Degradation potential and microbial community ...

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Degradation potential and microbial community structure of heavy oil-enriched microbial consortia from mangrove sediments in Okinawa, Japan a

a

Hernando P. Bacosa , Koichi Suto & Chihiro Inoue a

a

Graduate School of Environmental Studie, Tohoku University, Aramaki, Sendai, Japan

To cite this article: Hernando P. Bacosa , Koichi Suto & Chihiro Inoue (2013): Degradation potential and microbial community structure of heavy oil-enriched microbial consortia from mangrove sediments in Okinawa, Japan, Journal of Environmental Science and Health, Part A: Toxic/Hazardous Substances and Environmental Engineering, 48:8, 835-846 To link to this article: http://dx.doi.org/10.1080/10934529.2013.761476

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Journal of Environmental Science and Health, Part A (2013) 48, 835–846 C Taylor & Francis Group, LLC Copyright ISSN: 1093-4529 (Print); 1532-4117 (Online) DOI: 10.1080/10934529.2013.761476

Degradation potential and microbial community structure of heavy oil-enriched microbial consortia from mangrove sediments in Okinawa, Japan HERNANDO P. BACOSA, KOICHI SUTO and CHIHIRO INOUE

Downloaded by [Tohoku University], [Hernando Bacosa] at 05:31 16 March 2013

Graduate School of Environmental Studies, Tohoku University, Aramaki, Sendai, Japan

Mangroves constitute valuable coastal resources that are vulnerable to oil pollution. One of the major processes to remove oil from contaminated mangrove sediment is microbial degradation. A study on heavy oil- and hydrocarbon-degrading bacterial consortia from mangrove sediments in Okinawa, Japan was performed to evaluate their capacity to biodegrade and their microbial community composition. Surface sediment samples were obtained from mangrove sites in Okinawa (Teima, Oura, and Okukubi) and enriched with heavy oil as the sole carbon and energy source. The results revealed that all enriched microbial consortia degraded more than 20% of heavy oil in 21 days. The K1 consortium from Okukubi site showed the most extensive degradative capacity after 7 and 21 days. All consortia degraded more than 50% of hexadecane but had little ability to degrade polycyclic aromatic hydrocarbons (PAHs). The consortia were dominated by Pseudomonas or Burkholderia. When incubated in the presence of hydrocarbon compounds, the active bacterial community shifted to favor the dominance of Pseudomonas. The K1 consortium was a superior degrader, demonstrating the highest ability to degrade aliphatic and aromatic hydrocarbon compounds; it was even able to degrade heavy oil at a concentration of 15%(w/v). The dominance and turn-over of Pseudomonas and Burkholderia in the consortia suggest an important ecological role for and relationship between these two genera in the mangrove sediments of Okinawa. Keywords: Mangroves, heavy oil, hydrocarbons, microbial community, biodegradation, Pseudomonas, Burkholderia.

Introduction Mangrove ecosystems are important components of the tropical and subtropical estuaries and have an indispensable role in coastal and deep-sea fisheries.[1] Mangroves serve as nurseries and support a diversity of living organisms. They also act as natural shields from storms, stabilizers of shorelines, and carbon sinks. However, mangroves are impacted by human activities and are at risk of pollution caused by oil tanker spills as well as chronic pollution from ports, chemical plants, and petrochemical industries.[2] Tanker ships carry and are mainly fueled with heavy oils. Mangrove wetlands, which are usually located in low wave energy shorelines, are highly vulnerable to oil spills. Oil can kill mangrove trees, extensively reduce animal populations and affect overall ecological processes for a considerable period of time. Toxic polycyclic aromatic hydrocarbons (PAHs), which are the major components of heavy oils, can Address correspondence to Hernando P. Bacosa, Graduate School of Environmental Studies, Tohoku University, 6-6-20 Aoba, Aramaki, Aoba-ku, Sendai 980-8579 Japan; E-mail: [email protected] Received September 17, 2012.

settle in coastal and mangrove sediments and present a risk for the habitat and associated benthic faunas.[3,4] Mangrove ecosystems support a rich microbial diversity, and these organisms play a valuable role in nutrient recycling.[5] The maintenance and restoration of microbial communities are thus fundamental to the productivity, conservation, and recovery of mangroves.[6] The presence of oil pollutants and hydrocarbons in mangrove sediments modify the structure of microbial communities.[7,8] These microbial populations hold great potential for the bioremediation of contaminated sites, such as those contaminated with hydrocarbons.[9,10] Bioremediation may be the most appropriate approach for the cleanup of mangroves.[11] However, to realize the potential of using bioremediation for the recovery of impacted mangrove environments, a proper understanding of the microorganisms present is required. One way to characterize the response of microbial populations to pollution is to use conventional enrichment methods coupled with molecular approaches. Molecular methods, such as polymerase chain reaction-denaturing gradient gel electrophoresis (PCR-DGGE), have received increasing attention in the characterization of microbial communities, and PCR-DGGE has been a useful tool in the


836 characterization of bacterial communities in mangrove sediments.[12–14] This technique provides a robust, costeffective, and rapid exploratory approach to ascertain the degree of compositional variation in microbial communities.[15] Some of the most studied mangrove areas in terms of microbial communities and hydrocarbonoclastic bacterial populations are located in Brazil.[2,10,14,16] The predominant microbial populations of different mangrove areas in India and China have also been investigated.[17,18] Finally, the oil- and hydrocarbon-degrading microbial populations of mangroves in Hong Kong,[4] Taiwan,[19] and Australia [8] have likewise been explored. Although many mangrove areas have been characterized, studies on the microbial communities in the mangrove areas of Okinawa, Japan are lacking. In this report, we examine the heavy oil-degrading bacteria obtained from sediment samples using an enrichment method and PCRDGGE. Our aims include the following: (i) to evaluate the heavy oil and hydrocarbon degradation potential of the enriched consortia obtained from the three mangrove sites in Okinawa, Japan; (ii) to investigate the changes in microbial community structure and identify the bacterial populations that are involved in the biodegradation process; and (iii) to obtain a stable and effective heavy oil-degrading consortium.

Materials and methods Field sites and sample collection The sampling sites are located in the central region of Okinawa Main Island and encompass approximately 20 km of coastline (Fig. 1). The identified mangrove swamps are situated in the estuarine areas of the Teima River in Nago City, the Oura River in Nago City, and the Okukubi River in Kin Town. Visual inspection of the sampling sites indicated that there was no contamination by oil pollutants. The mangrove areas are located far from industries and human villages and have no known history of previous oil spills in the surrounding areas. Thus, the sampling sites can be considered as pristine mangrove areas. Surface sediment (top soil) samples of approximately 2 cm were collected randomly using a spoon as a corer. The soil samples were directly placed in sterilized sampling bottles and homogenized. Three samples were obtained from the Teima site (T1, T2, and T3), one sample was obtained from Oura (O), and two samples were obtained from the Okukubi site (K1 and K2). The samples were transported from Okinawa to the laboratory on ice. Enrichment procedure The enrichment procedure was performed immediately upon sample arrival in the laboratory. Microorganisms

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Fig. 1. Map of Okinawa’s main island showing the three sampling sites (Teima, Oura, and Okukubi mangrove areas).

from the mangrove sediments were obtained by selective enrichment using heavy oil A and mineral salt medium (MSM). The MSM composition is described elsewhere.[20] The medium was sterilized by autoclaving at 121◦ C for 20 min. Trace elements from the sediments were filtered through a 0.20-µm filter (Millipore syringe filter unit) prior to addition to the culture medium. The final pH of the medium was adjusted to 7.1 using 1.0 M HCl or 1.0 M NaOH. Initial enrichment was performed to determine the optimal NaCl concentration at which the microbial samples grew. One gram of soil from each homogenized sample was transferred to 30 mL test tubes followed by the addition of 10 mL of sterilized MSM and 1% (v/v) heavy oil. NaCl at concentrations of 0, 1%, 2% and 3% (w/v) was then added to each sample in duplicate. The tubes were incubated at 30◦ C on a rotary shaker at 120 rpm for a period of 14 days. After 14 days, the sediment was allowed to settle, and the supernatants were then diluted with fresh MSM to attain an absorbance of approximately 0.100 at OD 600 nm. The incubation process was repeated with four different NaCl concentrations and 1% (v/v) heavy oil for another 14 days. Microbial growth was measured periodically using a spectrophotometer (Spectronic 20D+) at an absorbance of OD 600 nm against a blank sample. After the second enrichment, bacterial consortia were continually enriched with NaCl at the salinity level that

Structure of heavy oil-enriched microbial consortia from mangrove sediments


showed the most favorable growth. An appropriate dilution of an aliquot from the test tube cultures was performed with MSM. Five milliliters of the diluted seed culture was then transferred into a 300 mL Erlenmeyer flask containing 95 mL of MSM with the addition of heavy oil A to a final concentration of 1% (w/v). The flask was shaken on an orbital shaker at 30◦ C at a speed of 120 rpm. Every 2 weeks, each culture was transferred into fresh MSM medium. To obtain a stable heavy oil-degrading consortium, enrichment was performed repeatedly for 9 months. To monitor the change in microbial community structure during the enrichment, microbial cells were collected at 4, 6, and 9 months. Degradation experiment The degradative ability of the bacterial consortia enriched for 9 months was evaluated using heavy oil A and individual hydrocarbon compounds, including hexadecane (C17), pristine (C19 branched), phenanthrene (3 rings), pyrene (4 rings), and dibenzothiophene (sulfur-containing). These compounds were selected because they represent important components of heavy oil. All hydrocarbons were of analytical grade and were procured from Wako Chemicals Ltd. (Osaka, Japan). To obtain a standard inoculum, individual consortia were grown for approximately 1 week in 1% (v/v) heavy oil at 30◦ C on an orbital shaker at 120 rpm. The cells were then harvested by centrifugation at 3,000 rpm (TOMYhigh speed refrigerated microcentrifuge) for 10 min, rinsed three times in sterile saline solution and suspended in sterile MSM to yield the desired cell density. Degradation experiments were performed using 10 mL of MSM in sealed 125 mL vials. The media containing heavy oil and hydrocarbons were inoculated with the seed culture to provide an initial cell concentration of approximately 105 cells/mL. Heavy oil and hydrocarbon compounds were added at an initial concentration of 1% (w/v) and 1,000 mg/L, respectively. Heavy oil, hexadecane and pristane were added directly to the medium. Phenanthrene and pyrene were prepared in stock solutions with dichloromethane, and dibenzothiophene was prepared in a stock solution of ethyl acetate at a concentration of 10 mg/mL. After the solvent was allowed to evaporate and MSM was added, an ultrasonic wave was applied prior to inoculation of the bacterial cultures. MSM containing the same amount of heavy oil and hydrocarbon substrates but without any microbial inoculum was used as a control to determine abiotic losses. The experimental cultures were conducted in triplicate with shaking at 30◦ C and 120 rpm. After 21 days, duplicate samples of hydrocarbon cultures were extracted for residual substrate. The residual heavy oil concentration was measured after 7 and 21 days. The degradation of K1 at higher oil concentrations was performed in sealed 300 mL Erlenmeyer flasks containing 5%, 10%, and 15% (w/v) heavy

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oil. All experiments were conducted under aerobic conditions. The efficiency of degradation was calculated using the following formula: Degradation efficiency (%) amount of residual substrate in the control medium − amount of the residual substrate in the inoculated medium = amount of residual substrate in the control medium × 100 The bacterial cells were mounted on a hemacytometer and counted under a binocular phase-contrast microscope (Olympus BH-2). Cultures of alkane mixes (containing decane, hexadecane and pristane) and PAH mixes (containing naphthalene, phenanthrene and pyrene) were also prepared for DNA extraction. Heavy oil analysis Residual heavy oil A was extracted using an equal volume of carbon tetrachloride (CCl4 ). Before extraction, the solution was acidified using hydrochloric acid to pH 2–3 and shaken vigorously for approximately 5 min. The mixture was allowed to stand for 10 min prior to extraction. An appropriate dilution was performed, and the sample was analyzed using a Horiba-Oil Content Analyzer (Model OCMA-350, Horiba, Kyoto, Japan). The residual concentration was determined by comparing the absorbance values against a five-point standard curve. Analysis of the residual heavy oil in the cultures containing 5%, 10% and 15% heavy oil was performed following the Japan Industrial Standard Method JIS K 0101 TPH extraction. Ten milliliters of hexane was added, and extraction was repeated twice in a separatory funnel. Approximately 2 g of Na2 SO4 was added to the extracted liquid to remove remaining traces of water. The extracted heavy oil in hexane was transferred into a beaker of known mass, and the solvent was allowed to evaporate on a hot plate at 80◦ C (Pasolina Hot Stirrer CT-3HA, AS ONE Corp., Osaka, Japan). The mass of the remaining oil was weighed immediately after all the solvent had evaporated. Hydrocarbon analysis Gas chromatography was used for the quantification of hydrocarbon compounds. Analyses were performed using a Hitachi (Tokyo, Japan) 663–50 GC equipped with a flame ionization detector (GC-FID). The column for the GCFID was a capillary column with the following features: a length of 30 m; ID, 0.25 mm; film thickness, 0.25 µm (GL Sciences, Inc., Tokyo, Japan). Sample extraction for hexadecane, pristane, phenanthrene and pyrene was performed by mixing the entire volume of the sample with an equal volume of dichloromethane. The mixture was shaken thoroughly


838 for approximately 5 min and centrifuged for 15 min at 3,000 rpm (Kubota 3700). The bottom layer was recovered and transferred to sealed vials in preparation for GC analysis. From the sealed vials, 1-µL samples were injected into the GC. The instrument conditions for the pristane and hexadecane analyses were the following: an initial temperature of 100◦ C for 1 min with a temperature ramp of 40◦ C per min and a final temperature of 300◦ C, maintained for 5 min. For the analysis of PAH, phenanthrene and pyrene, the conditions were as follows: an oven temperature program that began at 60◦ C for 2 min and increased to 300◦ C at a rate of 10◦ C/min and maintained at 300◦ C for 5 min. The injector and detector temperatures were set at 280◦ C and 300◦ C, respectively. Dibenzothiophene was extracted using an equal volume of ethyl acetate. The ethyl acetate extract was centrifuged at 10,000 × g for 2 min and analyzed by GC-FID. The column temperature was maintained at 125◦ C for 2 min, increased to 250◦ C at 20◦ C/min and maintained at this temperature for 7 min. The injector and detector temperatures were set at 270◦ C and 300◦ C, respectively. The residual hydrocarbon concentration was determined by comparing the peak areas of the chromatogram to those of the standard curve. All of the biodegradation results were obtained based on reference to a sterile control. The remaining substrates in the cultures with alkane and PAH mixes were not analyzed. DNA extraction Bacterial cells were obtained from heavy oil, the alkane mix, the PAH mix, and pure hydrocarbon cultures of hexadecane, pristane, phenanthrene, pyrene, and dibenzothiophene. About a grain size of the pelleted cells was used for DNA extraction. Cells were suspended in 40 µL of sterile distilled water. Freezing and thawing of the suspension was performed twice for 10 min at −20◦ C and 10 min at room temperature. Total genomic DNA was extracted by according to a procedure described elsewhere.[21] The supernatant containing the extracted DNA was then collected and stored at −20◦ C prior to further downstream applications.

Bacosa et al. (denaturing), 65◦ C for 1 min (annealing) and 72◦ C for 1 min (extension) with a 1◦ C decrease in annealing temperature every second cycle up to 55◦ C; this was followed by 10 cycles at an annealing temperature of 55◦ C and a final extension for 10 min at 70◦ C. PCR amplification was performed in a 9700 Thermal Cycler (Applied Biosystems, Foster City, CA, USA). The PCR amplification products were verified by 1.5% agarose gel electrophoresis in 1X TAE buffer.

Denaturing Gradient Gel Electrophoresis (DGGE) After verification of the PCR products, the bands were resolved by DGGE using the DCodeTMUniversal Mutation Detection System (Bio-Rad, Hercules, CA, USA). The gel was composed of 8% polyacrylamide (acrylamide: N,N’methylene bisacrylamide ratio, 19:1, Biorad) and a 30% to 70% linear denaturant consisting of urea and formamide. After cooling at 4◦ C, 180 µL of 10% APS (ammonium peroxodisulfate solution) and 18 µL of TEMED (N,N,N,N, tetramethyl ethylendiamine) were added to the 30% and 70% solutions separately and then gradually added to the glass stand. The wells of the acrylamide gels were washed repeatedly with TAE buffer to remove unpolymerized acrylamide. Subsequently, 7 µL of PCR product and 3 µL of loading buffer were mixed and loaded into the wells of the polyacrylamide gel. Electrophoresis was performed at 60◦ C and 70 V for 960 min. The gels were then stained with ethidium bromide for 30 min with shaking and visualized using a UV transilluminator. The bands of interest were excised and incubated overnight in 50 µL of sterile Milli-Q water (Millipore, Billerica, MA, USA) at 4◦ C. The eluted DNA was re-amplified using primers 341F and 518R. A PCR was performed with hot start at 95◦ C for 5 min and cycling parameters of 94◦ C for 1 min, 55◦ C for 1 min, and 72◦ C for 1 min for 25 cycles followed by a final extension at 72◦ C for 7 min. After confirming the presence of bands by gel electrophoresis, the amplified products were purified using a Gene Elute Mammalian Total RNA Miniprep Kit (Sigma-Aldrich) according to the manufacturer’s instructions.

Polymerase Chain Reaction (PCR) PCR was performed in a total volume of 50 µL containing 25 µL of Promega PCR Master Mix, 5 µL of 2 mM MgCl2 , 2 µL of 2% DMSO, 0.5 µL of primer 341 F (5 -CCTAC GGGAGGCAGCAG-3 ) with a GC clamp (5 -CGCCCG CCGCGCGCGGCGGGCGGGGCGGGGGCACGGG GGG-3 ), 0.5 µL of 518 R (5 -ATTACCGCGGCTG CTGG-3 ), 2 µL of the DNA template and nuclease-free water. Amplification using the above primers was performed using the following touchdown PCR program: a hot start of 95◦ C for 7 min followed by 20 cycles at 94◦ C for 1 min

DNA sequencing Purified DNA was used as a template for the sequencing reaction using the Bigdye Terminator v3.1 Cycle Sequencing Kit following the product protocol. Primers 341F and 518R were used to sequence the V3 region of the 16S rRNA gene fragment. The sequencing reaction products were purified by sodium dodecyl sulfate (SDS) using CENTRISEP Spin Columns (Princeton Separations, Inc., Freehold, NJ). Sequencing was performed using a Genetic Analyzer 3130 (Applied Biosystems). The DNA sequences were compared to the sequences available in GenBank.

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Structure of heavy oil-enriched microbial consortia from mangrove sediments Results

Table 1. 16S rRNA gene sequences of the DNA bands detected by PCR-DGGE.


Microbial community dynamics during heavy oil enrichment The microorganisms in the collected sediments were initially cultured at different salinities to determine the optimal NaCl concentration that favored their growth. The microbial communities demonstrated different growth capacities under four different conditions of salinity (0, 1%, 2%, 3% NaCl). Most of the consortia showed optimal growth in the absence of NaCl with the exception of T3 and K1, which exhibited the best growth in the presence of 1% NaCl (data not shown). The microbial consortia were continually enriched with heavy oil at the salinity at which they showed the most favorable growth. The obtained consortia were enriched with heavy oil up to 9 months to determine how the microbial community changed during the process. The DGGE profiles of the bacterial community dynamics of the six consortia during the enrichment culture and the identity of the sequenced bands are presented in Figure 2 and Table 1, respectively. T1 was characterized by four bands representing Burkholderia at 4 months, but these bands tended to decrease as the band corresponding to Pseudomonas increased. T2 was dominated by bands representing Pseudomonas at 4 and 6 months, but the bands corresponding to Burkholderia gradually increased and dominated at 9 months, leading to the disappearance of Pseudomonas bands. In contrast, T3 appeared to be dominated by Burkholderia and other bacteria (represented by several bands) during the initial enrichment, but Pseudomonas bands dominated in the 9-month enrichment culture. The Pseudomonas band tended to increase and finally dominated in the 9-month O consortium. Whereas the T and O consortia were characterized by dramatic changes, the K1 consortium appeared to be stable after 6 months, and it was characterized by prominent bands representing Pseudomonas.

Band 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

Closest relative in GenBank (access as of August 5, 2012) Pseudomonas delhiensis RLD-1 Burkholderia cepacia 717 Pseudomonas delhiensis RLD-1 Pseudomonas delhiensis RLD-1 Burkholderia cepacia 717 Burkholderia cepacia 717 Burkholderia cepacia 717 Burkholderia cepacia 717 Burkholderia sp. R-5630 Pseudomonas delhiensis RLD-1 Pseudomonas delhiensis RLD-1 Burkholderia cepacia 717 Pseudomonas delhiensis RLD-1 Pseudomonas delhiensis RLD-1 Achromobacter insolitus LMG 6003 Pseudomonas delhiensis RLD-1 Pseudomonas jinjuensis Pss 26 Achromobacter insolitus LMG 6003 Pseudomonas stutzeri ATCC 17588 Pseudomonas delhiensis RLD-1 Achromobacter insolitus LMG 6003 Pseudomonas stutzeri ATCC 17588

Accession No.

Similarity

NR043731 192/192 (100%) NR029209 189/190 (99.5%) NR043731 190/191 (99.5%) NR043731 190/193 (98.4%) NR029209 NR029209 NR029209 NR029209 NR042632 NR043731

190/191 (99.5%) 194/195 (99.5%) 194/194 (100%) 191/192 (99.5%) 190/190 (100%) 194/195 (99.5%)

NR043731 194/194 (100%) NR029209 193/194 (99.5%) NR043731 192/193 (99.5%) NR043731 194/194 (100%) NR025685 192/193 (99.5%) NR043731 190/191 (99.5%) NR025226 191/191 (100%) NR025685 192/192 (100%) NR041715 188/192 (97.9%) NR043731 193/193 (100%) NR025685 193/194 (99.5%) NR041715 190/194 (97.9%)

Degradation of heavy oil and hydrocarbon compounds

Fig. 2. Bacterial community dynamics within the microbial consortia after 4, 6 and 9 months of enrichment culture with heavy oil as determined by PCR-DGGE. The denaturant concentration ranges from 30% to 70%.

The consortia enriched for 9 months were used to determine the heavy oil and hydrocarbon degradation potential of the obtained samples. The enriched consortia demonstrated different abilities to degrade heavy oil (Fig. 3). After 7 days, the best degradative capacity was observed for K1, exhibiting 17%, which was almost twice that of the T1, T3 and O consortia. At the end of the 21-day incubation, all consortia degraded more than 22% of heavy oil with K1 as the superior degrader. T1 also degraded 30% of oil in 21 days. Heavy oil is a mixture of several components, and some compounds were degraded favorably by the consortia. The


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Fig. 3. Degradation of heavy oil (1% w/v) by the enriched microbial consortia. Error bars represent the standard deviation of three replicates.

enriched consortia were incubated in MSM containing hexadecane, pristane, phenanthrene, pyrene and dibenzothiophene to determine the hydrocarbon components that were degraded by the enriched consortia. The degradation results are presented in Figure 4. Hexadecane was degraded well by all the consortia. T2, O, K1, and K2 almost equally degraded hexadecane (approximately 70%) in the samples, whereas T1 and T3 degraded hexadecane at approximately 60%. The branched pristane was degraded at between 15–20% by most of the consortia except T1. Phenanthrene and pyrene were not degraded well by T1, T2, and T3 with a degradation percentage below 5%. In contrast, K1 and K2 exhibited the highest degradation of phenanthrene and pyrene. Degradation of dibenzothiophene showed a high degree of variation. T1 and K2 degraded more than 30% of dibenzothiophene, whereas T2 and O degraded only approximately 17%, and T3 had the lowest dibenzothiophene degradative capability. Pyrene was less degraded than phenanthrene. The K1 consortium demonstrated the greatest ability to degrade alkanes and aromatics in this study. Microbial community structure and dynamics in hydrocarbon cultures The microbial consortia enriched with heavy oil was used as a seed culture or a source of inoculant for the degradation of mixed alkanes (decane, hexadecane, pristane), mixed PAHs (naphthalene, phenanthrene, pyrene), and pure hydrocarbon culture media (hexadecane, pristane, phenanthrene, pyrene, and dibenzothiophene). The PCR-DGGE profiles of the consortia enriched in heavy oil and those cultured in hydrocarbon compounds are shown in Figure 5. The identity of the sequenced bands is presented in Table 1.

Fig. 4. Degradation of hydrocarbon compounds by the enriched microbial consortia. Error bars represent the standard deviation of three replicates.

The T1 consortium in heavy oil was dominated by Burkholderia. It is clear that the structures of the bacterial communities in the cultures with different hydrocarbon substrates were completely different from those of the original culture in heavy oil. The shift of the dominant bacterial populations from Burkholderia to Pseudomonas was observed using specific hydrocarbon substrates. Dramatic changes in the community structure were evident in the T3 consortium. Here, in the presence of heavy oil, hexadecane, and pristane, the most intensive band corresponded to Pseudomonas, whereas Burkholderia dominated in mixed aliphatic and mixed aromatic cultures. However, neither Pseudomonas nor Burkholderia were present in aromatic cultures of phenanthrene, pyrene, and dibenzothiophene. It appears that mixtures of alkane and mixtures of PAH favored Burkholderia. The T2 consortium was dominated by Burkholderia in heavy oil, the alkane mix and the PAH mix. The bacterial communities within the hydrocarbon media were distinct from those of the initial community in heavy oil and the alkane and PAH mixes. The active bacterial community shifted towards Pseudomonas when the bacteria were cultured in the presence of the pure hydrocarbon substrates. Thin bands of Burkholderia were also present in pristane, pyrene and dibenzothiophene cultures. The O consortium was generally stable and was composed predominantly of Pseudomonas when it was cultured with different hydrocarbons, except the PAH mix. Similarly,


Structure of heavy oil-enriched microbial consortia from mangrove sediments

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Fig. 5. PCR-DGGE profiles of the enriched microbial consortia cultured in heavy oil and various hydrocarbon compounds. (Oheavy oil; A-alkane mix; P-PAH mix; H-hexadecane; R-pristane; E-phenanthrene; Y-pyrene; D-dibenzothiophene). The denaturant concentration ranges from 30% to 70%. The three gels were run independently.

in the presence of heavy oil and the various hydrocarbon substrates, the predominant populations in the K1 consortium were Pseudomonas, and these were relatively stable. A closer examination of the DGGE profile revealed variations in the location of the dominant bands depending upon the substrate. For instance, bands 16 and 17 were dominant in the heavy oil and alkane mix, respectively. The position of band 16 is distinct from that of 17. The location of the dominant band, 16, in heavy oil appeared to be identical to that of major bands in hexadecane and pristane. The major bands in phenanthrene, pyrene and dibenzothiophene was located in the same position in the gel but was different from that of heavy oil. Sequencing analysis of bands 16 and 17 revealed that they differ in 5 bases of the DGGE-loaded DNA fragment such that band 16 was identified to be closely related to Pseudomonas delhiensis, and band 17 was closely affiliated with Pseudomonas jinjuensis. Although all bands were not sequenced, it can be deduced that there was a shift in the bacterial community depending upon the substrates used, but the shift was within the Pseudomonas genus only. K2 shared very similar DGGE profiles to those of K1, and the profiles were characterized by little change in the microbial community. Minor changes in the position of the bands indicated that the shift was only within the Pseudomonas genus. Achromobacter was also detected in the microbial community in heavy oil, but these bacteria appeared to be reduced or lost in the presence of the pure hydrocarbon substrates. Band 20 of K2 in heavy oil appeared at the same position as that of hexadecane and pristane. Additionally, communities found in phenanthrene, pyrene, and dibenzothiophene

appeared identical to each other but were distinct from those of hexadecane and pristane. Bands with identical positions could represent the same microorganism because they came from the same seed culture. Another characteristic of K2 was the presence of band 22 in pristane, but this band did not lead to significant degradation of pristane. This band has the same 16S rRNA gene sequence as band 19 in the K1 consortium. Unlike band 19, band 22 was not detected in phenanthrene, pyrene and dibenzothiophene cultures. Conversely, band 19 was absent in pristine-cultured K1. One explanation is that bands 19 and 22 represent different strains and have different degrading abilities because they came from different consortia. It should be noted that the DGGE-loaded 16S rRNA gene was just about a 200–bp fragment. Similarity in a 200-bp region does not guarantee similarity to the rest of the segment of the ∼1500 bp 16S rRNA gene. Degradation of different concentrations of heavy oil by the K1 consortium A heavy oil spill in the sea may lead to accumulation of high concentrations of heavy oil in mangrove sediments. Another objective of this study was to obtain a superior and stable consortium. The preceding experiments demonstrated that the K1 consortium had a remarkable ability to degrade heavy oil, demonstrating superior degradation of the 3-ring and 4-ring PAHs, and a relatively stable microbial community in heavy oil and hydrocarbon compounds. K1 was further investigated for its ability to degrade different concentrations of heavy oil. The ability of the K1 consortium to degrade different concentrations of heavy


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Fig. 6. Degradation of heavy oil at different concentrations by the K1 consortium. Error bars represent the standard deviation of three replicates. For comparison, the values for the 1% heavy oil culture were obtained from the previous experiment and expressed as (v/v).

oil is shown in Figure 6. There was no evident difference in biodegradation efficiency after 7 days when heavy oil concentrations were increased from 0.1% to 15%. The biodegradation efficiency was approximately 15–20% except for concentrations of 0.05% (w/v), which resulted in a biodegradation efficiency below 10%. After 21 days, 40% was degraded in 5% (w/v) and approximately 50% was degraded in 10% (w/v) heavy oil cultures. A small increase in degradation was observed in concentrations of 15% (w/v). Only approximately 22% was degraded in 0.05% (w/v) and 0.1% (w/v) heavy oil cultures after the 21-day experiment. In other words, heavy oil was barely degraded at 0.05% and 0.1%. Higher degradation efficiency occurred at higher oil concentrations. Oil degradation efficiency began to decline at 15%, although degradation at this concentration was still greater than 30% and was equal to the amount degraded in 10% (w/v) concentrations after 21 days. As a result of high microbial activity, the pH in 10% and 15% heavy oil cultures dropped to around 5 after 21 days. In contrast, 0.05% and 0.1% heavy oil cultures characterized by low bacterial growth and lower degradation had negligible changes in pH throughout the study period. The K1 consortium cultured at 0.05% and 0.10% heavy oil showed only a 100-fold increase in bacterial density, whereas the 5%, 10%, and 15% cultures demonstrated an approximate 10,000-fold increase, with bacteria reaching a density of 1010 cells/mL at day 14 of incubation (data not shown). There was a general increase in cell density with increasing oil concentrations of up to 15% (w/v).

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Fig. 7. Effect of the different concentrations of heavy oil (% w/v) on the microbial community structure of the K1 consortium. The denaturant concentration is 30% to 70%.

The microbial community structure was nearly identical in 0.05% and 0.10% heavy oil concentrations and was characterized by several bands representing diverse bacterial populations. At increasing oil concentrations, the number of bands was reduced but was accompanied by an increasing intensity of a band that corresponded to Pseudomonas (Fig. 7). At 15% heavy oil, only the single band affiliated with Pseudomonas was clearly detected by PCR-DGGE.

Discussion Hydrocarbon degradation is influenced by a number of environmental factors, such as salinity, temperature and nutrients.[22] Salinity is one of the most variable environmental factors in mangrove sediments, and it can periodically fluctuate in response to tides and freshwater input from rivers and land. The bacterial communities in mangrove sediments are primarily influenced by salinity and total phosphorus levels.[13] The optimal growth shown by most consortia in the absence of NaCl implies that the bacterial communities were acclimatized to lower salinities in the sampling sites. The PCR-DGGE profile demonstrated that even after 9 months of enrichment using heavy oil, the microbial consortia were not yet stable. The presence of several microorganisms was evident in the early stage of the enrichment culture, and this diversity generally decreased, mostly as a result of the selection of bacterial populations that were


Structure of heavy oil-enriched microbial consortia from mangrove sediments able to degrade heavy oil components. The enrichment culture represents a selective pressure strategy that is effective in enriching the hydrocarbon-degraders from environmental samples by enhancing the adaptation of the microbial flora to the available substrates and selecting the microbial populations that are capable of transforming the substrates involved.[23,24] Burkholderia and Pseudomonas were the dominant genera and played key roles in the degradation of oil components. The increasing band representing Pseudomonas and accompanied by a decrease in Burkholderia in T1, and the turn-over of the previously dominating Pseudomonas to a mainly Burkholderia-dominated T2 consortium at 9 months suggest that these two bacteria have an important ecological relationship in the mangroves of Okinawa in the advent of an oil spill. The DGGE cluster of bacteria obtained from the sediments of mangrove areas in Guanabara, Brazil revealed that they harbored different microbial communities, but no differences were found among replicates from the same mangrove area.[14] In the latter studies, the group also detected Pseudomonas and Burkholderia among the predominant microbial populations in the area. In the current study, we did not extract DNA from the sediments, but the DGGE profiles of the consortia enriched for 4 months showed a different microbial community profile for each sampling area and in replicates from the same mangrove area. The pressure exerted by the enrichment culture could have selected for Pseudomonas and Burkholderia as the predominant bacteria after 9 months of enrichment. The number of DGGE bands appeared to have decreased with time. Enrichment cultures in the laboratory usually show lower diversity than those of in-situ mesocosm communities because the selective pressure imposed by laboratory conditions is much stronger than the pressure that occurs in nature.[13] Both the exposure time and the PAH concentration reduced the microbial diversity in mangrove sediments from Hong Kong, and DGGE bands representing Vibrio, Roseobacter, and Ferrimonas were the most abundant after PAH exposure.[7] However, in another study using pyrosequencing of the 16S rRNA gene sequences where Pseudomonas was not detected as predominant, the number of different OTUs detected in contaminated mangrove sediment samples was significantly higher than the number of OTUs detected in non-contaminated samples.[25] The heavy oil degradation potential of the enriched consortia was higher than that of the consortium obtained from the Gulf of Mexico (17% of weathered oil in 14 days) reported by Shelton et al.[26] and of the consortia (8.8% to 29% of Maya heavy crude oil in 15 days) cited by Okoh.[27] Further, the degradative ability of the K1 consortium was comparable to Terrazyme, a commercial microbiological culture (Oppenheimer Biotechnology, Inc.) studied by Hozumi et al.,[28] which degraded 35% of Nakhodka heavy oil in a three-week period. These results suggest that the K1 consortium has the potential to be used for biore-

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mediation due to its superior ability to utilize heavy oil components. In this study, we showed how the enriched bacterial community responded to a variety of substrates. Heavy oil contains various hydrocarbons and related compounds, with its major constituents consisting of saturated and aromatic fractions. Generally, straight alkanes are degraded more readily than branched hydrocarbons. As a result, hexadecane represented an easily degradable hydrocarbon for the consortia. The similarity of the community structure in heavy oil, hexadecane and pristane suggested that alkanes in heavy oil were mainly degraded by the consortia. This result is not surprising, as alkanes or aliphatic compounds are the most degradable components of petroleum oil mainly due to their less complex chemical structures.[22,29] The numbers of alkane degraders in both oiled and bioremediated mangrove sediments were generally higher than the numbers of aromatic-degraders.[8] The more complex PAH compounds are highly recalcitrant substances.[27] Phenanthrene, pyrene and dibenzothiophene are polyaromatic hydrocarbons in which the initial attack of the aromatic ring is mediated by a dioxygenase enzyme. However, the results here showed that in most cases, the degradation of phenanthrene and pyrene was lower than that of sulfur-containing dibenzothiophene. One possible explanation is that dibenzothiophene was not just oxidized by dioxygenase but was also desulfurized. Although not investigated in the current study, some authors have reported Pseudomonas species that desulfurize dibenzothiophene.[30] In the present study, only T1 and T2 showed different bands for phenanthrene and pyrene. Zhou et al.[31] demonstrated that Sphingomonas strains grew rapidly in three-ring, but not four-ring, PAHs, whereas only Mycobacterium-degrading strains dominated in the four-ring PAH-spiked slurry of mangrove sediments, suggesting differences in the dominant bacteria depending on the types of PAH compounds used. The dioxygenase genes involved in PAH degradation are prevalent in various bacterial isolates obtained from surface mangrove sediments.[32] The use of pure hydrocarbon compounds in enrichment cultures of bacterial communities from the sediments may result in a different microbial community structure because different substrates may select different microbial community compositions.[33] We identified the key bacterial genera involved in heavy oil and hydrocarbon degradation. Pseudomonas and Burkholderia were the dominant genera involved in heavy oil degradation, whereas Pseudomonas was mainly responsible for the degradation of aliphatic and aromatic compounds when they were present as sole substrates. Because the consortia were obtained from different locations, Pseudomonas strains in each consortium may be distinct and have varying potential to degrade heavy oil components. As shown in the heavy oil degradation results (Fig. 3), K1, which was dominated by Pseudomonas, had the most efficient degradative capability, whereas T3,


844 which was similarly dominated by Pseudomonas, had the lowest degradation potential. The microbial community in the sediment could vary within site samples and between sites in mangrove areas.[17] Many species of microorganisms stably coexist by interacting with each other in a microbial consortium.[34] A consortium can adapt well to a change in the environment (i.e., a change in the predominant species). When a specific species decreases during stressful conditions, its members can be restored under certain conditions. The relationship is quite complex in the consortium. Some members of the consortium produce the enzymes necessary for the initial attack of the contaminant. Other members may produce the necessary micronutrients for another species, decompose the toxic metabolites produce by other microbes, or produce biosurfactants. These characteristics make a bacterial consortium more suitable for the degradation of complex substances, such as heavy oil and crude oil. The Pseudomonas genus is known to be ubiquitous in many environments and is arguably the most diverse and ecologically significant of the bacteria.[35] Pseudomonas members have been commonly detected as hydrocarbon degraders in mangrove sediments,[4,10,14] and they are known to have simple metabolic systems, including encoding genes for catabolic enzymes and harboring a degradative plasmid,[36] which allows them to utilize a wide range of aliphatic and aromatic hydrocarbons as well as oil mixtures.[37–39] Pseudomonas strains were also the dominant degraders of the Nakhodka oil spilled in the Sea of Japan.[40] Burkholderia was cited by many authors as a species with remarkable nutritional and physiological versatility.[41,42] Several members of this genus possess oxygenase, which is an essential enzyme for the initial attack of hydrocarbon components, and a plasmid that contributes to the versatility of the bacteria.[42] Burkholderia members from various oil contaminated sites have been reported to degrade a wide range of PAHs.[43,44] However, in the current study, Burkholderia did not dominate in the presence of pure PAH compounds. Furthermore, the remarkable dominance of Pseudomonas and Burkholderia together in the consortium, and the turnover of one genus in favor of another is not commonly reported in literature. The dominance of a single band corresponding to Pseudomonas at higher heavy oil concentrations in the K1 consortium suggests the resistance of these bacteria to higher oil concentrations and lower pH levels. The lower degradative capabilities observed at 0.05% and 0.10% heavy oil can be ascribed to two factors. First, it is likely that other bacteria in the K1 consortium did not utilize the hydrocarbon components of heavy oil but instead made use of the intermediate products of oil degradation. Second, these bacterial species could have produced intermediate products that were inhibitory to the activities of Pseudomonas as the main degrader. Evidence for this was the abrupt increase in degradation% (v/v) in the heavy oil culture when

Bacosa et al. these bands were almost negligible in the medium. Higher heavy oil concentrations could already have been toxic to these populations so that their growth was inhibited at 5%, 10% and 15%. Hydrocarbon concentration plays an important role in selecting the hydrocarbon-degrading bacteria.[7] Bacterial density began to decline at 6% heavy crude oil, attaining a maximum cell density of only 109 cells/mL.[42] Similar results were reported by Vieira et al.,[45] who reported that the percent degradation of 10% bunker oil and crude oil by bacterial consortia reached only approximately 26%. The results of this study suggest a high heavy oil degradation potential for the K1 consortium. The capacity of the K1 consortium to grow and to metabolize heavy oil over a wide range of concentrations demonstrates its high degree of tolerance. This consortium was highly versatile and could have potential in bioremediation applications, specifically in treating oil wastes in the environment. Microbial communities respond to oil pollution depending on the history of contamination.[46] Communities in pristine mangroves diverged greatly when compared with those of the contaminated sites, and this was a result of adaptation to the challenges presented by oil pollution.[46,47] Because the mangrove areas sampled were far from human habitation and had no reports of previous oil contamination, the results of the current study represent a typical result for pristine mangroves where microbial communities have greatly diverged from the original microbial structure in soil samples. In a neighboring subtropical mangrove area of Taiwan, the microbial community of phenanthrene- and pyrene-enriched mangrove sediments was consistently dominated by Dyella and Bacillus.[19] The bacterial communities in sediments of urban mangrove forests in Guanabara Bay (Rio de Janeiro, Brazil) were dominated by Alteromonadales, Burkholderiales, Pseudomonadales, Rhodobacterales and Rhodocyclales.[14] Actinobacteria and Bacteroidetes/Chlorobi dominated the sediments of a Florida mangrove forest.[19] The hydrocarbonoclastic bacteria in mangrove sediments in Okinawa appear to be less diverse; the one prominent feature of this community is the dominance of mainly Pseudomonas and Burkholderia.

Conclusions The enriched microbial consortia from mangrove sediments in Okinawa, Japan have comparatively high heavy oil-degrading potential and primarily degraded the alkane components of heavy oil. They had relatively lower PAHdegrading ability. The K1 and K2 consortia from the Okukubi site had higher PAH degradation potential when compared with the Teima consortia. Pseudomonas and Burkholderia were the dominant genera in the enriched consortia. When incubated in the presence of hydrocarbon compounds, the active bacterial community shifted to favor


Structure of heavy oil-enriched microbial consortia from mangrove sediments the dominance of Pseudomonas, even in the Burkholderiadominated T1 and T2 consortia from the Teima site. The K1 consortium from the Okukubi site was identified as the most efficient consortium, having high ability to degrade alkanes, PAHs and heavy oil at high concentrations. K1 demonstrated a very good capability to degrade heavy oil and hydrocarbons. This consortium has potential for future bioremediation applications. The dominance of Burkholderia and Pseudomonas and the turn-over of the populations depending on the substrate is an interesting feature of the enriched consortia. These two genera are known to be hydrocarbon degraders, but the relationship between them with respect to how they influence each other requires further investigation. The findings of this study suggest that Pseudomonas and Burkholderia have critical roles in the removal of hydrocarbons in mangroves in Okinawa, and these two genera thus serve an important ecological function in hydrocarbon degradation. This study revealed that Pseudomonas and Burkholderia could be the most dominant bacteria in the advent of a sudden oil spill in the mangrove areas of Okinawa, Japan. To the best of our knowledge, our work is the first to characterize the bacterial populations in Okinawa, and we anticipate that our findings will contribute to the understanding of the bacterial populations that could be involved during sudden oil spills. Studies focusing on the unexplored oil-degrading microbial populations of the pristine mangrove areas of Okinawa could be useful in informing the coastal management and bioremediation measures that may be required in the advent of a sudden oil spill.

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