• mangrove;
  • petroleum;
  • rhizosphere;
  • hydrocarbonoclastic bacteria;
  • Guanabara bay


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

In this study, the combination of culture enrichments and molecular tools was used to identify bacterial guilds, plasmids and functional genes potentially important in the process of petroleum hydrocarbon (PH) decontamination in mangrove microniches (rhizospheres and bulk sediment). In addition, we aimed to recover PH-degrading consortia (PHDC) for future use in remediation strategies. The PHDC were enriched with petroleum from rhizosphere and bulk sediment samples taken from a mangrove chronically polluted with oil hydrocarbons. Southern blot hybridization (SBH) assays of PCR amplicons from environmental DNA before enrichments resulted in weak positive signals for the functional gene types targeted, suggesting that PH-degrading genotypes and plasmids were in low abundance in the rhizosphere and bulk sediments. However, after enrichment, these genes were detected and strong microniche-dependent differences in the abundance and composition of hydrocarbonoclastic bacterial populations, plasmids (IncP-1α, IncP-1β, IncP-7 and IncP-9) and functional genes (naphthalene, extradiol and intradiol dioxygenases) were revealed by in-depth molecular analyses [PCR-denaturing gradient gel electrophoresis and hybridization (SBH and microarray)]. Our results suggest that, despite the low abundance of PH-degrading genes and plasmids in the environmental samples, the original bacterial composition of the mangrove microniches determined the structural and functional diversity of the PHDC enriched.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

Sediment contamination with petroleum hydrocarbons (PH) is especially dangerous for mangrove forests, because low-molecular-weight aromatic hydrocarbons (e.g. BTEX, naphthalene and phenanthrene) can be phytotoxic and affect plants at all stages of growth (Proffitt et al., 1995; Hoff, 2002; Kummerová & Kmentová, 2004). Curiously, despite their clear symptoms of stress (death of mangrove saplings, smaller trees and the loss of forest cover), mangrove trees are still often found in urban areas under chronic exposure to PH contamination (Proffitt et al., 1995; Gomes et al., 2007; Benson & Essien, 2009). The survival of these plants may be due to their inherent physiological resistance, but could also be the result of beneficial interactions with the associated microbiota. Specifically, it had been documented that microbial PH degraders can be enriched in the plant rhizosphere due to the release of root exudates and can either contribute to soil detoxification surrounding the root or inside of the plant (Kuiper et al., 2001; Germaine et al., 2009). Surprisingly, despite the well-known ability of inland plants to promote bioremediation of contaminated soil, no studies have hitherto performed an in-depth analysis of the catabolic potential of microorganisms associated with mangrove roots for PH degradation. Regarding neotropical coastal areas, mangroves are native trees that can be planted in those areas, potentially acting as reservoirs of PH-degrading microbial guilds for bioremediation purposes.

It is assumed that horizontal gene transfer by mobile genetic elements plays an important role in the acclimation of bacterial populations to environmental contamination (Bale et al., 1988; Rasmussen & Sørensen, 1998; Wilson et al., 2003). However, in spite of its potential ecological importance, a limited number of studies have addressed plasmid diversity in coastal environments (Sobecky, 1999; Beeson et al., 2002), and no reports exist thus far that specifically focus on mangrove ecosystems. Moreover, although it is well known that the degradation of environmental pollutants is enhanced in plant rhizospheres, the diversity and heterogeneity of plasmids and the functional genes involved in this process remain almost completely unknown in natural mangrove settings.

The in vitro enrichment of functional guilds has been used commonly for the isolation of hydrocarbon degraders from terrestrial, marine and freshwater sources (MacCormack & Fraile, 1997; Head, 1998; Yakimov et al., 2003). Often, the resulting enrichments allow the acquisition of specific information, such as identification of microbial degraders and metabolic pathways. For example, due to the difficulties associated with the culturing of environmental microorganisms, the utilization of DNA from enrichment cultures can allow the study of several new microbial traits and improve access to new potential sources of genes of biotechnological interest (Entcheva et al., 2001). Furthermore, enrichments can be used as environmental inoculants to speed up natural biological processes (bioaugmentation) (Venosa et al., 1996).

The utilization of plant inoculants has also proven to be a reliable approach to promote plant growth and phytoremediation of PH-polluted environments (Kuiper et al., 2001; Germaine et al., 2009). In fact, the plant–microorganism interactions have become an important topic in restoration ecology (Harris, 2009). The development of root inoculants with the ability to colonize and to degrade phytotoxic PH compounds in the root system (rhizoremediation) might contribute to the development of novel strategies for environmental recovery, which may couple both reforestation and remediation of PH-impacted environments. A primary step to achieving this promise relies on the isolation and identification of the structural and functional components involved in the degradation of PH in the rhizosphere.

In this study, bulk sediment and rhizosphere (microniches) of healthy plants of three different species (Avicenia schaueriana, Laguncularia racemosa and Rhizophora mangle) were sampled in a mangrove site exposed to chronic levels of PH contamination. The microbial communities retrieved from the samples were then used in serial batch enrichment subculturing in the presence of petroleum to select for mangrove microbial guilds involved in the process of PH degradation. A range of molecular tools were used to investigate the influence of the original composition of the microbial communities of different mangrove microniches on the structural and functional diversity of enriched PH-degrading consortia (PHDC). The PHDC obtained will be used for future mangrove reforestation projects in PH-polluted sites.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

Growth of mangrove plants and sampling

The sampling approach of this study was designed to ensure sampling access to rhizospheres from A. schaueriana, L. racemosa and R. mangle at similar developmental stages. Briefly, mature mangrove propagules of R. mangle, A. schaueriana and L. racemosa were collected in mangrove forests located in Guanabara Bay (Rio de Janeiro, Brazil) and grown in a mangrove nursery. Sixteen plant saplings (∼75 days old) from each mangrove species were replanted in a mangrove forest exposed to chronic levels of oil hydrocarbon contamination close to the petrochemical complex of Duque de Caxias (22°44′08″S/43°13′55″W) (Rio de Janeiro, Brazil). The sampling site characteristics were described previously (Gomes et al., 2007, 2008).

The saplings were planted randomly in high intertidal zones with a distance of at least 1 m between each other. Bulk sediment and rhizosphere samples were taken 28 days after planting. Three composite replicates of bulk sediment (∼20 cm of top sediment with 4 cm diameter) and roots of individual plants (three replicates) of each species were sampled. A spatula was used to remove the sediment that could be easily detached from the roots. Only the remaining sediment adhering to the plant root system was considered as the rhizosphere fraction. Each rhizosphere sample consisted of the total root system, which was cut and thoroughly mixed. Microbial cells were detached from rhizosphere and bulk sediment samples (5 g) as described previously in Gomes et al. (2007). The resulting microbial suspensions were used in the batch enrichment of PHDC.

Enrichment of PHDC

The enrichment of PHDC was performed by means of a serial batch culture approach. The first enrichment round consisted of inoculation with 5 mL of the microbial community suspensions obtained from rhizosphere and bulk sediment samples (as described above) in Erlenmeyer flasks containing 45 mL mineral salt medium (MSM) (Margesin & Schinner, 1997) supplemented with light petroleum (0.1% v/v) (cordially provided by Petrobras S.A.) as the sole added carbon source. Before medium amendment, the petroleum was sterilized in an oven at 180 °C for 2 h. Flasks without inoculation were used as a control. The flasks were closed with nonabsorbent cotton wool and incubated at 27 °C in the dark under constant shaking for 5 days. Enrichments were continued for two more rounds by serial subculturing in the MSM medium amended with petroleum as described above, using a 10% inoculum from the previous culture. Aliquots of each enriched PHDC were stored with glycerol at −80 °C for follow-up studies. The concentration of total PH was determined in extracts from the batch cultures after the last enrichment round for each replicate by gas chromatography (GC), as described previously in the Environmental Protection Agency ( standard method 8015. anova was applied to determine whether there was significant removal of total PH by PHDC.

DNA extraction

After the third enrichment round, 1 mL from each enriched consortium was transferred to a Lysing Matrix E tube (Q Biogene) and the microbial cells were pelletized by centrifugation. Total community DNA was extracted using the BIO101 DNA extraction kit (Q Biogene) according to the manufacturer's recommendations. Mechanical lysis was achieved using the FastPrep FP120 bead-beating system (Q Biogene) 2 × for 30 s at a maximum vertical velocity of 5.5 m s−1.

PCR amplification of 16S rRNA gene fragments and denaturing gradient gel electrophoresis (DGGE)

Amplified 16S rRNA gene fragments suitable for DGGE fingerprint analyses were obtained with bacterial DGGE primers F984-GC and R1378 (∼473 bp) according to Heuer et al. (1997). A nested-PCR approach (25 thermal cycles) was also applied for the amplification of 16S rRNA genes of Alphaproteobacteria, Betaproteobacteria and Pseudomonas groups as described previously (Heuer et al., 1997; Gomes et al., 2001; Milling et al., 2004). The DGGE of the amplified 16S rRNA gene fragments was performed using the INGENY PhorU System (INGENY, Goes, the Netherlands). Gels were prepared with a double gradient of 30–65% denaturants (100% denaturant: 7 M urea and 40% formamide) and 6–9% acrylamide. The electrophoresis run was carried out in 1 × Tris-acetate–EDTA buffer at 58 °C at a constant voltage of 240 V for 20 h. The gels were silver stained according to Heuer et al. (2001).

PCR amplification of naphthalene dioxygenase (ndo) gene fragments and ndo DGGE analyses

The detection of genes encoding enzymes involved in the degradation of low-molecular-weight polycyclic aromatic hydrocarbon (PAH) in the enrichment cultures was achieved by a nested-PCR approach targeting ndo genes using the method developed by Gomes et al. (2007). This PCR system targets environmentally relevant ndo genes belonging to the main clade of group III according to the classification system proposed by Nam et al. (2001). Amplified GC-clamped ndo gene fragments obtained after the second PCR round were applied to a double-gradient DGGE as described above for bacterial fingerprints.

Detection of broad host range (BHR) plasmids and ndo genes by PCR from total community DNA and Southern blot hybridization (SBH)

Before SBH, a PCR-based approach was used for the amplification of BHR plasmid-specific sequences and ndo genes from DNA extracted from environmental samples and from the enrichment consortia. The PCR amplification of ndo genes (first step ndo PCR) was performed as described previously (Gomes et al., 2007). The amplification of plasmid-specific sequences of IncP-1 α and β (trfA2 gene), IncP-7 (rep gene) and IncP-9 (ori-rep genes) was carried out as described previously (Götz et al., 1996; Krasowiak et al., 2002; Izmalkova et al., 2005). The amplicons from ndo and BHR plasmid PCR were Southern blotted onto HYBOND N nylon membranes (Amersham Pharmacia Biotech) according to Sambrook et al. (1989). The probes for ndo genes were generated from PCR products of nahAc from Pseudomonas putida KT2442 (pNF142) (Gomes et al., 2005), phnAc from a cloned phnAc gene fragment (Gomes et al., 2005) and from the cloned nagAc gene fragment 4NDO-S3 (Gomes et al., 2007). The probes for BHR plasmids were PCR-generated as described previously for IncP-1 α and β, IncP-7 and IncP-9 (Götz et al., 1996; Krasowiak et al., 2002; Izmalkova et al., 2005). Hybridization was performed under conditions of medium stringency following the protocol published by Fulthorpe et al. (1995). Hybridization of DIG-labelled probes was detected using a DIG luminescent detection kit (Roche) as specified by the manufacturer and exposed to an X-ray film (Roche).

Comparative analyses of enriched PHDC

The DGGE gels were scanned transmissively and the digitalized DGGE profiles were analyzed using the software package gelcompar 4.0 (Applied Maths, Sint-Martens-Latem, Belgium) as described by Smalla et al. (2001). The sets used for band detection were 5% minimal profiling (area along the densitometric curve) and 0.5% minimal area. The positioning and quantification of bands were carried out by setting tolerance and optimization at eight points, i.e. 0.8%. The band positions and their corresponding intensities from each PHDC were exported to Excel files and the band surface was converted to relative intensity by dividing its surface by the sum of all band surfaces in a lane. Bray–Curtis similarities were calculated based on the band position and intensity. The matrices of similarities were then used for multivariate analyses of DGGE profiles using analysis of similarities (anosim) with the primer 5 software package (Primer-E Ltd, Plymouth, UK). The anosim was used to test whether communities differed significantly. The R statistic in anosim ranges from 0 to 1, with higher values indicating greater variation in the composition among the samples (Clarke, 1993). Moreover, the extent to which the variation in the presence and relative abundance of DGGE bands is explained by the origin of PHDC (i.e. sediment or rhizosphere samples) was addressed. To this end, ordination of PHDC fingerprinted by DGGE and their corresponding sources of isolation was performed following the procedures of Costa et al. (2006). Briefly, detrending correspondence analysis and detrending canonical correspondence analysis were carried out to estimate the gradient lengths of band relative intensity datasets. Based on these assessments, linear species response models – i.e., principal components analysis (PCA) and redundancy analysis (RDA) – were deemed to better fit the dataset and therefore used in unconstrained (PCA) and constrained (RDA) ordination analyses (Ramette, 2007). Focus on sample distances was used with further default parameters, as implemented in the software package canoco for windows 4.5 (Microcomputer Power, Ithaca, NY).

Sequence analyses

The most dominant bands selected from bacterial DGGE profiles were excised, cloned, screened and sequenced as described previously (Gomes et al., 2008). One clone per band carrying the right insert with the DGGE mobility of each selected band was sequenced. The partial 16S rRNA gene sequences obtained were classified according to the Naive Bayesian ribosomal RNA Classifier (Version 1.0) of the Ribosomal Database Project II ( and compared with sequences available in the GenBank database using blast-n ( Sequences were subjected to the Chimera Check program from RDP and deposited in the GenBank database under the accession numbers (HM003918HM003925). The ndo gene sequence fragments for cloning were screened as described previously (Gomes et al., 2007). Clones containing inserts that shared the electrophoretic mobility with dominant DGGE bands were selected for sequencing. blast-n was used for similarity searches for ndo gene sequences (∼740 bp). Sequences of cloned ndo gene fragments were aligned with their closest relatives. A phylogenetic tree was then inferred using the neighbor-joining algorithm applied to a matrix of evolutionary distances calculated with the Maximum Composite Likelihood method and bootstrapping analysis using the Molecular Evolutionary Genetics Analysis (mega4) integrated software. The nonredundant ndo gene sequences obtained were deposited in the GenBank database under the accession numbers (HM003909HM003917).

Microarray analyses

The microarray designed in this study was composed of probes targeting 16S rRNA gene sequences (137 probes) representing distinct bacterial groups across a variety of phyla (including known degraders of aromatic hydrocarbons) and gene members of the main evolutionary clusters (subfamilies) of extradiol dioxygenases type I (EXDO, vicinal chelate superfamily) (197 probes) and intradiol dioxygenases (INDO) (257 probes) involved in aromatic degradation. The probes targeting 16S rRNA genes were designed, according to the bacterial taxonomy of the Ribosomal Database Project (, on a selection of sequences from type strains representing 19 phyla: Chloroflexi, Thermomicrobia, Nitrospira, Deferribacteres, Cyanobacteria, Chlorobi, Actinobacteria, Planctomycetes, Chlamydiae, Spirochaetes, Fibrobacteres, Acidobacteria, Bacteroidetes, Fusobacteria, Verrucomicrobia, Dictyoglomus, Gemmatimonadetes, Firmicutes and Proteobacteria. In the cases of Firmicutes, sequences from type strains of the different classes inside the families of Clostridia, Mollicutes and Bacilli were included. For Proteobacteria, we included expanded representation of type strain sequences of the following orders (in parentheses) inside Classes of Alphaproteobacteria (Rhodospirillales, Rickettsiales, Rhodobacterales, Sphingomonadales, Caulobacterales, Rhizobiales, Parvularculales); Betaproteobacteria (Burkholderiales, Hydrogenophilales, Methylophilales, Neisseriales, Nitrosomonadales, Rhodocyclales); Gammaproteobacteria (Acidithiobacillales, Thiotrichales, Chromatiales, Xanthomonadales, Cardiobacteriales, Legionellales, Methylococcales, Oceanospirillales, Pseudomonadales, Aeromonadales, Alteromonadales, Vibrionales, Enterobacteriales, Pasteurellales); Deltaproteobacteria (Desulfurellales, Desulfovibrionales, Desulfobacterales, Desulfuromonales, Syntrophobacterales, Myxococcales); and Epsilonproteobacteria (Campylobacterales).

The probes against catabolic genes were designed on two custom CDS databases for EXDO type I and INDO gene families, which were built based on protein sequences from reference enzymes, representing different evolutionary branches described for INDO (Eulberg et al., 1998) and for EXDO type I (Eltis & Bolin, 1996). These sequences were used as seeds for blastp searches in order to upgrade the sequence diversity currently found in the databases. By collecting the CDS corresponding to the related proteins of blastp results and excluding the redundant representation of the same or near-identical sequences (>99% similarity), the resulting databases had been used to design 50-oligomer probes (Chou, 2010) able to specifically detect, under the hybridization conditions assayed, each of their sequence targets with a similarity higher than 84%. According to megablast searches (GenBank nonredundant nucleotide database), all the probes designed have matches with highly related genes (95% similarity over the complete alignment of the target and the related gene CDS or 16S rRNA gene). The full list of the genes targeted (including the GenBank number, abbreviation of the bacterial genus or species where the gene was found, strain name, gene name abbreviation) conforming to the final dataset used for probe design can be found in the Supporting Information, Fig. S1. These probes arrayed are able to collectively detect all the various types of evolutionary branches across the gene families or in the case of 16S rRNA gene, taxonomically highly related type strain sequences.

About 20 ng of DNA of the third PHDC replicate (Avi-c, Lag-c, Rhi-c and Sed-c) of each treatment was amplified by phi29 DNA polymerase using the Ultrafast Repli-G kit (Qiagen) according to the specifications of the manufacturer and the appropriate test of controls. After digestion with DNase turbo (Ambion), fragments were biotinylated using dUTP-biotin (Roche) and concentrated to a final volume of 20 μL in a vacuum centrifuge. Afterwards, the volume was adjusted to 100 μL with formamide buffer and hybridization was performed at 55 °C for 18 h. Slides were stained with streptavidine-Cy3 in TNB buffer at room temperature and washed with SSC buffer 0.1 × SDS 0.2% at 48 °C for 10 min. Before scanning, slides were rinsed with SSC buffer 0.1 × at room temperature. Scanning was performed in an Agilent system according to the manufacturer's instructions. Raw data intensities were normalized against the background using the formula (spot intensity−intensity background)/intensity background. Values for signals considered above the background as threshold were considered as positives. Controls of type genomes for which hybridization profiles were predicted as well as specific target gene copies vs. probes intensities were used to define the intensity threshold at which the experimental results fully matched the expected profiles (R. Vilchez-Vargas, unpublished data). All the microarray results presented are the average intensities of triplicate measurements per probe.

Results and discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

Enrichment of PHDC

The quantitative and qualitative aspects of root exudates released by different plant species and at different plant growth stages will affect root colonization by responding bacteria and the synergistic interactions among these (Hedge & Fletcher, 1996; Jaeger et al., 1999). Therefore, the sampling approach of this study was designed to ensure rhizosphere samples from different mangrove plant species at similar growth stages.

The GC analysis of A. schaueriana (129.6 ± 62 mg L−1) and L. racemosa (134.6 ± 35.2 mg L−1) rhizosphere PHDC (third transfer) showed a significant reduction in the total PH concentration in comparison with the control flasks (301 ± 109.3 mg L−1) (P<0.05). The final concentration of the total PH in the flasks containing R. mangle (166.1 ± 13.1 mg L−1) and bulk sediment (235.8 ± 38.1 mg L−1) PHDC were not significantly different from the control flasks (P>0.05). Although not significant, the values for R. mangle and bulk sediment PHDC still show a trend toward an improved PH degradation. The petroleum volatile compounds (several n-alkanes and low-molecular-weight aromatic molecules) belong to the petroleum light fraction and are known to be more readily degraded than high-molecular-weight PH (Venosa et al., 1996). The lack of significant differences between the control flasks and R. mangle and bulk sediment PHDC may be explained by the dominance of microbial guilds more specialized in the degradation of volatile PH compounds in these two cultures. Obviously, petroleum volatile compounds were also lost due to volatilization in the control flasks during the 5 days of the incubation period. However, this loss was not measured in this work. In contrast, flasks inoculated with A. schaueriana and L. racemosa rhizosphere PHDC showed a stronger decrease in the final concentration of the total PH. The higher efficiency of PH removal by A. schaueriana and L. racemosa rhizosphere PHDC suggests that the relative abundance and activity of specific PH-degrading guilds in these two enrichments were different from R. mangle and bulk sediment enrichments. Studies on plant–microorganisms interactions have shown that the influence of root exudation on the composition of microorganisms colonizing the rhizosphere is plant species specific and can selectively enhance specific microbial guilds (Grayston et al., 1998; Neumann & Römheld, 2001). In order to demonstrate that the enrichments display a microniche-dependent structural and functional diversity of PHDC, a thorough DNA-based analysis was performed.

DGGE analyses of PHDC

Bacteria and taxon-specific DGGE analyses were used to compare the relative abundance of PHDC enriched from rhizosphere and bulk sediment microbial communities (see Fig. S2). While Bacteria and Alphaproteobacteria DGGE fingerprint analyses revealed complex profiles, a rather low complexity of DGGE ribotypes was detected for Betaproteobacteria and Pseudomonas. Differences in the complexity (number of bands) of Bacteria fingerprints were also observed between PHDC enrichments. With the exception of one replicate of R. mangle PHDC, the Bacteria fingerprints of rhizosphere PHDC showed a higher number of bands compared with PHDC from bulk sediment, suggesting that a larger number of populations adapted to growth on PH.

In this study, anosim of bacterial DGGE profiles were used to test whether the differences observed between PHDC communities were significant (R) (Table 1). Values of R>0.75 are considered well separated, values >0.5 as moderately separated and values <0.25 as poorly separated (Clarke, 1993; Ramette, 2007). With the exception of R. mangle PHDC, the statistical analysis of Bacteria community fingerprints revealed significant separation between rhizosphere PHDC of different plant species and between rhizosphere and bulk sediment PHDC. The anosim also indicated significant separation of Alphaproteobacteria communities from rhizosphere and bulk sediment PHDC. However, the comparison of rhizosphere PHDC only showed significant separation between A. schaueriana vs. R. mangle (R=0.70). The analyses of Betaproteobacteria and Pseudomonas communities did not show significant differences between rhizosphere PHDC from different plant species nor between rhizosphere and bulk sediment PHDC. In general, the statistical analyses indicated that differences in the bacterial community structure of each PHDC were best depicted by Bacteria fingerprint analyses.

Table 1. anosim statistics of Bray–Curtis similarity measures (R) of rhizosphere and bulk sediment PHDC
PHDCBacteria (R)Alphaproteobacteria (R)Betaproteobacteria (R)Pseudomonas spp. (R)
  • *


  • Bulk sediment.

  • Avi, Avicenia schaueriana.

  • §

    Lag, Laguncularia racemosa.

  • Rhi, Rhizophora mangle.

Rhiz* vs. Sed
Rhiz vs. Rhiz
 Avi vs. Lag10.1400.40
 Lag vs. Rhi0.630.2200
 Rhi vs. Avi0.110.7000.14

Further multivariate statistics performed using ordination techniques such as PCA and RDA supported the overall trends observed via anosim. The PCA ordination biplot of PHDC bacterial DGGE fingerprints and their underlying environmental samples (Fig. 1) illustrate the clear differentiation between bulk sediment, A. schaueriana and L. racemosa PHDC across the ordination space. Taken together, these results indicate for the first time that the inoculum from different mangrove microniches determined the structural composition of the PHDC enrichments. Similar trends have been observed in previous studies based on terrestrial plants, which have shown that root exudates can induce plant species-specific changes on the structural diversity of rhizosphere bacterial communities (Gomes et al., 2001; Smalla et al., 2001; Costa et al., 2006).


Figure 1.  PCA ordination biplot of PHDC of bacterial PCR-DGGE fingerprints and their corresponding sources of isolation. Sed, bulk sediment; Avi, Avicenia schaueriana; Lag, Laguncularia racemosa; Rhi, Rhizophora mangle.

Download figure to PowerPoint

Sequence analysis of dominant DGGE bands

Selected dominant bands from Bacteria and taxon-specific DGGE fingerprints of PHDC were chosen for cloning and subsequent sequencing. The sequence analysis of all selected DGGE ribotypes revealed bacterial populations, which were assigned to three orders: Pseudomonadales, Rhizobiales and Burkholderiales (Table 2). The sequences affiliated to the orders Pseudomonadales and Burkholderiales were associated with the most abundant populations detected in the PHDC enrichments. These sequences were often closely related to known PH degraders (Table 2). Sequence analyses of bands excised from Bacteria profiles revealed the dominance of Acinetobacter venetianus in nearly all PHDC, with the only exception of L. racemosa PHDC. In contrast, the results indicated that Acinetobacter baumannii was only enriched in L. racemosa PHDC. Members of the genus Acinetobacter are known for their ability to catabolize a wide range of carbon sources and for their importance in the process of emulsification and degradation of PH (Sar & Rosenberg, 1983; Pleshakova et al., 2001; Koren et al., 2003). The two sequences derived from the Betaproteobacteria PHDC fingerprints were assigned with high confidence to Comamonas testosteroni and Burkholderia sp., bands 1bet and 2bet (see Fig. S2c, Table 2), respectively.

Table 2.   Results of partial 16S rRNA gene sequence analysis and tentative affiliations of selected DGGE ribotypes
DGGE band type*Accession no.Closest phylogenetic relative
blast-n identity%Accession no.
  • *

    Codes refer to the bands shown in Fig. S1.

  • GenBank sequence accession numbers of the respective clone.

  • GenBank sequence accession number of the most closely related bacterial sequence.

1bacHM003919Acinetobacter venetianus100DQ912805
2bacHM003923Acinetobacter baumannii100FN563424
1alpHM003918Bosea sp.100DQ104981
2alpHM003922Uncultured bacterium99EU234233
1betHM003920Comamonas testosteroni99GQ259481
2betHM003924Burkholderia sp.99AB299596
1pseHM003921Pseudomonas aeruginosa100GU447238
2pseHM003925Pseudomonas putida99GU396284

The 16S rRNA gene sequence analyses of bands excised from Pseudomonas profiles revealed the dominance of populations affiliated with Pseudomonas aeruginosa and P. putida (Table 2). These species are known versatile degraders of PH (Van Hamme et al., 2003; Wongsa et al., 2004). We have shown previously that P. aeruginosa populations were enhanced in mangrove sediments contaminated with PH (Gomes et al., 2008). Brito et al. (2006) also reported the isolation of a P. aeruginosa strain from petroleum-contaminated mangrove sediments able to degrade fluoranthene and octadecane. Several environmental and clinical isolates of P. aeruginosa were also found to contain alk genes (Smits et al., 2003), which can be assumed to belong to the core genome of this species (Palleroni et al., 2010). The alk genes code for alkane hydroxylase, which is an important enzyme involved in the process of environmental degradation of petrogenic n-alkanes. These data suggest that P. aeruginosa might have a potential ecological role in the process of PH detoxification in mangrove sediments.

In contrast to the sequence analyses of bands retrieved from Bacteria, Betaproteobacteria and Pseudomonas fingerprints, which revealed typical PAH degraders in the enrichment consortia, the dominant bands retrieved from Alphaproteobacteria profiles (see Fig. S2b, 1alp and 2alp) were assigned to phylogenetic groups belonging to the Rhizobiales order. This order comprises several plant symbionts and nitrogen-fixing bacteria. Only recently have some members of this group been reported as degraders of aromatic hydrocarbon compounds (Baek et al., 2003). Even though their potential role in the process of PH degradation in the plant root system remains largely unknown, a recent survey of aromatic degrading properties encoded in the genomes of >900 sequenced bacteria indicated strains of the Rhizobiales order to be metabolically highly versatile (Pérez-Pantoja et al., 2010). The sequence from the DGGE ribotype 1alp was similar to Bosea sp., a non-nitrogen-fixing Rhizobiales able to generate energy from the oxidation of reduced sulfur compounds (chemolithoheterotroph) (Das et al., 1996), which was previously shown to be able to degrade PAH (Seo et al., 2007). The closest relative of the cloned sequence retrieved from band 2alp could not be assigned with good confidence to any family and its closest relative in the GenBank was an uncultured bacterium clone (Table 2).

PCR-SBH analyses

The PCR-SBH analysis of total community DNA extracted from environmental samples revealed positive signals for only a few replicates (Table 3), suggesting that ndo genotypes (nag, phn and nah) and BHR plasmids (IncP-1α, IncP-1β, IncP-7 and IncP-9) were in low abundance in the rhizosphere and bulk sediments analyzed. Nevertheless, the different plasmid groups and ndo genes were enriched from these same samples after PHDC enrichments (Table 3). These results indicate that several biological components involved in the process of PH removal are initially present below the detection limit of the technique applied in both mangrove rhizosphere and bulk sediments. Despite the differences in the hybridization signals, SBH analysis showed that the ndo genotypes and BHR plasmids were strongly enriched in the PHDC. Populations carrying plasmid groups belonging to IncP-9, IncP-1α and IncP-1β were enriched in most of the PHDC (Table 3). The IncP-9 plasmid is a known group of self-transmissible plasmids responsible for horizontal transfer of catabolic genes encoding PH (e.g. xylene, toluene and PAH) catabolism (Dennis, 2005; Sota et al., 2006). Probably, the enrichment of this group of plasmids in nearly all PHDC occurred due to the selection of plasmid-encoded PH catabolic traits. Curiously, SBH signals for IncP-1α and β were stronger in A. schaueriana, L. racemosa and bulk sediment PHDC than in R. mangle PHDC. The IncP-1 group encompasses the most promiscuous self-transmissible plasmids known to date, which often carry genes coding for the degradation of man-made chloroaromatic compounds and resistance against antibiotics and metals (Dennis, 2005; Smalla et al., 2006; Schlüter et al., 2007). The IncP-1 group may have been spread through the PHDC bacterial population due to the growth of a plasmid-containing population or horizontal spread as different studies have shown that IncP-1 plasmids can also be efficiently transferred to bacterial populations even in the absence of selection pressure (Pukall et al., 1996; Fox et al., 2008). Surprisingly, plasmids belonging to IncP-7 were poorly enhanced in the PHDC. Plasmids belonging to this group are known to possess complex genetic rearrangements that can help the host cell to adapt to environmental changes (Dennis, 2005). This group is also deemed to include important mobile elements for spreading genes encoding PH degradation.

Table 3.   PCR SBH analyses of plasmid groups and naphthalene dioxygenase genes of mangrove environmental samples (MES) (rhizospheres and sediment) and their respective enriched PHDC
MESA. schauerianaL. racemosaR. mangleSediment
  1. (+), weak hybridization (when faint bands were detected);+, hybridization (dark and sharp bands);++, strong hybridization (dark oversized band area).

IncP1β  +    + +  
nagAc    +    +  
IncP-1α++(+)+++++++(+) (+)++(+)++
IncP1β+(+)+++++++ (+)++++++
IncP-7+(+)     (+)  ++ 
nahAc(+)(+)(+)(+)+ (+)+  (+)(+)
phnAc++++++   +++    

The SBH analyses of ndo genes detected the enrichment of nahAc, nagAc and phnAc genes in the PHDC (Table 3). However, while nahAc and phnAc-like genes were enhanced in few PHDC, the results indicated the enhancement of genotypes closely related to nag genes in all cultures. In agreement with these results, the ndo DGGE analyses (Fig. 2) revealed the dominance of one band (ndo DGGE band type 1) in nearly all PHDC. The phylogenetic analysis of cloned sequences matching these bands showed a close phylogenetic relationship with the protein encoded by the pahAc gene of C. testosteroni H (Fig. 3). The pah gene of C. testosteroni H has a high sequence similarity to the nag operon (Moser & Stahl, 2001). The abundance of populations carrying the nagAc-like genes was previously shown to be enhanced in a mangrove sample from sites under chronic exposure to PH (Gomes et al., 2007). Bulk sediment samples from the same sampling site were used in the PH enrichments performed in this work. The nagAc genotype has been detected in PH-contaminated sites in different geographic locations all over the world (Widada et al., 2002; Jeon et al., 2003; Gomes et al., 2007). A study by Dionisi et al. (2004) has shown that indigenous bacteria carrying nagAc-like genes play an important ecological role in the process of in situ degradation of PAH. Previous studies have also indicated that phnAc genotypes similar to those found in Burkholderia sp. RP007 are ubiquitous genes often detected in PH-contaminated soils (Laurie & Lloyd-Jones, 1999; Lloyd-Jones et al., 1999; Wilson et al., 2003; Gomes et al., 2005). Both molecular approaches used in this study for ndo gene analyses (SBH and DGGE) revealed that phnAc-like genes related to Burkholderia sp. RP007 were abundant only in replicates of A. schaueriana PHDC and two replicates of R. mangle PHDC. Burkholderia sp. RP007 was isolated previously from contaminated soil samples in New Zealand (Laurie & Lloyd-Jones, 1999). Our results suggest that while nag genotypes might be ubiquitously distributed in bulk and rhizosphere sediments, bacterial guilds carrying phn-like genes may have a more specific association with the rhizospheres of A. schaueriana and R. mangle.


Figure 2.  DGGE fingerprints of ndo gene fragments amplified from DNA templates extracted from PHDC. Sed, Bulk sediment; Avi, Avicenia schaueriana; Lag, Laguncularia racemosa; Rhi, Rhizophora mangle. The bands x and y are silver-stained single-strand DNA. The band positions indicated in the gel correspond to the melting behavior of selected representative ndo gene clones that matched dominant genotypes. From top to bottom, ndo gene fragments used as markers (M): phnAc (environmental clone), nahAc (Pseudomonas putida KT2442 – pNF142) (Gomes et al., 2005) and nahAc (Pseudomonas sp. ARS 10) (I. Kosheleva, unpublished data).

Download figure to PowerPoint


Figure 3.  Phylogenetic relationships of DNA sequences of the large α-subunits of naphthalene dioxygenase genes. The sequences were aligned with related sequences retrieved from GenBank. The narAa gene from a PAH-degrading gram-positive bacterium (Rhodococcus sp. NCIMB12038) was used as an outgroup. The numbers on the branches indicate the percent of bootstrap values.

Download figure to PowerPoint

Diversity and affiliation of 16S rRNA and aromatic ring-cleavage genes based on microarray analyses

A microarray was specifically designed (unpublished) to determine profiles of 16S rRNA genes and aromatic ring-cleavage (meta- and ortho-cleavage pathways) encoding genes. The arrays can detect all the main branches described for the families targeted, and as such, can detect a range of highly divergent branches in the family, but, nevertheless, rely on previous sequence knowledge and each probe is also able to detect highly similar genes (>90%) to the sequence targeted. With the exception of L. racemosa PHDC, patterns of positive signals above the background threshold were obtained for DNA samples extracted from PHDC. Microarray analysis of L. racemosa PHDC showed hybridization problems that could not be solved and only two taxon-specific probes (Bacillus cereus ATCC27877 Z84581 S000005826 and Pseudomonas psychrophila E-3 AB041885 S000002852) had signals above the background threshold (data not shown). It was found that the majority of taxa microarray signals (see Table S1) were detected by probes targeting Pseudomonadales 16S rRNA genes, in accordance with DGGE results (see Fig. S2 and Table 2). However, some dominant populations detected by PCR-DGGE were not detected by microarray analysis. Such a difference may be due to differences in the sensitivity of both techniques and most probably due to the fact that the microarray designed in this study has a limited number of taxon-specific (16S rRNA gene) probes designed based on species type strains.

Probes targeting EXDO and INDO detected genes encoding various enzymes responsible for the ortho- and meta-cleavage of catechols and chlorocatechols (catechol 1,2 dioxygenases, C12O) (Tables 4 and 5). The INDO family comprises different evolutionary branches, such as catechol 1,2-dioxygenases of Proteobacteria, those of Actinobacteria, chlorocatechol 1,2-dioxygenases, hydroxyquinol dioxygenases and the α and β subunits of protocatechuate 1,2-dioxygenases (Pérez-Pantoja et al., 2010). Prominent signals were observed, indicating the abundance of catechol 1,2-dioxygenase and protocatechuate dioxygenases encoding genes of Burkholderia and Pseudomonas spp., genes typically belonging to the core genome of members of these genera (Pérez-Pantoja et al., 2010). These results are in accordance with the observation of a high abundance of such species in the samples analyzed. Also, genes encoding catechol 1,2-dioxygenases of Actinobacteria could be observed in all samples. This group of bacteria has not been targeted in PCR assays, but may be important and must be tracked in detail in future studies of the ecosystem under analysis. Probes specifically designed for chlorocatechol 1,2-dioxygenase encoding genes, such as the ones targeting Pseudomonas sp. P51 (P27098), Delftia acidovorans P4a (AAC35836) or P. aeruginosa JB2 (AAC69474) only showed hybridization signals for A. schaueriana enrichment. The reason for the specific abundance of such genes remains to be elucidated. Hydroxyquinol dioxygenase encoding genes were observed in high abundance in rhizosphere PHDC, possibly due to the enrichment of organisms harboring such genes by root exudates.

Table 4.   Positive hybridization signals of DNA extracted from PHDC with the microarray probes against 257 CDS of protein members representing all the main phylogenetic branches of the INDO family (see Fig. S2)
INDO gene target*A. schauerianaR. mangleSediment
  • DNA extracted from PHDC; Avicenia schaueriana; Rhizophora mangle and bulk sediment.

  • *

    INDO gene targets are described with the genus or species name and bacterial strain identification where the aromatic ring-metacleavage protein member of the INDO family has been found, followed by the gene abbreviation and its GenBank accession number.

  • Signal intensities detected in DNA extracted from the PHDC of bulk sediment (Sediment), A. schaueriana and R. mangle above the significant relative intensities are shown as the triplicate mean values with the corresponding standard deviations.

  • C12O, catA or catB: catechol 1,2-dioxygenase.

  • pPC34DO, pca, or pcaH: putative (predicted from annotations of genome projects) protocatechuate 3,4-dioxygenase.

Acinetobacter lwoffii K24 catA U77658 0.68 ± 0.18 
Arthrobacter sp. BA-5-17 catA BAD11154 0.99 ± 0.80 
Aspergillus fumigatus Af293 pcatA AAHF010000081.17 ± 1.09  
Brevibacterium linens BL2 pPC34DO NZAAGP010000020.85 ± 0.300.92 ± 0.821.75 ± 1.22
Brucella suis 1330 pcaC AE0142920.61 ± 0.031.16 ± 1.270.54 ± 0.48
Burkholderia cenocepacia AU pPC34DO2 YP_6240540.86 ± 0.321.23 ± 0.63 
Burkholderia cenocepacia AU10 pPC34DO NZAAHI010000200.71 ± 0.241.41 ± 1.590.55 ± 0.26
Burkholderia mallei 10399 pPC34Do NZAAHN010000250.56 ± 0.14  
Burkholderia mallei ATCC23344 pcaH NC0063490.84 ± 0.430.95 ± 0.540.61 ± 0.38
Burkholderia mallei NCTC10247 pPC34DO NZAAHP010000431.67 ± 0.28  
Burkholderia pseudomallei 171 pPC34DO NZAAHS010000300.60 ± 0.390.61 ± 0.33 
Burkholderia pseudomallei 668 pPC34DO NZAAHU010000012.72 ± 1.682.90 ± 1.522.38 ± 1.12
Burkholderia pseudomallei 668 pPC34DO NZAAHU010000320.88 ± 0.520.76 ± 1.32 
Burkholderia pseudomallei K96243 pca NC0063510.70 ± 0.470.72 ± 0.400.50 ± 0.36
Burkholderia pseudomallei NCTC 10229 pPC34DO YP_0010240700.66 ± 0.381.05 ± 1.19 
Burkholderia pseudomallei S13 pPC34DO NZAAHW010000911.04 ± 0.250.66 ± 0.40 
Burkholderia sp. c12o TH2 BAC167690.54 ± 0.350.56 ± 0.44 
Burkholderia vietnamiensis G4 NZAAEH02000012  0.59 ± 0.24
Cupriavidus necator CH34 pC12O YP_587012  0.50 ± 0.32
Cupriavidus necator JMP134 catA YP_2983350.57 ± 0.27  
Delftia acidovorans P4a chloro-C12O AAC358360.56 ± 0.29  
Jannaschia sp. pPC34DO YP_5121051.37 ± 0.740.81 ± 0.680.83 ± 0.82
Nocardia sp. H171 catA AY613438 0.80 ± 0.850.79 ± 0.68
Polaromonas sp. JS666 pPC34DONZAAFQ020000220.97 ± 0.790.81 ± 0.570.69 ± 0.50
Polaromonas sp. pC12O YP_5488920.54 ± 0.301.05 ± 0.59 
Pseudomonas aeruginosa FLH04754001 catA AAT511800.62 ± 0.460.59 ± 0.65 
Pseudomonas aeruginosa JB2 chloro-C12O AAC694740.69 ± 0.36  
Pseudomonas aeruginosa PAO1 catA NC0025160.72 ± 0.180.52 ± 0.18 
Pseudomonas balearica catA CAG15365 1.23 ± 0.400.97 ± 0.45
Pseudomonas putida KT2440 pC12O NP_7458460.70 ± 0.131.42 ± 1.061.97 ± 0.68
Pseudomonas putida mt-2 pCN12 D377820.87 ± 0.511.19 ± 0.960.68 ± 0.41
Pseudomonas putida PRS1 catA U125570.64 ± 0.29  
Pseudomonas sp. CA10 catA AB0472720.82 ± 0.570.95 ± 0.570.59 ± 0.47
Pseudomonas sp. P51 chloro-C12o P270980.63 ± 0.36  
Pseudomonas stutzeri ATCC 27951 catA AJ6175141.03 ± 0.501.17 ± 0.230.77 ± 0.57
Pseudomonas stutzeri CCUG11256 catA AJ617513 0.68 ± 0.25 
Pseudomonas synthetic construct catA AAT511800.61 ± 0.290.85 ± 0.160.98 ± 0.92
Pseudomonas syringae pv. syringae B72 pPC34DO CP0000751.23 ± 0.601.14 ± 0.610.55 ± 0.37
Rhodobacterales bacterium Y4I pPC34DO1 YP_0026939410.56 ± 0.140.70 ± 0.190.52 ± 0.35
Rhodococcus opacus 1CP pcaH AF003947 0.57 ± 0.29 
Rhodococcus rhodochrous NCIMB 13259 C12O AAC330030.51 ± 0.42  
Roseovarius nubinhibens ISM pPC34DO ZP_009613950.64 ± 0.31  
Rubrobacter xylanophilus DSM 9941 PC34DO YP_6443550.61 ± 0.51  
Ruegeria pomeroyi DSS-3 pPC34DO YP_1648751.45 ± 1.120.75 ± 0.381.26 ± 0.65
Solibacter usitatus Ellin6076 pPC34DO YP_8215440.52 ± 0.18 1.15 ± 0.46
Streptomyces coelicolor A3(2) SCO66 AL9391280.61 ± 0.29  
Streptomyces setonii catA AAK140650.56 ± 0.140.70 ± 0.190.52 ± 0.35
Streptomyces setonii catB AF2770510.71 ± 0.281.65 ± 1.220.83 ± 0.39
Streptomyces setonii catA AAK140650.66 ± 0.300.60 ± 0.380.82 ± 0.52
Streptomyces sp. 2065 PC34DO AAD05270 0.56 ± 0.34 
Uncultured organism C12O AAW82364 2.56 ± 1.821.59 ± 0.67
Xanthomonas axonopodis pv. citri str. 306 pPC34DO NP_6407231.76 ± 0.345.39 ± 2.765.10 ± 1.12
Xanthomonas campestris pv campestris pPC34DO NC003902a0.52 ± 0.15  
Xanthomonas campestris pv. campestris str. ATCC pPC34DO NP_6357620.77 ± 0.61 0.85 ± 0.40
Xanthomonas oryzae pv. oryzae KACC10331 pPC34Do YP_1991210.88 ± 0.511.01 ± 0.390.78 ± 0.45
Table 5.   Positive hybridization signals of DNA extracted from PHDC with the microarray probes against 197 CDS of protein members representing all the main phylogenetic branches of the EXDO family (see Fig. S2)
EXDO gene target*A. schauerianaR. mangleSediment
  • *

    EXDO gene targets are described with the genus or species name and bacterial strain identification of the microorganism where the aromatic ring-metacleavage protein member of the EXDO family has been found, followed by the gene abbreviation and its GenBank accession number.

  • Signal intensities detected in DNA extracted from the PHDC bulk sediment (Sediment), Avicenia schaueriana, Rhizophora mangle above the significant relative intensities (above threshold of 0.5 normalized against the overall background) are shown as the triplicate mean values with the corresponding standard deviations.

Burkholderia vietnamiensis G4 c23o AAEH020000690.92 ± 0.590.77 ± 0.310.53 ± 0.39
Comamonas testosteroni TA441 tesB AB040808 0.84 ± 0.23 
Nocardia farcinica IFM10152 nfa30490 AP0066180.52 ± 0.021.01 ± 0.431.28 ± 0.63
Pseudomonas putida MT53 xylE pWW53 AF1028911.14 ± 0.711.05 ± 0.620.86 ± 0.66
Pseudomonas stutzeri OX1 cdo AJ4967390.94 ± 0.410.66 ± 0.280.65 ± 0.30
Sphingomonas aromaticivorans F199 xylE pNL1 AF079317 1.04 ± 0.261.13 ± 0.14
Sphingomonas sp. CHY-1 phnC AJ633552  0.57 ± 0.19

Whereas multiple signals for INDO-encoding genes were observed, which, with the exception of chlorocatechol 1,2-dioxygenases, show a typical chromosomal location and therefore can be regarded as a kind of phylogenetic marker (Pérez-Pantoja et al., 2010), only a few signals due to the presence of genes encoding EXDO of the vicinal chelate superfamily were evident. In fact, microarray results detected genes encoding EXDO of the subfamily I2A (according to the nomenclature of Eltis & Bolin, 1996), which have been described previously as being responsible for extradiol cleavage of catechol generated from toluene after monooxygenation of the side chain (Nakai et al., 1983), from naphthalene after 1-hydroxylation of intermediate salicylate (Bosch et al., 2000), from phenol by soluble diiron multicomponent oxygenases (Bartilson & Shingler, 1989) or from benzene and toluene after dioxygenolytic attack (Junca & Pieper, 2004), and that are typically harbored by Pseudomonas spp. strains. Also, the presence of genes encoding EXDO of the subfamily I2C, often observed as chromosomally encoded in Burkholderiales involved in phenol degradation, was observed. In contrast, signals indicating the presence of genes similar to Ralstonia sp. U2 nagC (GenBank accession number AF036940) encoding 1,2-dihydroxynaphthalene dioxygenases involved in naphthalene degradation and typically encoded in the same gene cluster as nagAc, observed in PHDCs by PCR-based profiling, were not detected. This obviously does not mean that the associated genes are not present or enriched in PHDCs, but may rather occur in quantities too low to yield signals above the threshold of normalization. Taking into account that a complex mixture of petroleum hydrocarbons was used as an enrichment substrate, this is not surprising and the enrichment may have overall selected for a complex community of alkane and monoaromatic degrading Proteobacteria and Actinobacteria, only some of which have the capability to degrade higher molecular weight aromatics. Light petroleum generally possesses about 15% of its weight in aromatic compounds, where the lightest aromatic fractions (benzene, toluene, ethylbenzene and xylenes) are by far the most abundant aromatic hydrocarbons (Wang et al., 2003). Therefore, it is possible that in this study, the initial mechanisms to break high-molecular-weight polyaromatics could be a minority compared with the number of genes encoding mechanisms to cleave, for example, n-alkanes and monoaromatics. It is certainly an aspect to be studied in further works applying tools to achieve the detection of a broader set of catabolic gene families (i.e. including the families responsible for ring activation mechanisms such as initial dioxygenases and monooxygenases), analogous to the approach for INDO and EXDO performed in this work.

Conclusions and future research

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

The identification of functionally important microbial components involved in the degradation of environmental pollutants is a primary step toward understanding the ecological mechanisms of environmental recovery. In this study, the detection of the functional components involved in the process of PH decontamination in mangrove microniches was enhanced due to the combination of batch enrichment cultures and molecular analyses. This approach allowed us to identify the bacterial phylotypes, plasmids and degrading genes potentially involved in the removal of PH in rhizosphere and bulk sediments of a mangrove chronically polluted with oil hydrocarbons. In general, the functional genes and plasmids involved in the process of PH removal detected in the PHDC were below the detection levels in the environmental samples. The molecular analyses of PHDC support the notion that differences in the composition of the original communities used for the enrichments triggered the structural and functional differences between PHDC. The differences in the composition of enriched hydrocarbonoclastic bacterial communities and specific degrading genes between PHDC may reflect the adaptation of the microbial communities to preferential substrates available in their original microniches.

The isolation of plant beneficial microorganisms for plant inoculation is a common agricultural practice already established for centuries. Basically, microorganisms are isolated from environmental samples (e.g. rhizospheres, plant tissues) and screened for the desired phenotypic characteristic for further use in agriculture as plant growth promoters. This same principle has been hypothesized to promote phytoremediation and plant growth in the presence of phytotoxic PH (Barac et al., 2004; Germaine et al., 2009). This approach can be especially interesting for the recovery of PH-polluted mangrove forests. Rhizoengineering approaches based on the combination of replanting mangrove methodologies and plant root inoculation with PH-degrading bacterial guilds and plasmids can be a promising approach that may couple both reforestation and remediation of PH-impacted mangrove forests. The rhizosphere PHDC recovered in this study can be evaluated in future investigations as a starting point to engineer efficient plant–microorganism consortia aimed at improving replanted sapling survival and growth in PH-impacted mangrove forests. In addition, they most likely constitute suitable reservoirs for future metagenomics exploration of gene clusters or for the domestication of novel bacteria involved in PH degradation.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

This study was funded by the Deutsche Forschungsgemeinschaft SM59/4-1 and 4-2 and by FAPERJ-Brazil. D.H.P., R.V. and H.J. thank Silke Kahl for the excellent technical assistance, and the European Commission for financial support to the projects consortia Biotool (STREP GOCE contract no. 003998) and BACSIN (proposal no. 211684). CGF acknowledges the support of Alexander-von-Humboldt-Stiftung and CONICET (Argentina).


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information
  • Baek SH, Kim KH, Yin CR, Jeon CO, Im WT, Kim KK & Lee ST (2003) Isolation and characterization of bacteria capable of degrading phenol and reducing nitrate under low-oxygen conditions. Curr Microbiol 47: 462466.
  • Bale MJ, Fry JC & Day MJ (1988) Transfer and occurrence of large mercury resistance plasmids in river epilithon. Appl Environ Microb 54: 972978.
  • Barac T, Taghavi S, Borremans B, Provoost A, Oeyen L, Colpaert JV, Vangronsveld J & Van Der Lelie D (2004) Engineered endophytic bacteria improve phytoremediation of water-soluble, volatile, organic pollutants. Nat Biotechnol 22: 583588.
  • Bartilson M & Shingler V (1989) Nucleotide sequence and expression of the catechol 2,3-dioxygenase-encoding gene of phenol-catabolizing Pseudomonas CF600. Gene 85: 233238.
  • Beeson KE, Erdner DL, Bagwell CE, Lovell CR & Sobecky PA (2002) Differentiation of plasmids in marine diazotrophs assemblages determined by randomly amplified polymorphic DNA analysis. Microbiology 148: 179189.
  • Benson NU & Essien JP (2009) Petroleum hydrocarbons contamination of sediments and accumulation in Tympanotonus fuscatus var. radula from the Qua Iboe Mangrove Ecosystem, Nigeria. Curr Sci India 96: 238244.
  • Bosch R, Garcia-Valdes E & Moore ER (2000) Complete nucleotide sequence and evolutionary significance of a chromosomally encoded naphthalene-degradation lower pathway from Pseudomonas stutzeri AN10. Gene 245: 6574.
  • Brito EM, Guyoneaud R, Goni-Urriza M, Ranchou-Peyruse A, Verbaere A, Crapez MA, Wasserman JC & Duran R (2006) Characterization of hydrocarbonoclastic bacterial communities from mangrove sediments in Guanabara Bay, Brazil. Res Microbiol 157: 752762.
  • Chou HH (2010) Shared probe design and existing microarray reanalysis using PICKY. BMC Bioinformatics 11: 196.
  • Clarke KR (1993) Non-parametric multivariate analyses of changes in community structure. Aust J Ecol 18: 117143.
  • Costa R, Gomes NCM, Peixoto R, Rumjanek NG, Berg G, Mendonça-Hagler LCS & Smalla K (2006) Diversity and antagonistic potential of Pseudomonas spp. associated to the rhizosphere of maize grown in a subtropical organic farm. Soil Biol Biochem 38: 24342447.
  • Das SK, Mishra AK, Tindall BJ, Rainey FA & Stackebrandt E (1996) Oxidation of thiosulfate by a new bacterium, Bosea thiooxidans (strain BI-42) gen. nov., sp. nov.: analysis of phylogeny based on chemotaxonomy and 16S ribosomal DNA sequencing. Int J Syst Bacteriol 46: 981987.
  • Dennis JJ (2005) The evolution of IncP catabolic plasmids. Curr Opin Biotech 16: 291298.
  • Dionisi H, Chewning C, Morgan K, Menn F, Easter J & Sayler G (2004) Abundance of dioxygenase genes similar to Ralstonia sp. strain U2 nagAc is correlated with naphthalene concentrations in coal tar-contaminated freshwater sediments. Appl Environ Microb 70: 39883995.
  • Eltis LD & Bolin JT (1996) Evolutionary relationships among extradiol dioxygenases. J Bacteriol 178: 59305937.
  • Entcheva P, Liebl W, Johann A, Hartsch T & Streit WR (2001) Direct cloning from enrichment cultures, a reliable strategy for isolation of complete operons and genes from microbial consortia. Appl Environ Microb 67: 8999.
  • Eulberg D, Kourbatova EM, Golovleva LA & Schlömann M (1998) Evolutionary relationship between chlorocatechol catabolic enzymes from Rhodococcus opacus 1CP and their counterparts in proteobacteria: sequence divergence and functional convergence. J Bacteriol 180: 10821094.
  • Fox RE, Zhong X, Krone SM & Top EM (2008) Spatial structure and nutrients promote invasion of IncP-1 plasmids in bacterial populations. ISME J 2: 10241039.
  • Fulthorpe RR, McGowan C, Maltseva OV, Holben WE & Tiedje JM (1995) 2,4-Dichlorophenoxyacetic acid degrading bacteria contain mosaics of catabolic genes. Appl Environ Microb 61: 32743281.
  • Germaine KJ, Keogh E, Ryan D & Dowling DN (2009) Bacterial endophyte-mediated naphthalene phytoprotection and phytoremediation. FEMS Microbiol Lett 296: 226234.
  • Gomes NCM, Heuer H, Schönfeld J, Costa R, Hagler-Mendonça L & Smalla K (2001) Bacterial diversity of the rhizosphere of maize (Zea mays) grown in tropical soil studied by temperature gradient gel electrophoresis. Plant Soil 232: 167180.
  • Gomes NCM, Kosheleva IA, Abraham WR & Smalla K (2005) Effects of the inoculant strain Pseudomonas putida KT2442 (pNF142) and of naphthalene contamination on the soil bacterial community. FEMS Microbiol Ecol 54: 2133.
  • Gomes NCM, Borges L, Paranhos R, Pinto FN, Krögerrecklenfort E, Mendonça-Hagler LCS & Smalla K (2007) Diversity of ndo genes in mangrove sediments exposed to different sources of PAH pollution. Appl Environ Microb 73: 73927399.
  • Gomes NCM, Borges LR, Paranhos R, Pinto FN, Mendonça-Hagler LCS & Smalla K (2008) Exploring the diversity of bacterial communities in sediments of urban mangrove forests. FEMS Microbiol Ecol 66: 96109.
  • Götz A, Pukall R, Tietze E, Prager R, Tschäpe H, Van Elsas JD & Smalla K (1996) Detection and characterization of broad-host-range plasmids in environmental bacteria by PCR. Appl Environ Microb 62: 26212628.
  • Grayston SJ, Wang S, Campbell CD & Edwards AC (1998) Selective influence of plant species on microbial diversity in the rhizosphere. Soil Biol Biochem 30: 369378.
  • Harris J (2009) Soil microbial communities and restoration ecology: facilitators or followers? Science 325: 573574.
  • Head IM (1998) Bioremediation: towards a credible technology. Microbiol 144: 599608.
  • Hedge RS & Fletcher JS (1996) Influence of plant growth stage and season on the release of root phenolics by mulberry as related to development of phytoremediation technology. Chemosphere 32: 24712479.
  • Heuer H, Krsek M, Baker P, Smalla K & Wellington EMH (1997) Analysis of actinomycete communities by specific amplification of genes encoding 16S rRNA and gel-electrophoretic separation in denaturing gradients. Appl Environ Microb 63: 32333241.
  • Heuer H, Wieland G, Schönfeld J, Schönwälder S, Gomes NCM & Smalla K (2001) Bacterial community profiling using DGGE or TGGE analysis. Environmental Molecular Microbiology: Protocols and Applications (RochelleP, ed), pp. 177190. Horizon Scientific Press, Wymondham, UK.
  • Hoff R (2002) Mangrove Recovery and Restoration. Oil Spills in Mangroves. Planning & Response Considerations (HoffR, ed), pp. 4855. Office of Response and Restoration, NOAA, Silver Spring, MD, USA.
  • Izmalkova TI, Sazonova OI, Sokolov SL, Kosheleva IA & Boronin AM (2005) Diversity of genetic systems responsible for naphthalene biodegradation in Pseudomonas fluorescens strains. Mikrobiologiya 74: 6068.
  • Jaeger CH, Lindow SE, Miller W, Clark E & Firestone MK (1999) Mapping of sugar and amino acid availability in soil around roots with bacterial sensors of sucrose and tryptophan. Appl Environ Microb 65: 26852690.
  • Jeon CO, Park W, Padmanabhan P, DeRito C, Snape JR & Madsen EL (2003) Discovery of a previously undescribed bacterium with distinctive dioxygenase that is responsible for in situ biodegradation in contaminated sediment. P Natl Acad Sci USA 100: 1359113596.
  • Junca H & Pieper DH (2004) Functional gene diversity analysis in BTEX contaminated soils by means of PCR-SSCP DNA fingerprinting: comparative diversity assessment against bacterial isolates and PCR-DNA clone libraries. Environ Microbiol 6: 95110.
  • Koren O, Knezevic V, Ron EZ & Rosenberg E (2003) Petroleum pollution bioremediation using water-insoluble uric acid as the nitrogen source. Appl Environ Microb 69: 63376339.
  • Krasowiak R, Smalla K, Sokolov S, Kosheleva I, Sevastyanovich Y, Titok M & Thomas CM (2002) PCR primers for detection and characterisation of IncP-9 plasmids. FEMS Microbiol Ecol 42: 217225.
  • Kuiper I, Bloemberg GV & Lugtenberg BJJ (2001) Selection of a plant-bacterium pair as a novel tool for rhizostimulation of polycyclic aromatic hydrocarbon-degrading bacteria. Mol Plant Microbe In 14: 11971205.
  • Kummerová M & Kmentová E (2004) Photoinduced toxicity of fluoranthene on germination and early development of plant seedling. Chemosphere 26: 387393.
  • Laurie AD & Lloyd-Jones G (1999) The phn genes of Burkholderia sp. strain RP007 constitute a divergent gene cluster for polycyclic aromatic hydrocarbon catabolism. J Bacteriol 181: 531540.
  • Lloyd-Jones G, Laurie AD, Hunter DWF & Fraser R (1999) Analysis of catabolic genes for naphthalene and phenanthrene degradation in contaminated New Zealand soils. FEMS Microbiol Ecol 29: 6979.
  • MacCormack WP & Fraile ER (1997) Characterization of a hydrocarbon degrading psychrotrophic Antarctic bacterium. Antarct Sci 9: 150155.
  • Margesin R & Schinner F (1997) Bioremediation of diesel-oil-contaminated alpine soils at low temperatures. Appl Microbiol Biot 47: 462468.
  • Milling A, Smalla K, Maidl FX, Schloter M & Munch JC (2004) Effects of transgenic potatoes with an altered starch composition on the diversity of soil and rhizosphere bacteria and fungi. Plant Soil 266: 2339.
  • Moser R & Stahl U (2001) Insights into the genetic diversity of initial dioxygenases from PAH-degrading bacteria. Appl Microbiol Biot 55: 609618.
  • Nakai C, Kagamiyama H, Nozaki M, Nakazawa T, Inouye S, Ebina Y & Nakazawa A (1983) Complete nucleotide sequence of the metapyrocatechase gene on the TOL plasmid of Pseudomonas putida mt-2. J Biol Chem 258: 29232928.
  • Nam JW, Nojiri H, Yoshida T, Habe H, Yamane H & Omori T (2001) New classification system for oxygenase components involved in ring-hydroxylating oxygenations. Biosci Biotech Bioch 65: 254263.
  • Neumann G & Römheld V (2001) The release of root exudates as affected by the plant's physiological status. The Rhizosphere – Biochemistry and Organic Substances at Soil Plant Interface (PintonR, VaraniniZ & NannipieriP, eds), pp. 4194. Marcel Dekker, New York, USA.
  • Palleroni NJ, Pieper DH & Moore ERB (2010) Microbiology of Hydrocarbon-Degrading Pseudomonas. Handbook of Hydrocarbon and Lipid Microbiology (TimmisKN, ed), pp. 17871798. Springer, Berlin, Germany.
  • Pérez-Pantoja D, Donoso R, Junca H, González B & Pieper DH (2010) Phylogenomics of Aerobic Bacterial Degradation of Aromatics. Handbook of Hydrocarbon and Lipid Microbiology (TimmisKN, ed), pp. 13551397. Springer, Berlin, Germany.
  • Pleshakova EV, Muratova AY & Turkovskaya OV (2001) Degradation of mineral oil with a strain of Acinetobacter calcoaceticus. Appl Biochem Microb 37: 342347.
  • Proffitt CE, Devlin DJ & Lindsey M (1995) Effects of oil on mangrove seedlings grown under different environmental conditions. Mar Pollut Bull 30: 788793.
  • Pukall R, Tschäpe H & Smalla K (1996) Monitoring the spread of broad host and narrow host range plasmids in soil microcosms. FEMS Microbiol Ecol 20: 5366.
  • Ramette A (2007) Multivariate analyses in microbial ecology. FEMS Microbiol Ecol 62: 142160.
  • Rasmussen LD & Sørensen SJ (1998) The effect of long term exposure to mercury on the bacterial community in marine sediment. Curr Microbiol 36: 291297.
  • Sambrook J, Fritsch EF & Maniatis T (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, New York.
  • Sar N & Rosenberg E (1983) Emulsifier production by Acinetobacter calcoaceticus strains. Curr Microbiol 9: 309314.
  • Schlüter A, Szczepanowski R, Puhler A & Top EM (2007) Genomics of IncP-1 antibiotic resistance plasmids isolated from wastewater treatment plants provides evidence for a widely accessible drug resistance gene pool. FEMS Microbiol Rev 31: 449477.
  • Seo J-S, Keum Y-S, Harada RM & Li QX (2007) Isolation and characterization of bacteria capable of degrading polycyclic aromatic hydrocarbons (PAHs) and organophosphorus pesticides from PAH-contaminated soil in Hilo, Hawaii. J Agr Food Chem 55: 53835389.
  • Smalla K, Wieland G, Buchner A, Zock A, Parzy J, Kaiser S, Roskot N, Heuer H & Berg G (2001) Bulk and rhizosphere soil bacterial communities studied by denaturing gradient gel e1ectrophoresis: plant-dependent enrichment and seasonal shifts revealed. Appl Environ Microb 67: 47424751.
  • Smalla K, Haines AS, Jones K, Krögerrecklenfort E, Heuer H, Schloter M & Thomas CM (2006) Increased abundance of IncP-1β plasmids and mercury resistance genes in mercury-polluted river sediments: first discovery of IncP-1β plasmids with a complex mer transposon as the sole accessory element. Appl Environ Microb 72: 72537259.
  • Smits TH, Witholt B & Van Beilen JB (2003) Functional characterization of genes involved in alkane oxidation by Pseudomonas aeruginosa. Antonie van Leeuwenhoek 84: 193200.
  • Sobecky PA (1999) Plasmid ecology of marine sediment microbial communities. Hydrobiologia 401: 918.
  • Sota M, Yano H, Ono A, Miyazaki R, Ishii H, Genka H, Top EM & Tsuda M (2006) Genomic and functional analysis of the IncP-9 naphthalene-catabolic plasmid NAH7 and its transposon Tn4655 suggests catabolic gene spread by a tyrosine recombinase. J Bacteriol 188: 40574067.
  • Van Hamme JD, Singh A & Ward OP (2003) Recent advances in petroleum microbiology. Microbiol Mol Biol R 67: 503549.
  • Venosa AD, Suidan MT, Wrenn BA, Strohmeier KL, Haines JR, Eberhart BL, King D & Holder E (1996) Bioremediation of an experimental oil spill on the shoreline of Delaware Bay. Environ Sci Technol 30: 17641775.
  • Wang Z, Hollebone BP, Fingas M, Fieldhouse B, Sigouin L, Landriault M, Smith P, Noonan J & Thouin G (2003) Characteristics of spilled oils, fuels, and petroleum products: 1. Composition and properties of selected oils. EPA/600/R-03/072. US Environmental Protection Agency, Cincinnati, OH.
  • Widada J, Nojiri H, Kasuga K, Yoshida T, Habe H & Omori T (2002) Molecular detection and diversity of polycyclic aromatic hydrocarbon-degrading bacteria isolated from geographically diverse sites. Appl Microbiol Biot 58: 202209.
  • Wilson MS, Herrick JB, Jeon CO, Hinman DE & Madsen EL (2003) Horizontal transfer of phnAc dioxygenase genes within one of two phenotypically and genotypically distinctive naphthalene-degrading guilds from adjacent soil environments. Appl Environ Microb 69: 21722181.
  • Wongsa P, Tanaka M, Ueno A, Hasanuzzaman M, Yumoto I & Okuyama H (2004) Isolation and characterization of novel strains of Pseudomonas aeruginosa and Serratia marcescens possessing high efficiency to degrade gasoline, kerosene diesel oil, and lubricating oil. Curr Microbiol 49: 415422.
  • Yakimov MM, Giuliano L, Gentile G, Crisafi E, Chernikova TN, Abraham W-R, Lünsdorf H, Timmis KN & Golyshin PN (2003) Oleispira antarctica gen. nov., sp. nov., a novel hydrocarbonoclastic marine bacterium isolated from Antarctic coastal sea water. Int J Syst Evol Micr 53: 779785.

Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Conclusions and future research
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1. Phylogenetic distribution of the genes types belonging to INDO (A) and EXDO type I (B) associated to the probes producing microarray signals detected in DNA extracted from the PHDC cultures.

Fig. S2. Comparison between rhizosphere and bulk PHDC by DGGE of 16S rRNA gene fragments amplified from sediment DNA templates.

Table S1. Major relative signal intensities of the probes targeting 16S rRNA gene of 137 different phylotypes in the DNA extracted from PHDC, Avicenia schaueriana, Rhizophora mangle and Bulk sediment.

Please note: Wiley-Blackwell is not responsible for the content or functionality of any supporting materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.

FEM_962_sm_suppinformation.doc6561KSupporting info item

Please note: Wiley Blackwell is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.