Curr Microbiol DOI 10.1007/s00284-014-0535-6

Effects of Three Polycyclic Aromatic Hydrocarbons on Sediment Bacterial Community Xin-Zhong Zhang • Jian-Jun Xie • Fu-Lin Sun

Received: 18 November 2012 / Accepted: 17 December 2013 Ó Springer Science+Business Media New York 2014

Abstract Mangrove sediment is susceptible to anthropogenic pollutants, including polycyclic aromatic hydrocarbons (PAHs). However, the effects of PAHs on the bacterial diversity in mangrove sediment have been rarely studied. In the present study, the effects of three types of PAHs (Naphthalene, Fluorene, and Pyrene) at three doses on sediment microbial populations were investigated by using denaturing gradient gel electrophoresis (DGGE). After 7 and 24 days of incubation of the three types of PAHs, markedly different patterns were observed in the bacterial communities. Overall, the diversity of bacterial community was suppressed before 7 days but was promoted after 24 days. Multidimensional scaling analysis suggested that the composition of bacterial communities after 7 days was distinctly distant from that after 24 days. Also despite a slight shift of bacterial abundance, the bacterial communities were relatively steady in these sediments after exposure to PAHs. In addition, DGGE suggested that the applications of three PAHs (especially PYR) had considerable effects on bacterial communities. For phylogenetic analysis, bacteria species belonging to Proteobacteria (a-, b-, and c-), Actinobacteria, Chloroflexi,

X.-Z. Zhang School of Life Sciences, Nantong University, Nantong 226019, China J.-J. Xie Zhejiang Ocean University, Zhoushan 316000, China F.-L. Sun (&) South China Sea Institute of Oceanology, Chinese Academy of Sciences, Guangzhou 510301, China e-mail: [email protected]

Bacteroidetes, and Planctomycetes were changed dramatically after treatment with PAHs. These results suggest that PAHs play key roles in the change of bacterial community, which may be important for understanding the relationship between PAHs and sediment microbial ecology.

Introduction Mangrove ecosystems as the intertidal wetlands between tropics and subtropics are a key ecological habitat that links terrestrial and marine environments. They are exposed to anthropogenic pollution from tidal water, river water, and land-based sources. Mangrove ecosystems supply substantially abundant carbon to coastal waters, and affect the global biogeochemical cycling of nutrients. The highly productive and diverse microbial communities, living in tropical and subtropical mangrove ecosystems, continuously transform nutrients from dead mangrove vegetation into sources of nitrogen, phosphorus, and other nutrients that can be used by the plants [11]. Polycyclic aromatic hydrocarbons (PAHs) are hydrophobic compounds composed of two or more fused aromatic rings. Although PAHs are ubiquitous in the environment (natural oil seeps, bush fire, and plant derivatives), anthropogenic activities such as disposal of coal processing, mining accidents, petroleum wastes, and vehicle exhaust have drastically increased their presence in the environment. Because of their hydrophobic nature, PAHs in the aquatic environment rapidly bind with particles and deposit in sediments [16]. The mangrove sediments which are rich in organic carbon and detritus are good at absorbing and preserving PAHs [15, 19]. The total PAH concentration, ranging from 237 to 726 ng in per gram of dry weight in mangrove surficial sediments of

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Deep Bay, shows significant correlation with total organic carbon (TOC), clay content, and Pb concentration. Fourand five-ring compounds are major components of composition pattern profiles of PAHs [33]. Reportedly, total PAH concentration ranges from 356 to 11,098 ng per gram of dry weight in the sediments collected from four mangrove swamps in Hong Kong, and the PAH profiles are dominated by naphthalene (NAP, two-ring PAH), fluorene (FLU), and phenanthrene (three-ring PAH) [27]. The mangrove sediments contain higher percentages of lowmolecular-weight PAHs, which indicates that PAHs in mangrove sediments may originate from oil or sewage pollution [13, 27]. PAHs have received much attention in recent years, because of their toxicity on plants, microorganisms, and invertebrates [20]. Organic pollutants are also toxic to microorganism and can reduce microbial abundance [3]. Reportedly, several Terrabacter spp. isolated from mangrove can degrade pyrene (PYR) below 5 mg/Kg, but microbial activity was completely inhibited by PYR higher than 10 mg/Kg [31]. In addition, petroleum pollution could alter microbial community structure and improve the microbial diversity at the species and subspecies levels [7]. Oil pollution can stimulate the growth of heterotrophic bacteria, alkane-degrading bacteria, and aromatic-degrading bacteria in mangrove sediments [23]. Soil microorganisms are of fundamental importance in energy flow, nutrient cycling, and organic matter turnover [14, 25, 30]. They are very sensitive to even low concentrations of pollutants and can respond rapidly to soil perturbation. Consequently, it is suggested that soil microbial community structure may be used as an indicator of soil quality [1, 24, 28]. Denaturing gradient gel electrophoresis (DGGE) of 16S rRNA gene is a powerful tool to study the bacterial community structures in complex environments [22]. Therefore, the aims of the study are: (1) to investigate the effects of three PAHs (e.g., NAP, FLU, and PYR) on the bacterial community structure of mangrove sediment and (2) to determine whether there is any difference in bacterial diversity and community structures after pollution by the three PAHs.

Materials and Methods Study Site and Sampling Samples were collected from a mangrove wetland in the Pearl River Estuary, Guangdong, China. Surface sediment samples at a depth of 0–5 cm were sealed in sterile sealed polythene bags and stored in an icebox in the dark. Then, the samples were transported to the laboratory immediately and treated within 6 h.

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Experimental Setup Moist sediment samples (200 g, moisture 160 %) were placed in plastic cylinders and stabilized at 25 °C overnight. NAP, FLU, and PYR were dissolved in acetone. The PAH-acetone solution (with different concentrations of PAH) were added to the sediments with the final concentrations of 1, 10, and 100 mg/Kg (wet weight). The controls were prepared by acetone mixed with sterilized water. The sediment samples were maintained moist by spraying water and mixed completely using a sterile glass rod every other day. After 7 and 24 days of incubation, these samples were used to assess the effects of the three PAHs on bacterial communities. Total Community DNA Extraction and PCR Amplification Total community DNA was extracted using a vitagen gel recovery kit from 1.0 g wet sediment according to the method by Zhou [32]. The amount and quality of the extracted DNA were estimated by electrophoresis and compared to a DNA marker. Then, DNA extracts were stored at -20 °C in the Tris–EDTA (TE) buffer before used. Polymerase chain reaction (PCR) was performed on a PTC-200 thermal cycler (Bio-Rad, USA). Then, 16S rRNA sequences were amplified with the primer pair F341GC/ R518, yielding a 230 bp DNA fragment suitable for total community fingerprinting [21].To reduce possible intersample variation, all PCRs were performed in triplicate and pooled together before loading on DGGE gel. The PCR products were analyzed by electrophoresis in 1.5 % agarose gels and ethidium bromide staining. DGGE Analysis DGGE was performed on an INGENYphorU-2 system (Ingeny, Netherlands). PCR products (50 lL) were loaded on 8 % polyacrylamide gels (wt/vol) in 1 9 tris- acetateEDTA (TAE) buffer containing a denaturant at the linear chemical gradient of 60–85 %. The gels were pre-run in 1 9 TAE at 80 V and 60 °C for 30 min before loading the samples, and then run at 120 V and 60 °C for 16 h. Then, the gels were stained with ethidium bromide solution for 30 min and rinsed with distilled water prior to viewing under ultraviolet light in an AlphaImager imaging system (Alpha Innotech, USA). Cloning of 16S rRNA Fragments After DGGE The dominant bands in DGGE were excised using a scalpel blade and incubated at 4 °C overnight in 20 lL of the TE

X.-Z. Zhang et al.: Effects of Three Polycyclic Aromatic Hydrocarbons

buffer before re-amplification. The positions of the excised bands were confirmed with repeated DGGE. The bands showing the expected melting position were amplified with a secondary primer without GC-clamp. The PCR products were purified with PCR purification kit (Takara, Japan), and then ligated into a pMD18-T vector(Takara, Japan). Positive clones were identified by PCR amplification with pMD-18T vector primer pairs T7 (50 -TAA TAC GAC TCA CTA TAG GG-30 ) and M13 (50 -CAG GAA ACA GCT ATG ACC-30 ), using the same program as 16S rRNA amplification. DNA Sequencing and Phylogenetic Tree Positive recombinants were then sequenced using an ABI3730 DNA Sequencer with M13 primer. All the sequences obtained in this study have been assigned to the nucleic acid sequence database GenBank with accession numbers from GU367871 to GU367882. Nucleotide sequences were compared with those in GenBank by the BLAST program for identification. Phylogenetic trees of 16S rRNA gene partial sequences were generated using the neighbor-joining algorithms in Mega 5.0. The level of support for the phylogenies derived from neighbor-joining analysis was gaged by 1000 bootstrap replicates. Statistical Analysis DGGE banding patterns were digitized and processed using BandScan 5.0 and manually corrected for further analyses. The bands were visually identified and distinguished by the migration distance and intensity in the gels. The bands that shared one migration position were considered as the same species. Based on these, each band was numbered and scored in each of the sediment samples. DGGE banding patterns were analyzed by multi-dimensional scaling (MDS) to assess the genetic diversity changes of bacterial communities. For this purpose, dissimilarity indices were recorded in a binary matrix, which was then analyzed on SPSS 18.0 for Windows. If the plots were closer to each other, the DGGE banding patterns were more similar.

Fig. 1 DGGE of 16S rRNA fragments of bacterial population from mangrove sediment samples incubated with three types of PAHs. Lanes CK1 and CK2 are PCR products amplified from the control group collected at 7 and 24 days, respectively; Lanes n1–n3 (7 days)/ N1–N3 (24 days) are PCR products from sediment samples amended with 1, 10, and 100 mg/Kg NAP, respectively; Lanes f1–f3 (7 days)/ F1–F3 (24 days) are PCR products from sediment samples amended with 1, 10, and 100 mg/Kg FLU, respectively. Lanes p1–p3 (7 days)/ P1–P3 (24 days) are PCR products from sediment samples amended with 1, 10, and 100 mg/Kg PYR, respectively. The labeled bands (B5–B16) were excised for sequencing and phylogenetic analysis

Results DGGE was used to examine the effects of the three PAHs on the sediment microbial communities. The number of bands per sample varies between 6 and 24, indicating diverse bacterial communities in the mangrove sediments. The differences between samples are either due to the presence/absence of individual DGGE bands or due to the intensity of co-migrating DGGE bands.

Fig. 2 Shannon diversity index values of PCR-DGGE profiles for mangrove sediments at 7 and 24-day incubation with the three PAHs. CK1, CK2, n1–n3 (7 days)/N1–N3 (24 days), f1–f3 (7 days)/F1–F3 (24 days), and p1–p3 (7 days)/P1–P3 (24 days) are the same as in Fig. 1

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X.-Z. Zhang et al.: Effects of Three Polycyclic Aromatic Hydrocarbons Fig. 3 Two-dimensional MDS plots of broad-scale differences in bacterial communities. CK1, CK2, n1–n3 (7 days)/N1–N3 (24 days), f1–f3 (7 days)/F1–F3 (24 days), and p1–p3 (7 days)/ P1–P3 (24 days) are the same as in Fig. 1

Bacterial Diversity in Sediment Based on PCR-DGGE The DGGE profile is highly diversified because of the numerous faint bands, suggesting that the structure of the microbial communities in mangrove sediments is rather complex (Figs. 1, 2). DGGE patterns reveal that the bacterial communities are in similar change amendments with NAP or FLU. Bacterial communities at 7 days, compared with those at 24 days, show that the PAH-polluted types (except PYR at 7 days) and incubation time do not cause a significant decrease in the number of DGGE bands (Fig. 2). On contrary, DGGE profiles at 24 days indicate that band intensity slightly increases with incubation time (Fig. 2). The bacterial communities after incubation with NAP are similar with those samples incubated with FLU within 7 days (Fig. 1), suggesting that the diversity does not change abruptly in these incubations. Minor changes of the bacterial community structure occur in the former 7 days under the incubation with NAP or FLU, but major changes are found in the former 7 days under 10 and 100 mg/Kg PYR (in B2 and B3). Many bands become faint and even disappear under PYR incubation. Bacterial community diversity changes and increases after 24 days of incubation (Fig. 1, 2). The predominant bands on the gels can be considered as the most abundant bacterial species in each incubation sample. Some bands are consistently present after 24 day incubation, suggesting that these bacterial species are more tolerant to PAHs or, are able to use PAHs as a carbon source. The quantities of some microbes increase (e.g., B6-B10, B12-B14), whereas those of others decrease (e.g., B5, B11, and B12) or even disappear compared with 7-day incubation. After 24 days

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of incubation, the ‘‘H’’ values of bacteria diversity increase slightly compared with the samples after 7-day incubation, indicating that it is affected by three PAHs at different time points. MDS of DGGE Banding Pattern To compare broad-scale differences between bacterial community profiles, MDS was employed to analyze DGGE band patterns. The two-dimensional plots of MDS scores for sediment samples are shown in Fig. 3. The results demonstrate spatial diversity in sediment bacterial community structures. The results also indicate that bacterial communities of samples after 7-day incubation are significantly different from the samples incubated for 24 days, with MDS stress value of 0.11 (a value below 0.2 indicates that MDS ordination plot is a good spatial representation of the differences). Samples polluted with PAHs (P1, P2, P3, F3, F2, and F1; p1, p2 p3, f3, f2 and f1; n1, n2, and n3) shared one bacteria community structure, separately. MDS shows that the samples incubated with PAHs between 7 and 24 days have distinct bacterial community structures. Identification and Phylogenetic Analysis of the Predominant Bacterial Phylotypes Table 1 lists the results of sequencing analysis of the predominant phylotypes. Most of them are similar to the 16S rRNA sequences cloned from uncultured organisms, which are commonly found in samples from sediment and soil. For phylogenetic analysis, the 16S rRNA sequences of the 12 clones from this study were compared to those in

X.-Z. Zhang et al.: Effects of Three Polycyclic Aromatic Hydrocarbons Table 1 The 16S rRNA sequences obtained from the respective bands in DGGE gels and the closest match to the sequence from GenBank Band

Accession number

Closest relative (accession number)

Class similarity (%)

source

B5

GU367871

Uncultured Chromatiales bacterium (AY711033)

96

Sediment

B6

GU367872

Uncultured beta proteobacterium (EF651588)

98

Cropland

B7

GU367873

Uncultured chloroflexi bacterium (AB448936)

95

Deep subseafloor sediments

B8

GU367874

Uncultured bacterium (EU133975)

100

Soil

B9

GU367875

Uncultured bacterium (FJ202756)

96

Coral

B10

GU367876

Sphingomonas desiccabilis (GU195183)

100

Activated sludge in wastewater

B11

GU367877

Uncultured bacterium (GQ243093)

96

Freshwater sediment

B12

GU367878

Uncultured bacterium (EF393056)

99

River sediment

B13

GU367879

Uncultured bacterium (GU196087)

99

Indian ocean ridge

B14

GU367880

Uncultured bacterium (GQ093936)

95

Skin

B15 B16

GU367881 GU367882

Uncultured alpha proteobacterium (AY922092) Robiginitalea sp. (FJ872534)

95 98

Farm soil Sediment

GenBank. These bacterial sequences can be divided by neighbor-joining into seven clusters: a-, b-, and c-Proteobacteria, Actinobacteria, Chloroflexi, Bacteroidetes, and planctomycetes (Fig. 4). In c-Proteobacteria, the 16S rRNA gene sequences of Band 5 exhibit 96 % similarity with the uncultured Chromatiales. In b-Proteobacteria, Band 6 has 98 % similarity with the uncultured Myxococcales. Band 15 is clustered in the a-Proteobacteria, and has 95 % similarity with Rhizobiales. Bands 13 and 14 belong to Actinobacteria. Bands 7 and 8 have 96 and 100 % similarity, respectively, with the uncultured Chloroflexi. Band 10 has 99 % similarity with the uncultured Planctomycetes.

Discussion Mangroves provide a unique ecological environment for diverse bacterial communities. In the present study, culture-independent analysis was used to reveal the changes in microbial community structure in mangrove sediments exposed to different PAHs. The concentrations of PAHs are close to the levels in natural mangrove sediments. Both incubation time and PAH type will change the microbial community structure, as determined by the number of DGGE bands, suggesting that the mangrove microbial community can be affected by PAH pollution. There are two opposite viewpoints concerning the effects of organic pollutants on microbial diversity. On one hand, organic pollutants can be used by microorganisms as carbon source, and thus, they can increase the microbial diversity [7]. On the other hand, organic pollutants pose a toxic threat to microorganisms, and thereby reduce the species abundance [2]. In the present study, the level of PAH pollution does not obviously correlate with bacterial diversity according to DGGE profile patterns. First of all,

PAHs were slightly toxic to microorganisms and inhibited the number and diversity of microbial community at the beginning of the experiment. But the 24-day incubation allowed the microorganisms to adapt and flourish, so the microbial number and diversity increased. Microbial communities in the sediments may be selected by PAH exposures [12]. Higher microbial diversity was detected after incubation with low and high concentrations of PAHs, indicating that PAHs can activate the microbial community. DGGE indicates that the microbial community structures of the PAH-incubated groups are similar to the control group after 24 days. In the present experiment, a slight shift in the abundance of major bacterial populations was observed in the sediments amended with three PAHs, indicating that the bacterial communities were relatively steady in these sediments. The bioavailability of PAHs is reduced because of their hydrophobicity and sorption onto soil minerals and soil organic matter. PAHs are relatively water insoluble and may be unavailable for microbial degradation. Sorption of organic pollutants onto sediment organic matter can significantly affect biodegradability, as well as biotoxicity [29]. In addition, PAHs may diffuse into sediment particle sites where they are inaccessible to microbes from the outside. On the other hand, in most cases, many bacteria can utilize organic pollutants as a carbon source and involve in the degradation of petroleum hydrocarbons [6]. Many species such as Mycobacterium sp. [4, 5] and Rhodococcus sp. [8] utilize PYR as a sole source of carbon and energy in soil. Although many high-molecular-weight PAH-degrading bacteria (e.g., actinomycetes) were found, a variety of nonactinomycete bacteria have the ability to metabolize fluoranthene, pyrene, chrysene, and benz[a]anthracene. Some bacteria, such as Pseudomonas putida, P. aeruginosa, and Flavobacterium sp. isolated from a soil-derived mixed culture, have the ability to metabolize fluoranthene and PYR

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Fig. 4 Phylogenetic tree based on 16S rRNA sequences representing the respective DGGE bands in Fig. 1. Bootstrap analysis was based on 1,000 replicates. Bootstrap values ([50 %) from distance analysis are depicted

when supplied with other forms of organic carbon [9, 10]. Under the pressure of pollutants, bacteria can proliferate by adjusting their metabolic mechanism. In summary, the three PAHs have slightly negative effects on the bacterial community; PAHs to some extent stimulate bacterial growth in the long term. The DGGE profiles of bacterial community indicate that the number of DGGE bands is significantly decreased after 7 days of PYR incubation. This may be partly because (1) PYR is slowly released from the soil particles, and thus the efficient dose is increased; (2) PYR has more rings, and it is not easy to be degraded. The biodegradation rates of PAHs as higher molecular-weight compounds in soils and sediments are usually much slower [17]; (3) The PYR-degraded products usually possess high toxicity [4, 8]. Other studies indicate that nitrogen-fixing bacteria are easier to be affected by PAHs than total bacterial community. In other words, a small amount of polycyclic aromatic hydrocarbons can drastically change the diversity of nitrogen-fixing bacteria in sediments, while the total bacterial community does not change significantly. Reportedly, particular substances (e.g., nonylphenol) in the

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sludge have special toxicity, and measurement of N2-fixation in sludge or soils is a useful method for predicting the biological quality of organic slurry [18]. PAHs have greater effect on functional microbial community than on the total microbial community. Therefore, nitrogen-fixing bacterial community, instead of total microbial community, can be used as an important parameter for assessing the effects of PAHs on mangrove sediment ecosystem [26]. This study suggests that the three PAHs may have a significant effect on the bacterial community in mangrove sediment. This may be important for understanding the relationship between PAHs and barriers in transformation of sediment nutrients, when assessing the potential adverse effects on the environment. In addition, further research is required to understand the relationship between functional sediment microbial community and ecosystem function. Acknowledgments This study were supported by the Open Foundation from Ocean Fishery Science and Technology in the most important subjects of Zhejiang (20110209), the Knowledge Innovation Program of the Chinese Academy of Sciences (SQ201220), the talent project of Nantong University (No. 13R38) and Public Services Project of Zhejiang Province Science Technology Department (2011F30017).

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References 1. Agnelli A, Ascher J, Corti G, Ceccherini MT, Nannipieri P, Pietramellara G (2004) Distribution of microbial communities in a forest soil profile investigated by microbial biomass, soil respiration and DGGE of total and extracellular DNA. Soil Biol Biochem 36(5):859–868 2. Andreoni V, Cavalca L, Rao MA, Nocerino G, Bernasconi S, Dell’Amico E, Colombo M, Gianfreda L (2004) Bacterial communities and enzyme activities of PAHs polluted soils. Chemosphere 57(5):401–412 3. Bachoon DS, Hodson RE, Araujo R (2001) Microbial community assessment in oil-impacted salt marsh sediment microcosms by traditional and nucleic acid-based indices. J Microbiol Methods 46(1):37–49 4. Balashova NV, Kosheleva IA, Golovchenko NP, Boronin AM (1999) Phenanthrene metabolism by Pseudomonas and Burkholderia strains. Process Biochem 35(3–4):291–296 5. Bird C, Martinez JM, O’Donnell AG, Wyman M (2005) Spatial distribution and transcriptional activity of an uncultured clade of planktonic diazotrophic c-Proteobacteria in the Arabian Sea. Appl Environ Microbiol 71(4):2079–2085 6. Dean-Ross D, Moody J, Cerniglia CE (2002) Utilization of mixtures of polycyclic aromatic hydrocarbons by bacteria isolated from contaminated sediment. FEMS Microbiol Ecol 41(1):1–7 7. Feris K, Hristova K, Gebreyesus B, Mackay D, Scow K (2004) A shallow BTEX and MTBE contaminated aquifer supports a diverse microbial community. Microb Ecol 48(4):589–600 8. Gadzala-Kopciuch R, Kluska M, Welniak M, Kroszczynski W, Buszewski B (1999) Aryl chemically bonded phases for determination of selected polycyclic aromatic hydrocarbons isolated from environmental samples utilizing SPE/HPLC. Pol J Environ Stud 8(6):383–388 9. Goyal A, Zylstra G (1997) Genetics of naphthalene and phenanthrene degradation by Comamonas testosteroni. J Ind Microbiol Biotechnol 19(5):401–407 10. Griffin LF, Calder JA (1977) Toxic effect of water-soluble fractions of crude, refined, and weathered oils on the growth of a marine bacterium. Appl Environ Microbiol 33(5):1092–1096 11. Holguin G, Vazquez P, Bashan Y (2001) The role of sediment microorganisms in the productivity, conservation, and rehabilitation of mangrove ecosystems: an overview. Biol Fertil Soils 33(4):265–278 12. Ka¨stner M, Mahro B (1996) Microbial degradation of polycyclic aromatic hydrocarbons in soils affected by the organic matrix of compost. Appl Microbiol Biotechnol 44(5):668–675 13. Ke L, Yu K, Wong Y, Tam N (2005) Spatial and vertical distribution of polycyclic aromatic hydrocarbons in mangrove sediments. Sci Total Environ 340(1–3):177–187 14. Kennedy A, Smith K (1995) Soil microbial diversity and the sustainability of agricultural soils. Plant Soil 170(1):75–86 15. Kragh T, Søndergaard M, Tranvik L (2008) Effect of exposure to sunlight and phosphorus-limitation on bacterial degradation of coloured dissolved organic matter (CDOM) in freshwater. FEMS Microbiol Ecol 64(2):230–239

16. Latimer S, Zheng JS (2003) Sources, transport, and fate of PAHs in the marine environment. In: Douben PET (ed) PAHs: an ecotoxicological perspective. Wiley, UK, pp 9–33 17. MacRae J, Hall K, Grabow W (1998) Biodegradation of polycyclic aromatic hydrocarbons (PAH) in marine sediment under denitrifying conditions. Water Sci Technol 38:177–185 18. Martensson AM, Torstensson L (1996) Monitoring sewage sludge using heterotrophic nitrogen fixing microorganisms. Soil Biol Biochem 28(12):1621–1630 19. Martins J, Peixe L, Vasconcelos VM (2011) Unraveling cyanobacteria ecology in wastewater treatment plants (WWTP). Microb Ecol 62(2):241–256 20. Menzie CA, Potocki BB, Santodonato J (1992) Exposure to carcinogenic PAHs in the environment. Environ Sci Technol 26(7):1278–1284 21. Muyzer G, de Waal EC, Uitterlinden AG (1993) Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 59(3):695–700 22. Muyzer G, Smalla K (1998) Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie Van Leeuwenhoek 73(1):127–141 23. Ramsay MA, Swannell RPJ, Shipton WA, Duke NC, Hill RT (2000) Effect of bioremediation on the microbial community in oiled mangrove sediments. Mar Pollut Bull 41(7):413–419 24. Salles JF, van Veen JA, van Elsas JD (2004) Multivariate analyses of Burkholderia species in soil: effect of crop and land use history. Appl Environ Microbiol 70(7):4012–4020 25. Schutter M, Sandeno J, Dick R (2001) Seasonal, soil type, and alternative management influences on microbial communities of vegetable cropping systems. Biol Fertil Soils 34(6):397–410 26. Sun FL, Wang YS, Sun CC, Peng YL, Deng C (2012) Effects of three different PAHs on nitrogen-fixing bacterial diversity in mangrove sediment. Ecotoxicology 21:1651–1660 27. Tam N, Ke L, Wang X, Wong Y (2001) Contamination of polycyclic aromatic hydrocarbons in surface sediments of mangrove swamps. Environ Pollut 114(2):255–263 28. van Bruggen AHC, Semenov AM (2000) In search of biological indicators for soil health and disease suppression. Appl Soil Ecol 15(1):13–24 29. Weissenfels WD, Klewer H-J, Langhoff J (1992) Adsorption of polycyclic aromatic hydrocarbons (PAHs) by soil particles: influence on biodegradability and biotoxicity. Appl Microbiol Biotechnol 36(5):689–696 30. Yao H, He Z, Wilson MJ, Campbell CD (2000) Microbial biomass and community structure in a sequence of soils with increasing fertility and changing land use. Microb Ecol 40(3):223–237 31. Zhou HW, Luan TG, Zou F, Tam NFY (2008) Different bacterial groups for biodegradation of three-and four-ring PAHs isolated from a Hong Kong mangrove sediment. J Hazard Mater 152(3):1179–1185 32. Zhou J, Bruns M, Tiedje J (1996) DNA recovery from soils of diverse composition. Appl Environ Microbiol 62(2):316–322 33. Zhang J, Cai L, Yuan D, Chen M (2004) Distribution and sources of polynuclear aromatic hydrocarbons in mangrove surficial sediments of Deep Bay, China. Mar Pollut Bull 49(5):479–486

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