anthracene, phenanthrene, and pyrene in the mineral medium, respectively, after 30 days of incubation. Six bacteria, identified by 16S rRNA sequencing as Mycobacterium fortuitum, Bacillus cereus, Microbacterium sp., Gordonia polyisoprenivorans, two Microbacteriaceae bacteria, and a fungus identified as Fusarium oxysporum were isolated from the enrichment culture. The consortium and its monoculture isolates utilized a variety of hydrocarbons including PAHs (pyrene, anthracene, phenanthrene, and naftalene), monoaromatics hydrocarbons (benzene, ethylbenzene, toluene, and xylene), aliphatic hydrocarbons (1-decene, 1-octene, and hexane), hydrocarbon mixtures (gasoline and diesel oil), intermediary metabolites of PAHs degradation (catechol, gentisic acid, salicylic acid, and dihydroxybenzoic acid) and ethanol for growth. Biosurfactant production by the isolates was assessed by an emulsification index and reduction of the surface tension in the mineral medium. Significant emulsification was observed with the isolates, indicating production of high-molecular-weigh surfactants. The high PAH degradation rates, the wide spectrum of hydrocarbons utilization, and emulsification capacities of the microbial consortium and its member microbes indicate that they can be used for biotreatment and bioaugumentation of soils contaminated with PAHs.

KEYWORDS anthracene, biosurfactant, degradation, microbial consortium, phenanthrene, pyrene


Polycyclic aromatic hydrocarbons (PAHs) are a group of chemical compounds consisting of several atoms of carbon and hydrogen, arranged in the form of two or more aromatic rings. Due to the possibility of ring fusion at different positions, there are more than 100 PAHs that are recognized by IUPAC (International Union of Pure and Applied Chemistry). PAH exposure occurs by breathing, ingestion, and dermal absorption. PAHs are highly lipid soluble and quickly absorbed by the gastrointestinal tract of mammals. PAH metabolism in human body produces epoxides with mutagenic and carcinogenic properties and cases of lung, intestinal, liver, pancreas, and skin cancers have been reported (Samanta et al., 2002).

PAHs are natural or anthropogenic in origin, formed due to incomplete combustion of organic substances such as coal, oil, gas, wood, and garbage and from petrochemical industries and oil refining. The residues generated by these activities contain a variety of PAHs, such as anthracene, phenanthrene, and pyrene. Release of PAH residues into the environment can lead to the contamination of natural resources. Their complex molecular structure and low solubility in water limit the application of conventional remediation techniques (Boopathy, 2000).

Although some microorganisms can metabolize PAHs to CO^sub 2^ and water, most of the in situ remediation study results show a low rate of degradation. This has been attributed to the inability of the microbes to degrade the PAHs, low solubility, and contaminant, and nutrient limitations (Huesemann et al., 2002; Johnsen et al., 2005).

PAH degrading bacteria include Pseudomonas, Aeromonas, Beijerinckia, Flavobacterium, Nocardia, Corynebacterium, Sphingomonas, Mycobacterium, Stenotrophomonas, Paracoccus, Burkbolderia, among others (Cerniglia, 1984; Zhang et al., 2004). Fungal genera reported to degrade PAHs are Cunnighametta, Pbanerochaete, Cndida, Penicillium, Pleorotus, Trametes, Aspergillus, Bjerkandera, Chrysosporium, and other (Cerniglia, 1997). One important advantage of using microbial consortia is they possess multiple metabolic capacities that increase the efficiency of the bioremediation process (Boonchan et al., 2000; Ghazalli et al., 2004).

The bioavailability of a compound in the environment is determined by the rate of mass transfer of the substrate to the microbial cells in relation to intrinsic catabolic activity (Johnsen et al., 2005). Microbial degraders can increase the PAH bioavailability through secretion of biosurfactants: molecules that contain a hydrophobic region as well as a hydrophilic region. These molecules reduce the interfacial and superficial tension in the growth medium, and they also form stable emulsions. The emulsions increase the solubility of the PAHs and consequently promote the bioavailability of such chemicals in the environment (Cameotra and Bollag, 2003).

Because previous studies on PAHs bioremediation reported low rates, the objective of this study was to isolate and characterize high-efficiency PAH-degrading microbial consortium from a 16-year- old petrochemical landfarming site with history of PAH disposal. The second objective was to evaluate the capacity of the consortium and its monoculture isolates to utilize a variety of toxic aromatic and aliphatic hydrocarbons that are likely to be found in sites contaminated with complex mixtures of PAHs.


Soil Sampling

Soil samples were collected aseptically from a layer 0 to 30 cm deep at a 16-year-old landfarming site for an effluent treatment plant of petrochemical industries and an oil refinery. Twelve landfarming cells of 15,000 m^sup 2^ each were built in 1989 for the disposal, at different rates, of the plant sludge, containing a complex mixture of PAHs, including anthracene, phenathrene, and pyrene. Three cells that had recently received residues were selected and soil samples were collected in two locations chosen randomly inside each cell, resulting in six samples that were brought to the laboratory at 4C.

Respirometric Evaluation of Landfarming Soil Samples

The six soil samples were evaluated for heterotrophic activity by respirometric assay using an uncontaminated soil to which a PAH was added. The uncontaminated soil was a paleodult collected from an agricultural area that had not received any PAH. The composition of the soil was as follows: clay content of 90.0 g kg^sup -1^; pH (H^sub 2^O) of 4.7; P of 4.1 mg dm-3; K of 20.0 mg dm^sup -3^; SOM (Soil Organic Matter) of 9.0 g kg^sup -1^; exchangeable Al of 0.2 cmol^sub c^ dm^sup -3^; exchangeable Ca of 0.4 cmol^sub c^ dm^sup – 3^; exchangeable Mg of 0.2 cmol^sub c^ dm^sup -3^; S of 3.4 mg dm^sup -3^; Fe of 3.4 g dm^sup -3^, and CEC (Cation Exchange Capacity) of 2.2 cmol^sub c^ dm^sup -3^. Soil pH was adjusted to 6.5 using CaCO^sub 3^ and MgCO^sub 3^ (3:1). The soil (50 g in 1.5 L respirometric flask) was contaminated with anthracene using a stock solution in acetone. The flasks were kept in an incubation chamber for 3 h at 50C for acetone to evaporate. A further 50 g of soil was added and thoroughly mixed. A total of 50 mg kg^sup -1^ of N and 25 mg kg^sup -1^ of P using NH^sub 4^NO^sub 3^ and KH^sub 2^PO^sub 4^, respectively, were added to the soil sample due to the soil’s low fertility. In parallel experiments, 1 g of each landfarming soil sample was used to inoculate separate respirometric flask and the control flasks received 1 g of the noncontaminated soil. The flasks were incubated in triplicate at room temperature and CO^sub 2^ was trapped in a 0.25 M NaOH solution. During a period of 176 days, the CO^sub 2^ trap was assessed by adding 1 mL of 1 M BaCl^sub 2^ solution and then titrated with 0.5 M HCl using phenolphthalein as the indicator. CO^sub 2^-C was quantified using the formula of Stotzky (1965) as follows: CO^sub 2^-C (mg soil kg^sup -1^) = (B – T) eq M 10; where B is the volume (in mL) of the solution of HCl used to titrate the blank (respirometric flask without soil); T is the volume (in mL) of the solution of HCl used to titrate the treatments; eq is the equivalent weight, in this case 6; M is the molarity of the standardized solution of HCl; and 10 is the conversion factor for soil to kilogram.

PAH Degrader Enrichment

One gram of soil from the flask with the highest production of CO^sub 2^-C was added to 125 ml Erlenmeyer flask containing 50 ml of the Tanner Mineral Medium (TMM) and spiked with 250 mg L^sup -1^ anthracene as the only source of C and energy. In all tests in vitro the anthracene was added to mineral medium as fine crystals (Merck). TMM used was composed of (g L^sup -1^ of deionized water) 0.04 CaCl^sub 2^ 2H^sub 2^O; 0.1 KH^sub 2^PO^sub 4^; 0.8 NaCl; 1.0 NH^sub 4^Cl; 0.2 MgSO^sub 4^ 7H^sub 2^O; 0.1 KCl. Micronutrients used were composed of (mg L^sup -1\^ of deionized water) 0.1 CoCl^sub 2^ 6H^sub 2^O; 0.425 MnCl^sub 2^ 4H^sub 2^O; 0.05 ZnCl^sub 2^; 0.015 CuSO^sub 4^ 5H^sub 2^O; 0.01 NiCl^sub 2^ 6H^sub 2^O; 0.01 Na^sub 2^MoO^sub 4^ 2H^sub 2^O; 0.01 Na^sub 2^SeO^sub 4^ 2H^sub 2^O. The pH was adjusted to 7.0 by adding aliquots of either HCl or NaOH. The medium was sterilized by autoclaving at 121C for 20 min. Cultures were incubated with orbital shaking (150 rpm) at 30C. Each week, 1 ml of culture was transferred to another flask containing the same sterile mineral medium and incubated.

Isolation of PAH-Degrading Consortia

After three enrichment transfers, the culture was diluted in saline solution (NaCl 0.85%) and plated in agar medium (3 g of meat extract, 5 g of peptone, 15 g of agar in 1 L of distilled water, pH 7.0) containing 250 mg L^sup -1^ of anthracene (Kiyohara et al., 1982) and incubated for 72 h at 30C. Morphologically different discrete colonies were purified by repeated streaking on the same medium and stored at 4C. The fungal colony was purified on malt agar (30 g of malt extract, 5 g of peptone, 15 g of agar in 1 L of distilled water at pH 5.4) containing 250 mg L^sup -1^ of anthracene and stored at 4C.

Carbon Concentration in the Mineral Medium

Dissolved carbon in the TMM was assessed using a total organic carbon analyzer (TOC-V; CSH, Shimadzu, Japan) to determine the concentration of organic C dissolved in the distilled water (as negative control). TOC of the test mineral medium and control mineral medium to which 250 mg L^sup -1^ of anthracene and 590 mg L^sup -1^ of glucose (equivalent to 236 mg L^sup -1^ of C as in anthracene), respectively, was also determined.

Effect of PAH Concentration

The effect of different concentrations of anthracene was tested in the range 0, 31.25, 62.5, 125, 250, and 500 mg L^sup -1^ of TMM. The medium contained 50 mg L^sup -1^ of TTC (tetrazolium triphenyl chloride). The flasks were incubated in triplicate with orbital shaking (150 rpm) at 30C. After 30 days, the reduction of TTC to TPF (triphenyl formazan) was evaluated, and used as indication of the metabolic activity of the microorganisms at each concentration (Alef, 1995).

Growth of the Isolates in the Mineral Medium

Growth of bacterial and fungi isolates in the mineral medium containing 250 mg L-1 of anthracene was evaluated for 36 days. The microbial consortium was inoculated into flasks containing 100 ml of mineral medium plus 250 mg L^sup -1^ of anthracene and incubated in triplicate with orbital shaking (150 rpm) at 30C. Colonyforming units (CFU) in culture samples was determined by plating on agar medium. Changes in culture pH were also determined.

Identification of the Microbial Monocultures by 16S rRNA Sequencing

For the initial characterization of the bacterial isolates, the cellular morphology was evaluated by Gram staining. The bacterial isolates selected were identified by 16S rRNA sequencing as described by Camargo et al. (2003). Briefly, bacterial colonies were suspended in nuclease-free water. DNA was extracted from the suspension according to the method described by Asubel et al. (1997). Colonies suspended in a mixture of TE (Tris EDTA) buffer, sodium dodecyl sulfate (SDS) (10%), and proteinase K were incubated for 1 h at 37C. Then, NaCl (5 M) and CTAB/NaCl solutions (4.1 g NaCl and 10 g N-cetyl-N,N,N-trimethylammoniumbromide [CTAB]) in 100 ml prewarmed distilled water) were added and incubated for 10 min at 65C. The solution was extracted with 780 μl chloroform-isoamyl alcohol (24:1), centrifuged for 5 min, and the aqueous phase further extracted with an equal volume of phenolchloroform-isoamyl alcohol (25:24:1). After centrifugation for 5 min, the DNA present in the aqueous phase was precipitated with 0.6 volume isopropanol and the precipitate washed with 70% ethanol. The DNA pellet was dried using a lyophilizer and resuspended in nuclease-free water. Universal bacterial primers corresponding to E. coli positions 27F and 519R were used for polymerase chain reaction PCR amplification of the 16S rRNA gene. A polymerase chain reaction (PCR) master mix (M7502; Promega, Madison, WI) was used according to the manufacturer’s instructions. Genomic DNA template (1 μl) was amplified using a 35-cycle PCR (initial denaturation, 95C for 3 min; subsequent denaturation, 95C for 1 min; annealing, 55C for 1 min; extension, 72C for 1 min; and final extension, 72C for 5 min). The PCR product was analyzed on 2% agarose gel and purified using a QIAEXII gel extraction kit (QIAGEN, Valencia, CA), according to the manufacturer’s instructions. Briefly, the gel slice was suspended in buffer QXI and the QIAEXII suspension was added and mixed by vortexing. The suspension was incubated at 50C for 10 min, centrifuged for 30 s, and the pellet washed twice with QXI buffer and once with buffer PE. After air-drying for 15 min, the DNA was eluted with nuclease free water. DNA cycle sequencing was performed using a BigDye terminator kit (Applied Biosystems, Foster City, CA) and an Applied Biosystems ABI 3100 genetic analyzer. The fungus DNA was extracted according to Schabereiter-Gurtner et al. (2001). Fungal ribosomal RNA (ITS1-5,8S-ITS2) was amplified, using the primers ITS1 (5′-TCCGTAGGTGAACCTGCGG3′) and ITS4 (5′- TCCTCCGCTTATTGATATGC-3′). The amplicons were purified and gene sequence was analyzed using MegaBACE 1000 sequencer (Amersham Biosciences, Piscataway, NJ). The homology of the bacterial and fungus sequences was obtained through MEGABLAST (Altschul et al., 1997). The nucleotide sequences were compared to all sequences in the National Center for Biotechnology Information (NCBI) database using Blast analysis (Altschul et al., 1997). Sequences obtained in this study were combined into a database with the most similar 16s sequences retrieved from GenBank and used to build a distance tree using the neighbor-joining algorithm. The consensus tree was generated using MEGA package version 3.1 (Kumar et al., 2004).

Analysis of PAH Degradation in the Mineral Medium by Gas Chromatography

The capacity of the microbial consortium to degrade PAHs was evaluated in 50 ml of TMM, plus 250 mg L^sup -1^ of anthracene or phenanthrene or pyrene. The medium was inoculated with the cultures or noninoculated (control) and incubated in triplicate at 30C for 30 days. Thereafter, each flask containing the growing culture was acidified to pH 2.0 and extracted with 20 ml dichloromethane (EM Science; >99.8%) in a separation funnel. Excess water was removed by adding sodium sulphate (Shuttleworth and Cerniglia, 1996). Extracted material was quantified in a gas chromatograph-mass spectrometer (Agilent, Palo Alto, CA), GC model 6890, mass selective detector model 5973, and a 30 m 0.25 mm 0.25 mm DB55% phenyl methyl siloxane capillary column. The injector and transfer line temperature were set at 290C, and the temperature program was 1 min at 40C, ramp to 220C at 6C min^sup -1^, maintaining isotherm for 1 min, and ramp to 300C at 15C min^sup -1^. A 0.2-ml aliquot was injected at a split rate of 1:50. The mass selective detector was operated in the scan mode to obtain data for identification of hydrocarbon components. Before injection into the gas chromatograph, deuterated phenanthrene (250 mg L^sup -1^) was added as a surrogate. The percentage of anthracene degradation was calculated relative to anthracene concentration in flasks without inoculum (controls) after 30 days of incubation.

Analysis of Surfactant Production by Isolates

Surfactant production by selected isolates was estimated using the emulsification index and by the reduction of surface tension analysis. The isolates were added to mineral medium (50 ml) containing 250 mg anthracene L^sup -1^ (three replicates) and incubated at 30C for 30 days with orbital shaking (150 rpm). Evaluation was performed in the presence and absence of cells in the MM (cells were removed by centrifugation at 10,000 rpm for 30 min, at 4C). A 2-mL aliquot of MM was mixed with diesel oil in a Pyrexglass tube (100 mm 15 mm) using a vortex for 2 min. After that, the tubes were rested for 24 h before the volume and the stability of the emulsion were measured. The surface tension of the supernatant was measured after sample equilibration (1 h at 25C), using a Gibertini tensiometer (Milan, Italy). Distilled water and ethanol were used as control (69.2 and 24 mN m^sup -1^, respectively).

Hydrocarbon Utilization by Microbial Consortium and Isolates

The microbial consortium and its monoculture isolates were evaluated for the capacity to degrade PAHs (pyrene, anthracene, phenanthrene, and naphtalene), monoaromatics hydrocarbons (benzene, ethylbenzene, toluene, and xylene), aliphatic hydrocarbons (1- decene, 1-octene, and hexane), hydrocarbon mixtures (gasoline and diesel oil), intermediary metabolites of PAHs degradation (catechol, gentisic acid, salicilic acid, and dihydroxybenzoic acid), and ethanol. All carbon substrates were added to the mineral medium in the same concentration of C (236 mg L^sup -1^) that was supplied by 250 mg L^sup -1^ of anthracene, except the gasoline and the diesel oil which were added to the mineral medium in the amount of 1% (v/ v). Triplicate cultures were incubated by orbital shaking (150 rpm) for 7 days. A sample of 1 ml of the growing cultures was diluted in saline solution, plated on agar medium, incubated at 30C for 72 hours and the CFU was calculated.


PAH-Degrading Activity of Landfarming Samples

Respirometric experiments revealed that all the landfarming samples displayed substantial production of CO^sub 2^-C compared to the control (Figure 1). This indicates that the 16 years of petrochemical residues treatment at this site promoted enrichment and selection of a microbial population with the capacity to degrade complex aromatic hydrocarbons. The highest production of CO^sub 2^- C was observed in the soil inoculated with th\e sample from area 5. The accumulated CO^sub 2^-C production in soil inoculated with area 5 soil was 90% higher than that inoculated with area 4 soil sample, which produced the second largest amount of CO^sub 2^-C (597 mg kg^sup -1^ of soil). The lowest degree of CO^sup 2^-C production was observed in soil inoculated with area 1 soil sample. The differences can be attributed to the heterogeneity of PAH-mineralizing microbial populations in the soil samples as well as landfarming management practices, chemical composition, rates and time of application of sludge, nutrient addition, irrigation, and soil tillage, among others. Area 5 landfarming soil sample was selected for the enrichment and isolation of PAH-degrading microorganisms.

Isolation and Identification of the PAH Microbial Degraders

A total of six bacteria and one fungus were isolated from further enrichments set up with area 5 sample using anthracene as the PAH after three transfers. Analysis of the rRNA 16S gene sequence and homology searches revealed isolate 1 belongs to the genus Mycobacterium (Figure 2). Members of the genus Mycobacterium have been used in past studies of PAHs biodegradation (Wick et al., 2002; Miyata et al., 2004; Leys et al., 2005; Mutnuri et al., 2005). Johnsen et al. (2005) reported that PAH degradation in soil is dominated by bacterial strains belonging to a very limited number of taxonomic groups especially Mycobactmum and others. The Mycobactmum strain isolated in our study is genetically very close to M.fortuitum (97% identity), reported to degrade aliphatic hydrocarbons and natural and synthetic rubber (Berekaa and Steinbchel, 2000; Linos et al., 2000).

Isolate 2 is most homologous to the genus Bacillus, with nearest type specie B. cereus (96% identity). This genus was reported to be involved in the degradation of aliphatic (Cybulski et al., 2003) and polycyclic aromatic (Kazunga and Aitken, 2000) hydrocarbons. The role of Bacillus spp. in the degradation of complex hydrocarbons has been characterized as that of secondary degraders, using metabolites produced by the primary hydrocarbon degraders (Chaillan et al. 2004). This can be considered as evidence of the metabolic complementarities among the isolated microorganisms. Isolate 3 clusters with the genus Microbacterium, with nearest type strain Microbacterium sp. B16SH (96% identity). Gauthier et al. (2003), Cavalca et al. (2004), and Zhang et al. (2004) isolated bacterial degraders of aromatic hydrocarbon belonging to this genus, notably, M. oleivorans and M. hydrocarbon oxydans, which are involved in crude oil degradation (Schippers et al., 2005). Isolate 4 is more closely related with Gordonia polyisoprenivorans (98% identity). Gordonia species have been shown to enhance solubilization, disintegration, and mineralization of natural and synthetic rubber, aliphatic hydrocarbon (Linos et al., 2000), petroleum (Chaillan et al., 2004), and only recently, PAHs (Mutnuri et al., 2005). Isolates 5 and 6 are both similar to members of the Microbacteriaceae family and are 98% similar to a naphthalene-utilizing bacterium. The only fungus isolated from the consortia was 100% similar to Fusarium oxysporum, reported to degrade pyrene (Cerniglia, 1997) and benzo(a)pyrene (Verdin et al., 2004, 2005).

Dissolved Organic Carbon Concentration in the Mineral Medium

Concentration of organic C dissolved in the medium was evaluated in order to assess possible use of an alternative source of carbon in the medium. The distilled water contained 0.868 mg L^sup -1^ of dissolved organic C. Addition of inorganic nutrients to the distilled water increased the concentration of dissolved C in the mineral medium to 3883 mg L^sup -1^. These levels of organic C are insignificant when compared to levels typically found in standard growth medium which contain up to 5000 mg L^sup -1^ dissolved organic carbon. In the mineral medium spiked with equal amounts of organic C using anthracene and glucose, 33,410 mg L^sup -1^ and 240,100 mg L^sup -1^ were found, respectively. The difference can be attributed to the low solubility of anthracene. The solubility of anthracene in water is about 0.076 mg L^sup -1^ (Verschueren, 2001). At normal temperature and pressure, anthracene, phenanthrene, and pyrene are in crystal forms and produce a superficial layer of micro crystals when added to mineral medium. This prevents dissolution into the aqueous solution and the low solubility is major factor that limits the cellular absorption of PAHs. It presents a methodological difficulty for several procedures in the laboratory, where overestimation is possible due to the presence of micro crystals of anthracene in the mineral medium that may be quantified without, however, being dissolved.

The reduction of TTC (tetrazolium triphenyl chloride), an artificial electron acceptor added to the growth medium, to TPF (triphenyl formazan), a visible rose color product, was taken as an indicator of microbial growth at different concentrations of anthracene (Alef, 1995). Appearance of rose coloration became evident at anthracene concentrations of 125, 250, and 500 mg L^sup – 1^ (data not shown). At a concentration of 62.5 mg L^sup -1^, the rose coloration was weak and was not detected at O and 31.25 mg L^sup -1^. In our study 250 mg L^sup -1^ of anthracene was selected for further studies.

Growth Curves and pH in the Mineral Medium

Figure 3 presents viable bacterial and fungal numbers (CFU) as well as pH changes in the anthracene mineral medium inoculated with the consortium and incubated for 36 days. Bacterial population was higher than the fungi during the whole experiment, and the pH profile showed strong reduction during the evaluation period. The growth curve of the bacteria and of the fungus presented typical behavior of microorganisms that grow under agitation, using crystals of PAHs as source of C and energy, in amount that exceeds its solubility in the mineral medium (Johnsen et al., 2005).

At the beginning of the experiment, the anthracene was at maximal aqueous concentrations in the mineral medium and the microorganisms grew at maximum rates. The growth was only limited by cellular metabolism and not by anthracene bioavailability, because PAH dissolution is fast enough to keep up with the rising substrate consumption by the growing population. This is the exponential phase and happened until the 5th day for bacteria and in 1st day after incubation for fungus.

With the increase in biomass, however, the anthracene demand exceeded its dissolution rate, dropping the concentration of soluble anthracene in the medium and the exponential growth ceased. From the end of the exponential phase to the 15th day for bacteria and during the period between the 2nd and 5th day for fungus, a pseudolinear growth phase, where the biomass growth was limited by rate of dissolution of the anthracene crystals, was observed. The growth rate decreased, but the biomass continued increasing. With the reduction of the anthracene concentration in the mineral medium, the dissolution rate continued to decrease, and the available substrate was enough only for the maintenance of viable biomass. This limited growth and marked the onset of pseudostationary phase, which is usually long due to insufficient substrate to increase population. There was, however, no death phase, because there is a continuous supply of anthracene for the aqueous phase. In our experiment, the pseudostationary was typical with fungus (after the 15th day) and occurred with the bacteria only during a short period after this day, because a decrease in number of bacterial cells was observed.

Analysis of pH changes with growth of the consortium revealed significant reduction in pH due to acidification (Figure 3). Growth of the microbial consortia in the mineral medium caused a rapid decrease in pH reaching 4.25 on the 19th day that corresponded with the onset of decrease in bacterial population, probably due to the accumulation of H^sup +^ or other acidic metabolites in the mineral medium. After the 19th day of incubation, no further drastic reduction in pH was observed. Interestingly, when the fungus was inoculated separately, in the same medium and incubated for 39 days, the pH was stable (approximately 7.0, data not shown). This is an indication that production of acidic metabolites was not associated with the fungus alone. This behavior can be an indication of metabolic complementary among bacteria and fungus in the consortium. Boonchan et al. (2000) reported that the fungus can accomplish the initial steps of the PAH degradation and excrete acidic metabolites partially oxidized that will be used as source of C and energy by bacteria.


The consortium capacity to degrade PAHs was evaluated by gas chromatography after 30 days of incubation in mineral medium containing 250 mg L^sup -1^ of PAHs (Table 1). The microbial consortium degraded the anthracene, phenanthrene, and pyrene added to the medium. The highest rate of degradation was observed with phenanthrene. This is possibly due to its higher solubility than the other PAHs added to the medium. The degradation of anthracene (48%) was higher than pyrene, which is more soluble. However, anthracene has three aromatic rings, whereas pyrene has four aromatic rings, making it more recalcitrant. Zhang et al. (2004) reported rates of degradation in mineral medium of 0.500, 0.333, and 0.083 mg L^sup “1^ day^sup -1^, for anthracene, phenanthrene, and pyrene, respectively.

The broad spectrum of PAHs degrading activity displayed by the consortium is an important characteristic in the selection of microorganisms for soil bioremediation. However, the selection of microorganisms for soil bioremediation should consider not only in vitro biochemical capacity but also the capacity of inoculum to colonize soil (Johnsen et al., 2005). The soil chemical, physical, and biological comple\xity can determine the decline of the inoculated population, by antagonic relationships imposed by the autochthonous populations, as predation and competition; and by physiologic stresses caused by abiotic factors, such as pH, availability of water and air, temperature, and, in the specific case of PAHs, bioavailability of C and energy sources (van Veen et al., 1997). So, even with the use of microbial consortium, the lack of attention to the PAH degradation principles in the soil has been leading the soil inoculation with microorganism PAH degraders (bioaugmentation) to several failures (Johnsen et al., 2005).

Biosurfactant Production during PAH Degradation

Biosurfactant production was monitored weekly for 42 days. At the end of this period, the superficial tension of the medium inoculated with the consortium was, on the average, about 62.7 mN m^sup -1^. The superficial tension of the medium inoculated with the fungus did not decrease from its initial value, staying close of 67 mN m^sup – 1^ throughout the analysis and was not different from the sterile medium (67 mN m^sup -1^) and distilled water (69 mN m^sup -1^). These results indicate that the production of low-molecular-weight biosurfactant was not the strategy preferentially used to increase the PAH bioavailability in the mineral medium. Jacques et al. (2005) selected a Pseudomonas citronellolis isolate that reduced the superficial tension of 69.2 for 36 mN m^sup -1^ after growing for 48 days in the mineral medium containing anthracene. According to Maier (2003), an efficient biosurfactant can reduce the superficial tension from 73 to 30 mN m^sup -1^.

The emulsification index was also evaluated weekly during the 42 days. No emulsification was recorded with the fungus. With the consortium, emulsion formation was detected in the mineral medium, with or without the microbial cells, indicating the production and excretion of high-molecular-weight biosurfactant to the environment. This emulsion was located in the oil/medium interface but preferentially positioned in the mineral medium, suggesting the presence of a biosurfactant with affinity for the hydrophilic phase (Maier, 2003). Johnsen and Karlson (2004) reported that microorganisms can be used other strategies to increase the PAH availability in the aqueous solution, such as the formation of biofilms in the surface of the PAH crystals.

Growth of the Isolates in Other Hydrocarbon Sources

The capacity of members of the microbial consortium to use 18 hydrocarbon C sources is presented in Table 2. The consortium displayed a wide spectrum of substrate utilization. It grew on all of the 18 C sources evaluated. Individually, the bacterial isolates 2, 4, 5, and the fungus showed higher metabolic versatility because they grew in 14 C sources compared to bacterial isolates 1, 3, and 6, which grew in 11 of the hydrocarbon C sources tested. There were great differences in the substrate utilization profile of the intermediary metabolites of the PAH degradation pathway. Gentisic acid was used as C source by all of the members of the consortium, indicating that this can be the main central intermediary metabolite of the PAH degradation pathway for the isolates from the PAH- degrading consortium.

For the monoaromatic hydrocarbons, the bacterial isolates 2, 4, 5 and the fungus grew in the presence of the four compounds. In spite of the simpler structure than PAHs, benzene, ethylbenzene, and xylene were not degraded by isolates 1 and 6. Aislabie et al. (2000) observed that bacterial isolates that degraded PAHs did not grow in the presence of any monoaromatic hydrocarbons. The amount of monoaromatic hydrocarbons added to the mineral medium (equivalent to C-anthracene) can be toxic for some microorganisms. However, the resistance of the isolates to the toxics effects of these compounds is also a characteristic to be considered in the selection of the degraders to soil bioremediation. In this work this toxics effects seem not to have happened because most of the isolates grew in BTEX.

The members of the microbial consortium demonstrated different capacities to use the aliphatic hydrocarbons. The longest chains were used by all the isolates whereas in the presence of hexane, there was growth of only three members of the consortium, supporting the results of Kastner et al. (1994) who also observed that bacteria degraders of PAHs did not grow in the presence of hexane. Utilization of ethanol by all of the members of the consortium is probably a function of its high solubility and simple chemical structure. Gasoline and diesel oil were used as C source by six members of the consortium. These fuels are complex mixtures of hydrocarbons, which include simple aliphatic chains to complex aromatic hydrocarbons that some of the isolates utilized.


The soil sample from landfarming area 5 displayed the highest microbial PAH mineralization activity in soil culture. A total of six bacterial monocultures and a fungus were identified as capable of degrading anthracene as only source of C and energy. The microbial consortium degraded significant amounts of anthracene, phenanthrene, and pyrene present in the mineral medium. The microbial consortium produced emulsification of the medium and degraded a variety of hydrocarbon substrates and its monoculture isolates displayed a wide substrate spectrum of activity, indicating the possibility of using the consortium for bioremediation of sites contaminated with mixtures of polynuclear aromatic and aliphatic hydrocarbons.


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Rodrigo J. S. Jacques

Department of Soil Science, Federal University of Rio Grande do SuL, Porto Alegre, RS, Brazil

Benedict C. Okeke

Department of Biology, Auburn University Montgomery, Montgomery, Alabama, USA

Fatima M. Bento

Department of Microbiology, Federal University of Rio Grande do Sul, Porto Alegre, RS, Brazil

Maria C. R. Peralba

Department of Inorganic Chemistry, Federal University of Rio Grande do Sul, Porto Alegre, RS, Brazil

Flvio A. O. Camargo

Department of Soil Science, Federal University of Rio Grande do Sul, Porto Alegre, RS, Brazil

Address correspondence to Flvio A. O. Camargo, Department of Soil Science, Federal University of Rio Grande do Sul, Porto Alegre, RS, 7712 Bento Conceives Ave. 91541-000, Brazil. E-mail: fcamargo@ufrgs.br

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