Hydrocarbon Degradation and Bacterial Community Responses During Remediation of Sediment Artiﬁcially Contaminated with Heavy Oil

Biocontrol Science, 2017, Vol. 22, No. 4, 187－203

Original Hydrocarbon Degradation and Bacterial Community Responses During Remediation of Sediment Artiﬁcially Contaminated with Heavy Oil

SHARON N. NUÑAL1, SHEILA MAE S. SANTANDER-DE LEON2, WEI HONGYI3, ADEL AMER REGAL4, TAKESHI YOSHIKAWA3, SUGURU OKUNISHI3＊, AND HIROTO MAEDA3

1Institute of Fish Processing Technology, College of Fisheries and Ocean Sciences, University of the Philippines Visayas, Miagao, 5023 Iloilo, Philippines 2Institute of Marine Fisheries and Oceanology, College of Fisheries and Ocean Sciences, University of the Philippines Visayas, Miagao, 5023 Iloilo, Philippines 3Education and Research Center for Marine Resources and Environment, Faculty of Fisheries, Kagoshima University, 4-50-20 Shimoarata, Kagoshima 890-0056, Japan 4National Institute of Oceanography and Fisheries, P.O. Box 182, Suez, Egypt

Received 13 February, 2017/Accepted 23 June, 2017

Natural biodegradation of heavy oil in the marine environment can be accelerated by the addition of nutrients or seeding of pre-selected microorganisms. In this study, a microcosm experiment was conducted to investigate the effects of inorganic nutrient supplementation（ biostimulation） and bacterial consortium amendment（ bioaugmentation） on the natural degradative processes of artiﬁcially contaminated sediment. Our results revealed that the addition of nutrients had greater effect on remediation than the addition of bacterial cells. Supplementation of inorganic nutrients promoted and sustained the growth of oil-degrading and heterotrophic bacteria throughout the experimental period. Highest reduction in the total petroleum hydrocarbons, and of their components, n-alkanes, polycylic aromatic hydrocarbons（ PAHs） and alkyl PAHs, were obtained in the biostimulated microcosms. Changes in the bacterial community were monitored by the PCR-DGGE（ polymerase chain reaction-denaturing gradient gel electrophoresis） method targeting the 16S rDNA gene. Results revealed different responses of the bacterial community to the addition of heavy oil and remediation agents. Shifts in the bacterial communities in the seawater were more dynamic than in the sediment. Results of this study showed that addition of remediation agents signiﬁcantly enhanced the natural biodegradation of heavy oil in a sediment- seawater microcosm trial.

Key words ： Microcosms / Hydrocarbon degradation / Bacterial community / Bioremediation.

INTRODUCTION column while some settle at the bottom through the process of sedimentation. Once the oil is interspersed When oil is spilled in the sea, it undergoes changes in in the sediment at the bottom, it can penetrate beyond form through the passing of time brought about by the surface, making its removal difficult and can thus wave action, sunlight and inherent microbial community persist for a longer time（ Amadi et al., 1994; Colombo （Mackay and McAuliffe, 1988; Kingston, 2002）. Some et al., 2005; Reddy et al., 2002）. Natural catalytic components of the spilled oil disperse into the water processes by the microbial community in the seawater and sediment can, however, break down the hydrocar- ＊Corresponding author. Tel & Fax: +81-99-286-4133, E-mail : bons in the oil which can be completely mineralized into

okunish（i a）ﬁsh.kagoshima-u.ac.jp CO2 and water（ Harayama et al., 1999）. In case of a 188 S. N. NUÑAL ET AL.

tanker accident in the sea, the large oil inﬂux causes an individually added. In the present study, the effective- imbalance in the carbon-nitrogen ratio which retards the ness of the bacterial consortium in degrading heavy oil growth of bacteria and subsequent utilization of the in the sediment, which presents more challenges in terms carbon sources（ Yakimov et al., 2007）. Furthermore, of accessibility to the oil as a substrate and to oxygen, the nutrients are also quickly assimilated by microorgan- was investigated. Furthermore, another bioremediation isms, thus depleting the nutrient reserves. This limitation approach, which is the addition of inorganic N and P in nutrient sources and dissolved oxygen, the toxicity of sources, was also explored. Temporal degradation of a high concentration of hydrocarbons and by-products the hydrocarbon components of heavy oil and growth of degradation to the bacterial community, and low of oil-degraders and other heterotrophic bacteria were numbers of the principal oil degraders in the sediment also measured. The effects of these remediation and seawater, can affect the rate and effectiveness of approaches to the natural succession of microbial natural biodegradation（ Boopathy, 2000）. communities were also determined by PCR-DGGE. Although natural biological remediation of an oil- Banding patterns were analyzed quantitatively by using polluted site happens without intervention, several tech- nonmetric multidimensional scaling（ NMDS） to describe niques aimed at enhancing the degradative capacities relationships of the microbial community in the different of the indigenous populations such as supplementation microcosms. Prominent bands were also excised and of nutrients or oxygen（ biostimulation） and addition of sequenced to determine their taxonomy. Generally this microbial cultures（ bioaugmentation） may be applied study is aimed at a better understanding of the degra- （Bento et al., 2005; Liang et al., 2009; McKew et al., dative processes and related shifts in the bacterial 2007; Simons et al., 2012; Thavasi et al., 2011; Tyagi et community structure, which can be helpful in the devel- al., 2010）. These human interventions are known to opment of more rational bioremediation strategies. impact the natural processes occurring within the different bacterial communities in the marine environment（ Tyagi MATERIALS AND METHODS et al., 2010; Vogel, 1996）. Thus, prior to the on site application of these bioremediation approaches, it is impor- Collection of seawater and sediment samples tant to determine microbial responses both at the meta- Sediment samples were collected from 2 locations in bolic and genetic levels. Yamagawa Bay, Kagoshima Japan（ Location 1: 31° The major bottleneck in studying microbial communi- 12'44. 48"N, 130°38'15.57"E; Location 2: 31°12'34.66"N ties is the fact that many environmental bacteria cannot 130°39'24.28"E） and were mixed in equal proportions. yet be cultured by conventional laboratory techniques Initial soil quality indices such as moisture（ gravimetric （Briones and Raskin, 2003; Ranjard et al., 2000）. Thus method）, grain size（ sieve analysis）, pH, Redox, total several advanced molecular methods have emerged to organic Carbon and total organic nitrogen（ CHNS provide more powerful tools to study bacterial commu- analyzer FLASH EA112, Thermo Fisher Scientific, nity dynamics by culture-independent techniques Massachusetts, USA）, and total organic Phosphorus

（Pushpanathan et al., 2014; Yoshida et al., 2006）. One （Concentrated H2SO4 and dilute base extraction method; of the most widely employed DNA fingerprinting tech- Bowman, 1989） were measured. Results of the physico- niques for analyzing phylogenetic diversity is PCR-DGGE chemical characterization are presented in Table 1. （Muyzer et al., 1993）. PCR-DGGE allows profiling of the bacterial community through the banding patterns Preparation of bacterial consortium by which its diversity can be analyzed quantitatively. The bacterial consortium with an established oil- Furthermore, the intensity of bands provides a rough degrading capacity was prepared as we have previously estimate of the relative abundance of each species, and described elsewhere（ Nuñal et al., 2014）. The consor- the nucleotide sequencing of the excised bands tium is composed of strains related to: Pseudomonas provides a phylogenetic and taxonomic interpretation of aeruginosa, Marinobacter mobilis, Gaetbulibacter sp. the obtained amplicons. and Halomonas sp.（ DDBJ Acc. Nos. AB649102, In our previous study, we described the isolation, AB649110, AB649114, AB649101 respectively）. identification and characterization of bacterial strains with potential use in the bioremediation of oil pollution Microcosms Set-up （Nuñal et al., 2014）. From these strains a bacterial The sediment mixture was equilibrated at 26℃ for consortium was formulated and evaluated for its oil- seven days prior to the microcosm assembly. The equili- degrading capacities in an in vitro seawater experiment. brated sediments were then placed in 100 mL Erlenmeyer Results showed significant increase in the degradation flasks covered with perforated paper caps to allow of heavy oil in all treatments receiving the bacterial aerobic conditions. For each treatment, nine flasks were consortium compared to when the bacterial strains are prepared. Each flask contained 50 g sediment and 10 REMEDIATION OF OIL CONTAMINATED SEDIMENT 189

TABLE 1．Physico-chemical characteristics of the sediment eluents were combined, spiked with an internal stan- used in the bioremediation microcosms. dard（ pyrene-d10） and concentrated until 0.2 mL with Variables Values nitrogen stream. Moisture content（ %） 41.82% ± 0.88 Measurements of n-alkanes, polycylic aromatic hydrocarbons（ PAHs） and alkylated PAHs in the extracted Grain size（ sand, %） 81.9 ± 2.6 residual oil were conducted using a gas chromatograph Grain size（ silt/clay, %） 18.1 ± 2.6 （Agilent 6890; Agilent Technologies, USA） equipped pH 8.0 with a mass spectrometer（ Agilent 5973 MSD）. One Redox potentia（l mV） 105 ± 7.5 microliter of the sample was injected by an Agilent 7683B Total organic carbon（ %） 1.72 automatic sampler into a DB-5MS capillary column（ 0.25 mm i. d.×60 m, film thickness 0.25 µm; J&W Scientific, Total organic nitrogen（ %） 0.06 USA）. The temperature pattern was programmed as Total organic phosphate（ %） 0.01 follows: 60℃ for 1 min, 60℃ to 200℃ at a rate of 20℃/ min, maintained at 200℃ for 1 min, 200℃ to 290℃ at a rate of 3℃/min, and then maintained at 290℃ for 5 min. mL of natural seawater（ from the same sampling site） The mass selective detector was operated in the SIM were added to avoid desiccation. The flasks were mode. wrapped in aluminum foil to prevent photooxidation. After the microcosm assembly, the flasks containing the Determination of total cultivable bacterial counts seawater-sediment mixtures were equilibrated again in Total cultivable bacterial counts in both seawater and a temperature-controlled room（ 26℃） for seven days. sediment were determined by the conventional plate Flasks were spiked with 1 % heavy oil at Day 0. The count technique. One g sediment and one mL sample

bacterial consortium was added to three ﬂasks, NH4Cl were collected from each flask. The sediment and

and KH2PO4 was added to another three flasks to seawater samples were suspended in 9 mL 0.75%（ W/ adjust the C: N: P ratio to 100:5:1, and the last three V） NaCl solution. The mixture was then serially diluted, flasks were used for natural attenuation or the control. and 0.1 mL of each dilution was spread on Zobell 2216E All flasks were randomly arranged and incubated at agar. Plates were then incubated at 26℃ and bacterial 26℃ and were gently shaken once daily for 60 days. counts were determined after 3-5 days of incubation. Sampling for residual oil, cultivable bacterial counts and bacterial community shifts were done at days 0, 3, 7, Determination of Most Probable Number（ MPN） 15, 30 and 60. of oil-degrading microorganisms MPN of oil-degraders in seawater and sediment were Extraction of Residual Oil and GC-MS analysis enumerated using the sheen-screen method as described Sediment samples from microcosms were dried at by Brown and Braddock（ 1990）. For seawater, a 1 mL room temperature until constant weights were obtained. sample from each replicate was added to 9 mL of 75% From each replicate, residual oil was extracted using natural seawater and was serially diluted. The same five grams of dry sediments using the method described procedure was followed for sediment samples using a 1 by Uno et al.（ 2010）. Sediment was ultrasonically g sample from each replicate. Three aliquots（ 0.1 mL） extracted twice with dichloromethane–hexane mixture of each serially diluted sample were then inoculated into （1:1, v/v） for 15 min. The extracts were pooled and sterile 24-well microtiter plates containing 1.75 mL of concentrated to 0.2 mL under nitrogen stream. To the sterile Bushnell Haas medium per well. Following inocu- concentrated extract was added 6 mL of 1 M potas- lation, 0.15 mL of heavy oil was applied to each well sium hydrate-ethanol and it was saponified under 90℃ making sure that the whole surface was covered with for 1 h. Then 20 mL of distilled water and 10 mL of oil. The plates were incubated without agitation at 26℃ dichloromethane-hexane mixture（ 1:3, v/v） were added for 21 days. After incubation, wells were scored positive to the solution which was then shaken and centrifuged when oil emulsification was indicated by the disruption at 760 xg for 10 min. The procedure was repeated of the oil sheen. Oil-degrading populations were then twice with the upper phase collected each time. The estimated using the standard MPN table. collected solutions were combined and was completely exchanged with hexane. The hexane solution was then DNA extraction concentrated to 1 mL and loaded on a silica-gel column Environmental DNA was extracted from sediment and （packed in a Pasteur pipette）. The column was washed seawater samples obtained from the different micro- with 10 mL hexane then eluted with another 10 mL cosms. Samples from three replicate microcosms were hexane and finally with 10 mL 1% acetone-hexane. The pooled prior to extraction. For sediment, DNA was 190 S. N. NUÑAL ET AL.

extracted using the PowerSoil DNA Isolation Kit（ MOBIO re-amplified and subjected to DGGE with the same Laboratories, Inc., Carlsbad, CA, USA） following the conditions as described above to compare and confirm manufacturer’s instructions. For seawater, samples the recovery from the first DGGE. The resultant single were pre-filtered with a mixed cellulose ester membrane bands were re-excised, recovered, and re-amplified in filter（ pore size, 0.20 µm; diameter, 25 mm, Advantec®, the reaction volume of 50 µL. The PCR products were Japan） to trap the bacteria. The filter was then trans- then subjected to 1.5% agarose gel electrophoresis at ferred into a 6-well microplate. DNA extraction was then 100 V. The bands were excised using a sterilized surgical performed using the DNEasy Plant Mini Kit（ Qiagen, blade and purified using the MonoFas DNA Purification Hilden, Germany）. Presence of the extracted DNA was Kit （I GL Science, Tokyo, Japan） with an elution volume then confirmed by agarose gel electrophoresis using 4 of 30 µL. Thermal cycle nucleotide sequencing of the µL DNA samples. PCR-amplified 16S rDNA was performed using the ABI PRISM BigDye Terminator Cycle Sequencing Kit Ver. 3.1 PCR amplification of 16S rDNA （Applied Biosystems, Carlsbad, CA, USA） with ≤5 ng The 16S ribosomal RNA gene（ 16S rDNA） was ampli- template DNA and a -357F primer without a GC clamp. fied using PCR primers -357F（ 5’-CCTACGGGAGGCA The products were purified using the BigDye XTerminator GCAG-3’） with a GC clamp（ Muyzer et al., 1993） and Kit（ Applied Biosystems, Carlsbad, CA, USA） and 907r（ 5’-CCGTCAATTCCTTTGAGTTT-3’）（ Yu and analyzed by the ABI PRISM 3500xl Genetic Analyzer Morrison, 2004）. The 20 µL PCR reaction mixture （Applied Biosystems, Carlsbad, CA, USA）. contained Ex Taq buffer, 100 µM of the dNTP mixture, 0.5 µM of the forward and reverse primers, and 0.0025 Cloning and sequencing of 16S rDNA PCR prod- units µL-1 of ExTaq DNA polymerase（ Hot Start Version, ucts with heterogeneous sequences Takara Bio, Otsu, Japan）. The amplification program Excised bands which yield heterogeneous sequences used was touch-down PCR with thermal cycling condi- with direct sequencing were subjected to cloning using tions as follows: initial denaturation of 95℃ for 1 min, 19 the pGEM®-T Easy Vector System cloning kit with JM109 cycles of denaturation at 95℃ for 1 min, annealing at High-Efficiency Competent Cells（ Promega Corp., 62℃ for 1 min with a decreasing temperature of -0.8℃ Madison, WI, USA）. The cloning protocol of the manu- at every cycle, and extension at 72℃ for 1 min, followed facturer was followed. The cloned DNA fragments were by 9 cycles of 95℃ for 1 min, 57℃ for 1 min, and 72℃ amplified by colony PCR with primers -21M13 Control for 1 min, and a final extension of 72℃ for 10 min. PCR Primer（ Applied Biosystems, Carlsbad, CA, USA） and amplifications were performed with the ASTEC PC320 Bca BEST Sequencing Primer RV-P（ Takara Bio, Otsu, thermal cycler（ ASTEC, Fukuoka, Japan）. The PCR Japan）. Nucleotide sequences of the amplified prod- products were visualized by agarose gel electrophoresis ucts were determined as described above. with staining of Gel RedTM（ Biotium Inc., Hayward, CA, USA）. Phylogenetic and Statistical analysis The closest relatives of the 16S rDNA sequences were Denaturing gradient gel electrophoresis and direct determined by the basic local alignment search tool nucleotide sequencing of 16S rDNA （BLAST）. The DGGE banding pattern in each lane was Amplified 16S rDNA fragments were subject to dena- converted into a binary matrix indicating the presence turing gradient gel electrophoresis（ DGGE） with the and absence of each band detected on all the lanes. D-Code System（ Bio-Rad, Hercules, CA, USA）. DGGE Similarity was then examined using the NMDS was performed in a 6% polyacrylamide gel with dena- conducted using PRIMER（ version 5） software turing gradient of 25-55%（ where 100% denaturing （PRIMER Enterprises, Plymouth, UK）. The degree to gels contain 7 M urea and 40% formamide in 0.5 x TAE which the plots matched was assessed using Kruskal’s buffer）. Electrophoresis was run at a constant voltage stress wherein the stress should be less than 0.15 and of 60 V for 16 h at 60℃. Gels were stained with SYBR ideally less than 0.10 for configurations of objects to be Gold（ Invitrogen, Carlsbad, CA, USA） and viewed with considered reliable（ Quinn and Keough, 2002）. the Safe ImagerTM 2.0 blue light transilluminator （Invitrogen, Carlsbad, CA, USA）. RESULTS The DGGE bands were excised from the gels using 200 µL pipette tips. Excised gels were suspended in Degradation of n-alkanes 100 µL TE buffer（ 10 mM Tris-HCl, pH 8.0, 1 mM In all treatments, degradation of n-alkanes decreased ethylene diamine tetra acetic acid） in 1.5 mL micro- with increasing chain length（ Table 2）. All microcosms tubes and were kept at 20℃ overnight to elute the receiving remediation agents showed significantly higher amplified DNA. The eluted DNA fragments were total n-alkane degradation than the control（ p<0.05）. REMEDIATION OF OIL CONTAMINATED SEDIMENT 191

TABLE 2．Reduction in the concentration（ %） of n-alkanes in the different microcosms after 60 days of incubation. Treatments n-alkanes Control Bioaugmented Biostimulated C8-C15 86.30 ± 0.18a 87.14 ± 0.71a 91.37 ± 0.32b C16-C25 54.15 ± 1.06a 66.93 ± 0.35b 86.07 ± 0.29c C26-C33 30.54 ± 1.74a 48.92 ± 0.91b 74.73 ± 0.49c Total n-alkane 52.46 ± 0.72a 65.30 ± 0.21b 84.59 ± 0.16c The means of two replicates are shown with the standard deviation. Different superscript letters indicate signiﬁcant difference at p<0.05.

3,000 (A) Temporal analysis showed that reduction in n-alkanes control ) occurred as early as the third day in all treatments bioaugmented （Fig.1A）. At day 15, a dramatic drop in residual n-alkanes (pp m

biostimulated 2,000 was observed in the treatment with nutrient supplemen-

kane s tation with ～75% of the original oil degraded. Further a l

n - degradation in all treatments occurred until the end of l 1,000 the incubation at 60 days with significantly highest values observed in the biostimulated microcosms. Residu a l 0 ot a

T 0 10 20 30 40 50 60 Degradation of polycyclic aromatic hydrocarbons （PAHs） PAHs are less biodegradable than n-alkanes with the

800 signiﬁcantly highest degradation of 62.82% found in the (B) biostimulated treatment after 60 days（ p<0.05）（ Table

) 3）. In all treatments, most degradation occurred in 2-4 600 ring PAHs including naphthalene, acenaphthylene, (pp m

s acenaphthene, ﬂuorene, phenanthrene, anthracene and

A H 400

P fluoranthene. For these compounds, more than 50% l were decomposed in microcosms receiving biological 200 and nutritional interventions while significantly lower Residu a l levels of degradation were observed in the control treat- ot a

T 0 ment. Generally lower levels of degradation were 0 10 20 30 40 50 60 observed in higher molecular weight（ HMW） PAHs in all treatments. Degradation of benzo［ g, h, i］ perylene and indeno［ 1, 2, 3, -c.d］ pyrene was not detected in the 4,500 control microcosm. Bioaugmentation and biostimulation (C) brought signiﬁcant differences in degradation compared s to natural attenuation in the cases of pyrene, benzo［ a］ A H P 3,000 anthracene and benzo［ k］ fluoranthene. Addition of kyl )

a l nutrients signiﬁcantly improved the degradation of benzo l ［b］ ﬂuoranthene compared to the bioaugmented only

(pp m 1,500 and control treatments. For more complex HMW PAHs Residu a

l such as benzo［ a］ pyrene, benzo［ g, h, i］ perylene,

ot a 0 T dibenzo［ a, h］ anthracene and indeno［ 1, 2, 3-c.d］ 0 10 20 30 40 50 60 pyrene, significant differences in degradation among Incubation time (d) treatments was not found. Temporal changes in total FIG. 1．Temporal changes in the residual n-alkanes（ A）, PAHs residual PAHs showed gradual but steady degradation （B） and alkyl PAHs in the control（ open circles）, bioaug- in all treatments（ Fig.1B）. The microcosm added with mented（ open squares） and biostimulated（ open triangles） nutrients showed faster degradation rates than without microcosms during 60 days of remediation. Values represent supplementation particularly in the first 15 days. the mean of two replicates. Thereafter the degradation rates in the treatments 192 S. N. NUÑAL ET AL.

TABLE 3．Reduction in the concentration（ %） of 16 priority polyaromatic hydrocarbons（ PAHs）（US-EPA） in the different microcosms after 60 days of incubation Treatments PAHs Control Bioaugmented Biostimulated Naphthalene 89.17 ± 3.30a 90.7 ± 4.36a 92.26 ± 1.79a Acenaphthylene 34.09 ± 0.10a 69.8 ± 1.22b 76.71 ± 0.33c Acenapthene 37.71 ± 0.34a 35.1 ± 0.81b 48.20 ± 0.02c Fluorene 72.43 ± 0.31a 74.9 ± 0.45b 79.81 ± 0.63c Phenanthrene 39.64 ± 1.70a 70.0 ± 0.16b 91.84 ± 0.17c Anthracene 58.76 ± 0.42a 63.1 ± 0.93b 69.96 ± 1.25c Fluoranthene 52.83 ± 4.74a 44.9 ± 4.85ab 55.50 ± 1.37b Pyrene 8.75 ± 0.65a 25.0 ± 0.14b 14.47 ± 0.34bc Benzo［ a］ anthracene 25.14 ± 5.1a 31.8 ± 0.37ab 36.36 ± 1.82b Chrysene 18.04 ± 2.14a 27.7 ± 9.96a 28.23 ± 5.25a Benzo［ b］ fluoranthene 19.69 ± 0.01a 20.4 ± 0.33a 25.58 ± 0.46b Benzo［ k］ fluoranthene 10.79 ± 3.33a 28.4 ± 1.75b 31.80 ± 2.60b Benzo［ a］ pyrene 13.99 ± 8.25a 20.7 ± 0.74a 23.25 ± 1.30a Benzo［ g, h, i］ perylene N.D. 20.11 ± 0.16a 22.03 ± 3.71a Dibenzo［ a, h］ anthracene 22.66 ± 2.31a 21.8 ± 0.88a 22.33 ± 0.35a Indeno［ 1, 2, 3-c, d］pyrene N.D. 8.3 ± 4.8a 14.62 ± 0.54a Total PAHs 38.73 ± 0.66a 56.53 ± 0.13b 62.82 ± 0.01c The means of two replicates are shown with the standard deviation. Different superscript letters indicate significant difference at p<0.05. N.D., no reduction was detected

started to diminish but still progressed until the end of differences（ p>0.05） were observed among the values. the 60 day incubation. Degradation of the following alkyl PAHs was not detected in any treatments: 2, 3-dimethyl anthracene, Degradation of alkyl PAHs 2-methyl fluoranthene, 1-methyl pyrene, 4-methyl chry- PAHs with alkyl substitution proved to be the least sene, 7, 12-dimethyl benzo［a］anthracene and perylene. biodegradable among the heavy oil fractions. Results of Slow and gradual degradation of alkyl PAHs was the degradation and the temporal residual total alkyl observed during the course of the 60 days incubation in PAHs are shown in Table 4 and Fig.1C, respectively. All all treatments（ Fig.1C）. By the end of the incubation bioremediation interventions resulted in significantly period, more than 50% of alkyl PAHs were still present higher total degradartion of alkyl PAHs than in the case in all treatments. of natural attenuation or the control（ p<0.05）. Nutrient and bacterial cell supplementations significantly improved Degradation of Total Petroleum Hydrocarbons the degradation of the following alkyl PAHs compared （TPH） to the control（ p<0.05）: all alkyl naphthalenes, diben- Reductions of TPHs in the three treatments after 60 zothiophene, 4-methyl dibenzothiophene and 1- and days of incubation are shown in Fig.2. All the treatments 2-methyl phenanthrenes and 7-methyl benzo［ a］ pyrene. receiving a bioremediation agent gave significantly Among the measured alkyl PAHs, the highest degrada- higher reductions than the contro（l p<0.05）. Significant tion of more than 70% was observed in 1-methyl phen- difference in TPH reductions was also found between anthrene and found in the biostimulated microcosm. the bioaugmented（ 51.93%） and biostimulated（ 58.57 Biostimulation also enhanced significantly the degrada- %） microcosms（ p<0.05）. Natural reduction in TPH tion of all alkyl naphthalenes, 4-methyl dibenzothio- found in the control microcosm only accounted for phene, and 4, 6-dimethyl benzothiophene compared to 36.05%. the control and bioaugmented treatments（ p<0.05）. Although degradation of 1-methyl benzo［ a］ anthra- Bacterial growth cene was detected in all treatments, no significant Temporal changes in viable bacterial counts in REMEDIATION OF OIL CONTAMINATED SEDIMENT 193

TABLE 4．Reduction in the concentration（ %） of alkyl PAHs in the different microcosms after 60 days of incubation. Treatments Alkyl PAHs Control Bioaugmented Biostimulated 2-methyl naphthalene 30.92 ± 0.71a 32.30 ± 1.7a 39.57 ± 0.89b 1-methyl naphthalene 35.21 ± 0.22a 41.74 ± 0.9b 58.78 ± 0.26c 1,2-dimethyl naphthalene N.D. 37.46 ± 0.8a 40.35 ± 0.97b 1-methyl fluorene N.D. N.D. 29.75 ± 0.09a 2,3,5-trimethyl naphthalene 23.08 ± 0.36a 40.78 ± 0.2b 54.71 ± 0.11c Dibenzothiophene 40.94 ± 2.55a 50.47 ± 0.5b 52.43 ± 1.20b 4-methyl dibenzothiophene 43.32 ± 0.62a 45.43 ± 1.4a 51.95 ± 0.08b 4,6-dimethyl dibenzothiophene 41.38 ± 2.21a 44.02 ± 0.5a 51.08 ± 0.57b 1-methyl anthracene 54.89 ± 3.13a 54.11 ± 0.4a 58.61 ± 0.21a 2-methyl phenanthrene 35.05 ± 0.22a 40.49 ± 0.7b 41.77 ± 0.02b 1-methyl phenanthrene 25.73 ± 3.13a 31.84 ± 0.9b 71.38 ± 0.29c 2,3-dimethyl anthracene N.D. N.D. N.D. 2-methyl fluoranthene N.D. N.D. N.D. 1-methyl pyrene N.D. N.D. N.D. 1-methyl benzo［ a］ anthracene 19.75 ± 13.66a 23.53 ± 2.2a 25.79 ± 5.12a 4-methyl chrysene N.D. N.D. N.D. 7,12-dimethyl benzo［ a］ anthracene N.D. N.D. N.D. Perylene N.D. N.D. N.D. 7-methyl benzo［ a］ pyrene 14.98 ± 1.41a 17.34 ± 1.43b 18.69 ± 0.49b Total Alkyl PAHs 27.31 ± 0.41a 40.30 ± 0.02b 43.37 ± 0.06c The means of two replicates are shown with the standard deviation. Different superscript letters indicate significant difference at p<0.05. N.D., no reduction was detected

seawater and sediment are shown in Figs.3A and 3B, ) c respectively. In the seawater, control and bioaugmented ( % 60 b microcosms started with lower bacterial counts at 2.6 x bons 106 and 9.9 x 105 CFU mL-1 respectively at day 0 compared to the microcosm to which nutrients（ 8.0 x 6 -1

Hydroca r 10 CFU mL ） had been added. Bacterial counts in the a u m 40 control microcosms dropped slightly at day 3 but ol e increased gradually until it reached the peak at day 15, Pet r

l then dropped dramatically on day 30 but increased ot a

T again on day 60. The microcosm supplemented with

o f 20 inorganic nutrients showed bacterial counts that remained fairly stable from days 3-7, slightly dropped at day 15 but increased continuously until day 60. Nutrient Reducion supplementation resulted in sustaining high bacterial 0 counts which were maintained until the end of the control bioaugmented biostimulated experimental trial. On the other hand, bacterial counts Treatments in the bioaugmented microcosm showed a similar trend FIG. 2．Degradation of petroleum hydrocarbon in the different to the counts in the control microcosm wherein the microcosms after 60 days of incubation. Values represent the numbers peaked at day 15, dropped at day 30 and mean of two replicates. Error bars show the standard devia- continuously increased until day 60, exhibiting the tion. Different letters above the bars indicate signiﬁcant differ- highest counts among the treatments. ence at p<0.05. In the sediment, highest initial bacterial counts were 194 S. N. NUÑAL ET AL.

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8 8 (B) 1) - (B) g

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N 5 M P u lti v a b l C

l a o t T 2 4 0 10 20 30 40 50 60 0 10 20 30 40 50 60 Incubation time (d) Incubation Time (d) FIG. 4．Changes in the Most Probable Number of oil degraders FIG. 3．Changes in the total cultivable bacterial counts in（ a） in（ a） seawater and（ b） sediment in the control（ open seawater and（ b） sediment in the control（ open circles）, circles）, bioaugmented（ open squares） and biostimulated bioaugmented（ open squares） and biostimulated（ open trian- （open triangles） microcosms during 60 days of remediation. gles） microcosms during 60 days of remediation. Values repre- Values represent the mean of three replicates. sent the mean of three replicates.

control and bacterial cell-augmented microcosm recorded for the microcosm to which inorganic nutrients （Fig.4A）. MPN continuously increased from day 0 to had been added（ Fig.3B）. Continuous bacterial growth day 30 in all treatments until day 30. Thereafter, was evident in this treatment until day 15. Although decreases in oil-degraders were observed on day 60 there was a decreasing trend thereafter, it still exhibited but numbers were higher than the values during the highest cell counts among the 3 treatments after the start of the experimental period. Higher numbers were 60-day incubation. For the control and bioaugmented sustained in the treatment with inorganic nutrients. treatments, bacterial counts showed similar growth In the sediment, the same initial trend as in the patterns. Cultivable bacterial counts decreased slightly seawater was observed wherein microcosms augmented on day 3 but that was followed by a steady increase with the bacterial consortium started with higher MPN until day 15, peaking on day 7. Both microcosms values than in the control and treatment receiving nutri- exhibited a drop in cell counts at day 30 and dramatic ents（ Fig.4B）. Continuous increases in MPN until day increase at day 60. 15 were observed in all treatments. For treatments to which remediation agents had been added, the peak of Changes in MPN of oil degraders oil-degrading populations was at day 7 but for the In the seawater, oil degraders in the nutrient-amended control, numbers still increased until day 30. MPN values treatment showed higher counts（～103） than in the in all treatments declined in all treatments on day 60 but REMEDIATION OF OIL CONTAMINATED SEDIMENT 195

FIG. 5．DGGE analysis of the ampliﬁed 16S rDNA fragments FIG. 7．DGGE analysis of the ampliﬁed 16S rDNA fragments from sediment and seawater samples of the control micro- from sediment and seawater samples of the microcosm cosm. The labels above the lane indicate the sampling days. supplemented with inorganic nutrients（ biostimulated）. The The bands subjected to nucleotide sequencing are indicated labels above the lane indicate the sampling days. The bands by the arrowheads. subjected to nucleotide sequencing are indicated by arrowheads.

were also higher than the initial counts.

Temporal changes in the bacterial population structures in the sediment and seawater of the microcosms The DGGE profiles in Figs.5, 6 and 7 show the temporal changes in the bacterial community structures of the different microcosms with or without remediation agents, while the quantitative analysis using NMDS is shown in Fig.8. For the control treatment（ Fig.8A） drastic changes in the community structure occurred in the seawater environment from day 0 to day 15. The bacterial community in the seawater stabilized from day 15 to 30 and shifted again at day 60. Bacterial structure in the sediment remained fairly stable from day 0 to day 60. For the treatment with added bacterial cells （Fig.8B）, the bacterial community changes in the seawater occurred at the following intervals: day 0 to day 3, day 7 to day 15 and day 30 to day 60. The sedi- FIG. 6．DGGE analysis of the ampliﬁed 16S rDNA fragments ment bacterial community, on the other hand, showed from sediment and seawater samples of the microcosm gradual changes but the positions on the NMDS map augmented with the bacterial consortium（ bioaugmented）. The were located close to one another, indicating similarity labels above the lane indicate the sampling days. The bands in the band proﬁles. with asterisks indicate the position of augmented bacteria as follows: 1, Gaetbulibacter sp.; 2, Pseudomonas sp.; 3, Nutrient addition resulted in temporal changes in the Marinobacter sp.; 4, Halomonas sp. The bands subjected to bacterial community in both the seawater and the sedi- nucleotide sequencing are indicated by arrowheads. ment（ Fig.8C）. Three major shifts in the seawater occurred as follows: from day 0 to day 3, day 7 to day 196 S. N. NUÑAL ET AL.

bands were affiliated with 21 genera distributed among different taxonomic groups such as alpha-, gamma- and deltaproteobacteria, Flavobacteria, Bacteriodetes, Firmicutes, Chlorobi and Chloroflexi. A number of genera were commonly found in more than one microcosm. Generally, changes in the community composition in the seawater were more dynamic than in the sediment in all microcosms. Greater diversities were found in the treatments with added remediation agents compared to the control. In the control treatment, fewer distinct bands could be found; therefore, fewer DNA sequences were retrieved （Table 5）. The bands related to Chlorobium phaeobacteriodes（ A1） and uncultured Chloroflexi（ A2）, both green non-sulfur phototrophic bacterial groups, were detected in both the sediment and seawater from day 0 to day 60. The band associated with Bacillus sp.（ A6） was found from day 0 to day 7 in the sediment but disappeared thereafter. Bands that appeared during the later stages were related to Marinobacter mobilis（ A5） and uncultured Deltaproteobacteria（ A3）. In the seawater, temporal bacterial community composition was also characterized by the appearance and disappearance of certain bands. The bands related to Sulfitobacter sp.（ A10）, a sulfite-oxidizing bacteria and Pseudomonas monteilli（ A12） were both present only at day 0 then disappeared subsequently. Bands appearing after day 0 were related to the following: uncultured Deltaproteobacterium（ A7, day 15-60）, Bacillus sp.（ A8, days 3 and 7）, Thalassobius sp.（ A9, days 7-60） and Bacillus firmus（ A11, days 15-60）. In the bioaugmented treatment（ Table 6）, bands found in both the sediment and the seawater throughout incubation were affiliated with the following species: Chlorobium phaeobacteriodes（ B1） and Uncultured Chloroflex（i B3）. Notable bacteria included Erythrobacter sp.（ B14）, a genera associated with bacterial commu- FIG. 8．Nonmetric multidimensional scaling analysis（ NMDS） nities in oil-contaminated sediments, and Alcanivorax map of the banding patterns from sediment and seawater sp.（ B18） a known alkane degrader, both found in the samples of the:（ A） control,（ B） bioaugmented and（ C） bios- bioaugmented microcosm. The added bacterial strains timulated microcosms. SW, the band profiles of the seawater samples; Sed, the band profiles of the sediment samples. The exhibited different survival trends throughout the 60-day sampling days are also shown as numbers following“ SW” or incubation（ Fig.6）. Gaetbulibacter sp. and Halomonas “Sed”. sp. had better survival rates, lasting until day 60 in the sediment. In the seawater, Gaetbulibacter sp. was detected until day 7. Pseudomonas sp. and Marinobacter 15 and day 30 to day 60. Changes in the sediment sp. survived in the sediment only until day 15 but were occurred only after day 3 and changed gradually until not detected in the seawater. day 60. When the microcosms were supplemented with nutrients, the number of identified genera was greater Taxonomy of the 16S rDNA sequences obtained compared to that in the control and bioaugmented from the excised DGGE bands treatments（ Table 7）. Organisms found in the nutrient- The results of the 16S rDNA sequencing analysis of amended microcosm were the bacteria related to the excised DGGE bands obtained from all the micro- Bacteriodetes（ C3）, Muricauda sp.（ C11）, Roseovarius cosms are shown in Tables 5, 6 and 7. The excised pacificus（ C14） and Bacillus sp.（ C16, C19）. Bands REMEDIATION OF OIL CONTAMINATED SEDIMENT 197

TABLE 5．Closest relatives of 16S rDNA sequences obtained from the excised DGGE bands of the control microcosm based on the BLAST homology search. Band No. Closest relatives in database GenBank Acc. No. Homology Taxonomic groups A1 Chlorobium phaeobacteriodes BS1 NR074363 100% Chlorobi A2 Uncultured Chloroflexi bacterium JQ579888 98% Chloroflexi A3 Uncultured delta proteobacterium AF229449 100% Deltaproteobacteria A4 Muriicola sp. KC839612 97% Flavobacteria A5 Marinobacter mobilis B17 GQ214550 96% Gammaproteobacteria A6 Bacillus sp. JQ030918 100% Firmicutes A7 Uncultured delta proteobacterium AM882608 94% Deltaproteobacteria A8 Bacillus sp. JQ030918 99% Firmicutes A9 Thalassobius sp. D7011 FJ161323 99% Alphaproteobacteria A10 Sulfitobacter sp. KC689814 99% Alphaproteobacteria A11 Bacillus firmus strain KJ-W9 KF011491 100% Firmicutes A12 Pseudomonas monteilii strain SeaH-As4w FJ607352 98% Gammaproteobacteria

TABLE 6．Closest relatives of 16S rDNA sequences obtained from the excised DGGE bands of the microcosm augmented with the bacterial consortium（ Bioaugmented） based on the BLAST homology search. Band No. Closest relatives in database GenBank Acc. No. Homology Taxonomic groups B1 Chlorobium phaeobacteriodes BS1 NR074363 100% Chlorobi B2 Uncultured deltaproteobacterium AM882608 94% Deltaproteobacteria B3 Uncultured Chlorofexi bacterium JQ579888 99% Chloroflexi B4 Uncultured Chlorofexi bacterium DQ811889 96% Chloroflexi B5 Uncultured Bacteriodetes clone GQ249615 95% Bacteriodetes B6 Desulfobulbaceae bacterium HQ400773 94% Deltaproteobacteria B7 Uncultured Bacteriodetes clone DQ394963 98% Bacteriodetes B8 Roseovarius sp. AMV6 FN376425 98% Alphaproteobacteria B9 Desulfobulbaceae bacterium HQ400825 97% Deltaproteobacteria B10 Sphingobacterium sp. BF02-S7 DQ677866 99% Bacteriodetes B11 Uncultured Firmicutes bacterium clone JQ516410 94% Firmicutes B12 Marinifilum sp. ONE-10 KF650761 94% Bacteriodetes B13 Muricauda aquimarina PR54-6 EU440979 99% Flavobacteria B14 Erythrobacter sp. MON004 KF042026 100% Alphaproteobacteria B15 Muricauda aquimarina PR54-6 EU440979 99% Flavobacteria B16 Muricauda ruestringensis DSM NR074562 99% Flavobacteria B17 Rhodobacteraceae bacterium DQ486502 98% Alphaproteobacteria B18 Alcanivorax sp. TK-23 KC161579 99% Gammaproteobacteria B19 Robiginitalea sp. ZH09 FJ872534 98% Flavobacteria B20 Bacillus firmus KJ-W9 KF011491 99% Firmicutes

related to Chlorobium phaeobacteriodes（ C2） and related to Idiomarina sp.（ C17）, a known PAH degrader, uncultured Chloroflexi（ C12） were also found to be was also found to persist in the seawater. Bands related persistent, appearing in both the seawater and the to genera that have a hydrocarbon-utilizing member, sediment from day 0 to day 60. Enrichment of bacteria Marinobacter mobilis（ C15）, were also present. 198 S. N. NUÑAL ET AL.

TABLE 7．Closest relatives of 16S rDNA sequences obtained from the excised DGGE bands of the microcosm supplemented with inorganic nutrients（ Biostimulated） based on the BLAST homology search. Band No. Closest relatives in database GenBank Acc. No. Homology Taxonomic groups C1 Peptostreptococcacea bacterium GU194175 97% Firmicutes C2 Chlorobium phaeobacteriodes NR074363 99% Chlorobi C3 Bacteriodetes bacterium JQ683778 99% Bacteriodetes C4 Marinifilum sp. ONE-10 KF650761 95% Bacteriodetes C5 Flexibacter sp. cu2i-19 JN594614 100% Flavobacteria C6 Myxosarcina sp. NTIA-05 EU583792 97% Cyanobacteria C7 Coccinistipes vermicola IMCC1411 EF108212 97% Bacteriodetes C8 Idiomarina sp. YCWA67 FJ984796 98% Gammaproteobacteria C9 Virgibacillus sp. B6B HQ433456 98% Firmicutes C10 Uncultured Chloroflexi bacterium JQ579888 99% Chloroflexi C11 Muricauda ruestringensis NR074562 99% Flavobacteria C12 Uncultured Chloroflexi bacterium AB433096 99% Chloroflexi C13 Chlorobium luteolum DSM273 NR_074096 98% Bacteriodetes C14 Roseovarius pacificus IMCC17041 KC593284 99% Alphaproteobacteria C15 Marinobacter mobilis HH2 HQ284162 100% Gammaproteobacteria C16 Bacillus sp. HWE-119 JQ723721 99% Firmicutes C17 Idiomarina sp. YCWA67 FJ984796 98% Gammaproteobacteria C18 Marinobacterium sp. IC961 AB196257 99% Gammaproteobacteria C19 Bacillus sp. Asd14 JQ030918 99% Firmicutes

DISCUSSION and Chandran, 2011; Welander et al., 2005）. It is a common observation in several studies that when the The present study was conducted to investigate the native microbial populations are allowed to grow opti- effects of bioaugmentation and biostimulation on hydro- mally through nutrient supplementation, hydrocarbons carbon degradation and bacterial community shifts are degraded more effectively compared to when a new during bioremediation in a marine environment. A sedi- population not inherent to the system is added ment-seawater microcosm experiment was designed to （Karamalidis et al., 2010; Kauppi et al., 2011; Malina determine temporal variations in total and oil-degrading and Zawlerucha, 2007）. Care must however be taken bacterial growth, residual hydrocarbons, and bacterial when implementing nutrient amendment as excessive community responses throughout the course of the addition of N and P may promote unnecessary growth different remediation interventions. of populations not specialized for hydrocarbon degra- Degradation reactions can take place without inter- dation which may outgrow the oil degraders in the vention but the speed at which these reactions proceed system（ Chaineau et al., 2005; Wang et al., 2010）. was improved greatly by the added remediation agents Furthermore, excess nitrogen can inhibit microbial in the present study. After the 60 days of incubation, growth due to ammonia toxicity（ Agarry and Ogunleye, higher total n-alkane, PAH and alkyl PAHs degradation 2013; Chaillan et al., 2006）. The 100:5:1 C: N: P ratio was found in treatments amended with a bioremedia- used in the present study proved to be suitable in tion agent of either nutrients or bacterial cells than in the promoting the growth of the indigenous and added case of natural attenuation or the control. Degradation microbial populations as seen in the temporal growth of of hydrocarbon components and total petroleum hydro- oil-degraders and other heterotrophic bacteria. carbons was highest in the biostimulated microcosm However, the addition of inorganic nutrients did not compared to the bioaugmented one. It has been sustain a high number of oil degraders in the seawater reported that in environments where physicochemical and sediment until the end of the 60-day experimental factors such as temperature and pH are constant and period. Decline in the populations at the later part of the optimal, the limiting factor of organic pollutant degrada- remediation process in the nutrient treatment may be tion is the shortage of nitrogen and phosphorus（ Das attributed to the depletion of the added N and P that REMEDIATION OF OIL CONTAMINATED SEDIMENT 199

were utilized during the early stages（ day 1 to day 7） PAHs in the heavy oil were not yet degraded. Addition which was characterized by a spur in the microbial of nutrients and the bacterial consortium may have growth and subsequent decrease in the residual hydro- increased the total degradation of alkyl PAHs; however, carbons. Although addition of the bacterial consortium the degradation of several HMW alkyl PAHs was not did not deliver the best performance compared to bios- detected at all. This may be due to several factors timulation in this study, it still brought significantly higher including recalcitrance of the alkyl PAHs, depletion of total degradation of the different hydrocarbon groups nutrients and limitations of the hydrocarbon-degrading compared to that in the control. The generally lower abilities of the indigenous bacteria and the added hydrocarbon degradation resulting from cell addition consortium（ Makkar and Rockne, 2003; Xu and Obbard, relative to nutrient amendment indicate that the limiting 2004）. factor of the Yamagawa Bay seawater and sediment In hydrocarbon-contaminated marine areas, the was the bioavailability of N and P and not the popula- natural succession of bacterial communities including tion of indigenous degraders. the increase of hydrocarbon-degrading microorganisms The bacterial consortium has already been tested for is usually observed（ Harayama et al., 1999; Pengerud its hydrocarbon-degrading ability in a seawater micro- et al, 1994; Yu et al., 2011）. Subsequently, bacterial cosm experiment and it was found to degrade short- groups that metabolize more recalcitrant oil compounds and long-chain n-alkanes, 2-4 ring PAHs and low replace the groups that have limited hydrocarbon- molecular weight alkyl PAHs（ Nuñal et al., 2014）. In the degrading ability. In this study, microcosms simulating present study, data of total n-alkane degradation show an oil-contaminated marine environment showed shifts significant statistical differences among the treatments in the bacterial communities after addition of heavy oil （p<0.05）, which signify that each intervention brought and remediation agents. Results of the NMDS analysis dynamic changes in the capacity of the system to revealed that:（ 1） responses of the bacterial communi- degrade the n-alkanes of heavy oil. Degradation of ties to the addition of the different remediation agents n-alkanes was faster in the biostimulated treatment with varied;（ 2） bacterial communities in the seawater ～75% of degradation occurring during the first 15 days changed more dynamically than in the sediment in of incubation. When left untreated, the indigenous response to oil and remediation agent inputs; and（ 3） populations were able to degrade n-alkanes albeit in a based on DGGE banding patterns, compositions of the much slower rate, degrading only half of its original bacterial communities in the seawater and the sediment concentration after 60 days. Faster and greater n-alkane were clearly different. degradation was also recorded in similar studies when Although their numbers are usually very low and hardly the following combination of remediation agents were detected（ Atlas, 1981; Leahy and Colwell, 1990）, used: bacterial consortium, rhamnolipid and NPK fertilizer marine sediment and seawater are known to harbor （Rahman et al., 2002） and nutrients and Pseudomonas hydrocarbon-degraders. Even if no intervention was sp. strain（ Stallwood et al., 2005）. made, the natural succession of bacterial communities PAH and alkyl PAHs are more recalcitrant than in the control microcosm occurred as indicated by the n-alkanes, thereby making them more reliable indices of temporal appearance and disappearance of bands in the effectiveness of a bioremediation intervention. the DGGE profile, although their diversity was lower Although relatively lower than those of the n-alkanes, than in the other microcosms. The most notable shift biodegradation rates of PAHs and alkyl PAHs were was observed at the later part of the experimental improved by the added remediation agents. Slow but period, where the bands related to Marinobacter sp. continuous temporal decreases in residual PAHs were and uncultured deltaproteobacteria appeared between observed in all treatments with reductions mainly attrib- day 15 and 60, with the disappearance of a distinct uted to the degradation of 2-4ring PAHs. Addition of the band related to Bacillus sp.. The genus Marinobacter is bacterial consortium brought about degradation of HMW known to include species that can degrade both aliphatic compounds such as benzo［ g, h, i］ perylene, indeno and aromatic hydrocarbons（ Yakimov et al., 2007; ［1, 2, 3-cd］ pyrene, and 1, 2-dimethyl naphthalene Zhuang et al., 2002）. In the seawater, a more dynamic which were not degraded by the indigenous population. shift occurred in a short time with a disappearance of This may indicate co-metabolism of the added cells bands related to non-hydrocarbon degraders such as with the indigenous population（ Jacques et al., 2008; S u l f i t o b a c t e r s p . （ P u k a l l e t a l . , 1 9 9 9 ） a n d Teng et al., 2010）. Pseudomonas monteilli（ Eloimari et al., 1997） and with The least degradable hydrocarbons were the alkyl an appearance of Bacillus firmus and uncultured PAHs showing the highest degradation of only about deltaproteobacterium. 53% in all the treatments. This indicates that by the end Addition of the oil-degrading consortium and nutri- of the 60-day incubation, more than half of the alkyl ents expanded the microbial diversity of the sediment 200 S. N. NUÑAL ET AL.

and seawater in the microcosms, causing more hydrocarbon-degrading capacity has not yet been pronounced shifts in the bacterial community structure documented, the bacterium was found in anaerobic than in the control. Significant increases in the total microbial communities in groundwater capable of degradation and degradation rates of different hydro- degrading aromatic hydrocarbons（ Yagi et al., 2010）. carbons in these treatments compared to those in the The degradation metabolism involved anaerobic control may be attributed to this mechanism. Most of consumption of aromatic compounds coupled to dissimi- the bacterial species included were affiliated with the latory nitrate reduction. Uncultured Chloroflexi, also groups that have oil-degrading capacity or found in oil- green non-sulfur bacteria, were found as well to be contaminated environments, such as Alcanivorax sp. abundant and persistent in almost all the microcosms. （Kasai et al., 2002; Yoshikawa et al., 2010）, Oceanicola They are known to include diverse members with both sp.（ Yuan et al., 2009）, Peptostreptococcaceae aerobic and anaerobic phenotypes. Chloroflexi have （Sherry et al., 2013; Sun et al., 2014）, Thalassospira been reported to be dominant in activated sludge sp.（ Zhao et al., 2009; Shao et al., 2010）, Erythrobacter （Bjornsson et al., 2002）. Due to their ubiquity, it can be sp.（ Chung and King, 2001; Liu and Liu, 2013）, assumed that they may also play an important role in Idiomarina sp.（ Zhao et al., 2009; Kleinsteuber et al., hydrocarbon degradation in the microcosms, although 2006） and Alcaligenes sp.（ Weissenfels et al., 1990）. underlying mechanisms need to be elucidated. Several facultative or strictly anaerobic bacterial In conclusion, results of this study show that addition groups with oil-degrading capability, such as Muricauda of remediation agents such as inorganic nutrients and sp.（ Hwang et al., 2009）, Desulfubulbaceae（ Savage the bacterial consortium have significantly enhanced the et al., 2010）, Roseovarius sp.（ Vila et al., 2010）, natural biodegradation of heavy oil in the sediment- Tenericutes（ Cardinali-Rezende et al., 2012） and seawater microcosm trial. The PCR-DGGE analysis Thiomicrospira sp.（ Voordouw et al., 1996）, were also indicated a shift in bacterial communities caused by oil enriched. These findings are controversial since the contamination and different bioremediation agents. microcosms were basically kept aerobic. However Sequencing of the excised bands revealed the presence heavy oil added to the microcosm setups, covered the of both aerobic and anaerobic bacterial groups in the surface of seawater or sediment layers, limiting the microcosms that were responsible for degradation of penetration of oxygen and resulting in a transition to an heavy oil. It should be, however, noted that elucidation anoxic condition in the subsurface layers（ Berthe-Corti of the bacterial community dynamics based on and Bruns, 2001）. Furthermore, the initial attack on PCR-DGGE is limited by the bias of PCR-amplification hydrocarbon substances by aerobic microorganisms efficiency among different bacterial species. Despite this consumes dissolved oxygen. In such cases, biodegra- limitation, the results show the differences in the adap- dation of hydrocarbons is carried out by strict or facul- tation mechanisms of the microbial communities in - 2- 3- tative anaerobic bacteria using NO3 , SO4 , Fe（III）, response to the bioremediation of oil contamination.

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