Environ Sci Pollut Res (2014) 21:3634–3645 DOI 10.1007/s11356-013-2365-7
RESEARCH ARTICLE
Bioremediation potential of microorganisms from a sandy beach affected by a major oil spill Izabela Reis & C. Marisa R. Almeida & Catarina M. Magalhães & Jaqueline Cochofel & Paula Guedes & M. Clara P. Basto & Adriano A. Bordalo & Ana P. Mucha
Received: 11 August 2013 / Accepted: 11 November 2013 / Published online: 24 November 2013 # Springer-Verlag Berlin Heidelberg 2013
Abstract The aim of this work was to evaluate the bioremediation potential of microorganisms from intertidal sediments of a sandy beach affected by a major oil spill 7 years before and subject to chronic petroleum contamination since then. For that, the response of microorganisms to a new oil contamination was assessed in terms of community structure, abundance, and capacity to degrade hydrocarbons. Experiments were carried out under laboratory-controlled conditions by mixing sediment with crude oil with three different nitrogen supplementations in 50 ml serum bottles under constant shake for 15 days. Autochthonous microorganisms were able to respond to the new oil contamination by increasing their abundance (quantified by DAPI) and changing the community structure (evaluated by DGGE). This response was particularly clear for some specific bacterial groups such as Pseudomonas, Actinomycetales, and Betaproteobacteria. These communities presented an important potential for hydrocarbon degradation (up to 85 % for TPHs and 70 % for total PAHs), being the biodegradation stimulated by addition of an appropriate amount of nitrogen.
Responsible editor: Robert Duran I. Reis : C. M. R. Almeida (*) : C. M. Magalhães : J. Cochofel : P. Guedes : A. A. Bordalo : A. P. Mucha CIMAR/CIIMAR—Centro Interdisciplinar de Investigação Marinha e Ambiental, Universidade do Porto, Rua dos Bragas, 289, 4050-123 Porto, Portugal e-mail: [email protected] I. Reis : J. Cochofel : P. Guedes : M. C. P. Basto CIMAR/CIIMAR and Faculdade de Ciências, Universidade do Porto, Rua do Campo Alegre, 4169-007 Porto, Portugal A. A. Bordalo Laboratório de Hidrobiologia, Instituto de Ciências Biomédicas, Universidade do Porto (ICBAS-UP), Rua Viterbo Ferreira, 228, 4050-313 Porto, Portugal
Keywords Oil contamination . Hydrocarbons . Microbial communities . Hydrocarbon degraders . DGGE
Introduction During the past decade, bioremediation of petroleumcontaminated soils has been a hot issue in environmental research, and many bioremediation strategies have been developed and improved to clean up soils/sediments polluted with petroleum and its derivatives (Xu and Lu 2010). Bioremediation involves highly heterogeneous and complex processes, being the success of oil bioremediation dependent on having the appropriate microorganisms in place under suitable environmental conditions (Zhu et al. 2001). Bacteria are considered to represent the predominant agents of hydrocarbon degradation in the environment (Röling et al. 2002). All hydrocarbon biological remediation processes have a shared principle, a technology based in specialized hydrocarbon-degrading microorganisms (HD). It is well known that HD are ubiquitous in the environment and have been found in habitats ranging from polar soils to marine environments, including salt marshes (Daane et al. 2001). Their distribution is also related to the historical exposure of a particular environment to hydrocarbons. In fact, while these degraders are generally found at much lower concentrations in pristine environments (Margesin et al. 2003), those environments with a recent or with a chronic oil contamination tend to have a higher percentage of hydrocarbon degraders than unpolluted areas (Zhu et al. 2001). It has been shown that when an environment is contaminated with petroleum, the proportion of hydrocarbon-degrading microorganisms increases rapidly, so the levels of these microorganisms generally reflect the degree of contamination of the ecosystem (Atlas 1995; Leahy and Colwell 1990; Vidali 2001). Nonetheless, it is
Environ Sci Pollut Res (2014) 21:3634–3645
important to test microbial potential to effectively degrade the hydrocarbons. Although several studies are available on the use of bioremediation on post-hydrocarbon contamination soil and beaches, the response of microorganisms from oil-impacted beaches to a new oil contamination skipped the attention of scientists. The aim of this work was to evaluate the bioremediation potential of microorganisms from intertidal sediments of a sandy beach 7 years after a major oil spill, a beach subject to chronic petroleum contamination since then. For that, the response of microorganisms to a new oil contamination was assessed in terms of community structure, abundance, and capacity to degrade hydrocarbons. Experiments were carried out under laboratory-controlled conditions in order to isolate microbial activity from physical–chemical factors. To our knowledge, no data is found on hydrocarbon degradation potential of beach microbial communities taken under laboratory-controlled conditions. The beach selected was the sandy beach of O Rostro in the northwestern Spanish coast. In November 2002, more than 800 km of this coast was affected by the wreck of the Prestige oil tanker that occurred near the Galician coast (Díez et al. 2005). At that time, Prestige was carrying 77,000 t of heavy fuel (Alonso-Gutiérrez et al. 2008), with a composition of 22 % of aliphatic hydrocarbons, 50 % aromatics hydrocarbons, and 28 % of resins and asphaltenes (Albaigés et al. 2006). Large spills of heavy fuel oil are barely dispersed in the water column, being mainly stranded on the shoreline or sediments in the form of patches or tar aggregates (AlonsoGutiérrez et al. 2008). Although a huge process of clean up was launched immediately after the spill several years after the accident, oil still remains in the affected beaches (Bernabeu et al. 2009). In fact, the surveys carried out in the O Rostro beach showed still the presence of oil sparsely dispersed throughout the beach 7 years after the incident either in the form of tar balls or in the form of gray sands, indicating a chronic petroleum contamination at this site (Bernabeu et al., unpublished results). In addition, the Atlantic coast of the Iberian Peninsula, including the Galician coast, is one of the main routes of oil cargo, and the possibilities of another occurring significant oil spill cannot be excluded. So, it is important to access if the indigenous microbial communities have effectively the potential to degrade hydrocarbons to a new contamination. Nutrients are required to support soil microbial activity and, therefore, are a key issue in any bioremediation process. Due to the high carbon content of oil, the levels of other nutrients such as nitrogen and phosphorus, essential for microbial growth, can be low, limiting the rate and extent of degradation (Prince 1997). For example, treating petroleumcontaminated soil with nitrogen can increase cell growth rates, decrease the microbial lag phase, help to maintain microbial
3635
populations at high activity levels, and therefore increase the rate of hydrocarbon degradation (Walworth et al. 2007). Therefore, in the present work, experiments were carried out with three different nitrogen supplementations.
Materials and methods Study area and sampling procedure and sediment characteristics In October 2009, sediment samples were collected from the intertidal area of the sandy beach O Rostro (42.960° N; 9.270° W). The beach is 2,000 m long and is very exposed to dominant and most frequent waves. It is characterized by an oxic medium-coarse sand (97 % of total grain size was between 0.25 and 1.0 mm), with a very low organic matter content (0.14±0.01 %). Nutrient concentrations in interstitial water were 28 μM NO3−, 0.16 μM NO2−, 1.4 μM NH4+, and 0.56 μM PO4−. In November 2002, this beach was directly affected by the Prestige oil spill, and since then, it has been monitoring, being constantly found superficial tar bars spread randomly through the beach (Bernabeu et al. 2009). Sand from the middle intertidal zone at low tide, without visual oil presence (without detected hydrocarbons concentrations), was collected from a 5 to 50 cm depth layer and thoroughly homogenized to avoid small-scale variations that could influence the experiment. Samples were collected in sterile plastic bags and transported to the laboratory in refrigerated ice chests. In the laboratory, the sediment was separated, and the portion which was in contact with the plastic bags was discarded. Laboratory experiments For the laboratorial evaluation of the biodegradation potential, experimental work involving three different nitrogen supplementations was performed in aerobic conditions, as no oxygen limitation exists in the environment were the sediments were collected. At the laboratory, the collected sediment was immediately used for the experiments, being distributed among the different glass vessels and doped with crude oil. The experimental design adopted was briefly 10 ml (volume) of sediment samples (S) were placed in 50 ml flasks, with 20 ml Bushnell–Haas (BH) medium supplemented with 2 % NaCl (M) and 0.5 ml of Arabian light crude oil (O) (supplied by an oil refinery). The crude oil had ca 50 % saturated hydrocarbons, 40 % aromatic hydrocarbons, and around 10 % resins, asphaltenes, and waxes. The BH medium already contains nitrogen (10 mM of KNO3 plus 9 mM of NH4NO3), being this the first treatment performed (M + O). Nitrogen was then extra supplemented into solution using two different concentrations, 20 mM of N (N) and 40 mM of N (2N) (both as
3636
KNO3), being these the second and third treatment tested (M + O + N and M + O + 2N, respectively). Initial triplicate flasks with sediment samples were collected for analysis and considered as T0 samples. The remaining flasks, with triplicate sediment samples per treatment, were incubated at room temperature in the dark under constant stirring at 100 rpm for 15 days. The flasks were also manually shaken once every day to improve blending between crude oil and sediment. Abiotic losses were assessed in flaks with sterile sediment spiked with oil. Sterilization was done by autoclaving the sediment at 121 °C. This sterilization process was not 100 % effective but reduced microbial abundance in four orders of magnitude (from 107 to 103 TCC g−1wet). Simultaneously, sediment samples, not spiked with crude oil, were also incubated in the abovementioned conditions to verify the effect of the media on the microbial community under different nitrogen supplementation. After 15 days of incubation, the samples were centrifuged, and the sediment samples were removed and considered as T15 samples. Samples (including T0) were frozen at −20 °C for further analysis of hydrocarbons content (store in aluminum foil) and DNA extraction. It was assumed that due to centrifugation (and to the relatively high grain size of the sediment), particulate sediment was deposit and that no significant amount would remain in solution. Therefore, all measurements were carried in the sediments. Hydrocarbon analysis For total petroleum hydrocarbon (TPH) measurements, a previous optimized method was used (Couto et al. 2011). Sediment, after thawing, was ultrasonic (Elma, Transsonic 460/H model) extracted with tetrachloroethylene (≥99 % spectrophotometric grade from Sigma-Aldrich) [1:10 (w/v)], being the extracts analyzed by Fourier transform infrared spectrophotometry (Jasco FT/IR-460 Plus) using a quartz cell of 10 or 40 mm path length (Infrasil I, Starna Scientific). Although no significant differences were observed in TPHs concentrations in dry and wet samples (Guedes 2010), analysis was carried out in wet samples to prevent any possible loss of volatile compounds during drying. Considering that the same type and amount of sediment and medium was used in the entire experiment, differences among sample moistures were considered negligible. To have a chemical profile of the saturated hydrocarbons (a smaller range than that covered by FTIR determination), analyses by gas chromatography with flame ionization detection (GC-FID) of dry sediments were also carried out. Sediments were firstly extracted with a mixture of n -hexane (95 %, UV-IR-HPLC, PAI-ACS, Panreac) and acetone (≥99.9 %, CHROMASOLV® plus, HPLC, Sigma-Aldrich) [1:1 (v /v )] in the abovementioned ultrasonic device for 30 min. The extracts were cleaned with Florisil® (60–100
Environ Sci Pollut Res (2014) 21:3634–3645
mesh, Fluka) and analyzed in a GC-FID (Varian 3800) with a column 30 m×0.25 mm×0.25 μm (Varian Factor Four, VF5ht), following the conditions described in Saari et al. (2007). The hydrocarbon range window measured was between nonane (C9) and tetracontane (C40). Polycyclic aromatic hydrocarbon (PAH) concentrations [the 16 considered a priority by the Environmental Protection Agency of the USA (USEPA and US Army Corps of Engineers 1991)] were measured as described in Rocha et al. (2011), after extraction from sediments in a microwave system [Panasonic NE-1037, with Parr reactor bombs (model 4782)] with acetone, followed by cleaning of the extract with florisil (adsorbent for chromatography, 60–100 mesh, Fluka), and analysis in a Varian CP-3800 gas chromatographer (GC) provided with a Saturn 2200 ion trap mass spectrometric detection system and a solid-phase micro-extraction device. Enumeration of microorganisms Total cell counts (TCC), expressed as in per gram wet sediment, were obtained by 4′,6′-diamidino-2-phenylindole (DAPI) direct count method (Porter and Feig 1980; Kepner and Pratt 1994). Triplicate sediment samples were immediately fixed with formaldehyde (0.2 μm filtered) to reach a final concentration of 4 % (v/v). For that, 0.1 g of homogenized samples were added to 2.5 mL of saline solution (0.2 μm filtered, 9 g L−1 NaCl), 200 μL of Tween 80 [0.2 μm filtered, 12.5 % (v/v)], and fixed with 1 mL of formaldehyde [0.2 μm filtered, 4 % (v /v )]. Samples were stirred at 150 rpm for 15 min followed by sonication for 20–30 s at low intensity (0.5 cycles, 20 % amplitude). Subsamples of fixed solution were then stained with DAPI and incubated in the dark for 12 min. Samples were filtered onto black Nuclepore polycarbonate filters (0.2 μm pore size, 25 mm diameter, Whatman, UK) under gentle vacuum and washed with autoclaved 0.2 μm-filtered distilled water. Membranes were mounted in glass slides and cells counted at ×1,875 using an epifluorescence microscope (Labphot, Nikon, Japan). Enumeration of hydrocarbon-degrading microbial populations Most probable numbers of culturable hydrocarbon degraders (MPN) were estimated according to the protocol developed by Wrenn and Venosa (1996) in 96-well microtiter plates. Prefiltered (0.2 μm) Arabian light crude oil was the selective substrate for determination of total hydrocarbon degraders. BH medium supplemented with 2 % NaCl was used as the growth medium for MPN procedures (180 μL BH/well). The crude oil was added to 96well microtiter plates (10 μL/well) after the plates were filled with growth medium. For each sample, 1 g of sediment was mixed in 1 mL BH, and the supernatant
Environ Sci Pollut Res (2014) 21:3634–3645
was diluted in a saline buffer solution containing 0.1 % sodium pyrophosphate (pH 7.5) and 2 % NaCl. Tenfold serial dilution was performed, and the plates were inoculated by adding 20 μL of each dilution to five rows of 11 wells. Five wells remained uninoculated to serve as sterile control. MPN plates were incubated for 2 weeks at room temperature. After incubation, 50 μL of filter sterilized Iodonitrotetrazolium violet (3 g L−1) was added to each well. Positive wells were scored after overnight incubation at room temperature. A computer program (Mike Curiale Calculator) was used to calculate the MPN. DNA extraction and PCR amplification The DNA from three independent replicates (1 g of homogenized sediments) was extracted from all treatments except those with nutrients only (M + N and M + 2N) using a modification of the CTAB (bromide–polyvinylpyrrolidone– βmercaptoethanol) extraction protocol (Coyne et al. 2001) described by Barrett et al. (2006). Each replicate of extracted DNA was amplified in 50 μL volumes containing general bacteria, Actinomycetales, Pseudomonas , Beta, and Alphaproteobacteria specific primers (Table 1; STABVIDA, Lisbon, Portugal) according to Gomes et al. (2001) and Milling et al. (2004). All polymerase chain reactions (PCR) were performed on a Mastercycler gradient (Eppendorf) thermal cycle. PCR mixtures for all amplifications contained 5– 20 ng of DNA, 100 μM of both primers (STABVIDA, Lisbon, Portugal), 200 μM dNTPs (Finnzymes), 1× Taq PCR buffer, 1 U (Beta and Alphaproteobacteria), 2.5 U (bacteria and Pseudomonas), and 1.25 U (Actinomycetales) of High Purity Taq DNA Polymerase (Citomed), 3.75 mM (bacteria, Beta, and Alphaproteobacteria), and 2.5 mM (Actinomycetales and Pseudomonas) of MgCl2 (Citomed). The PCR mixture was held 5 min at 94 °C, followed by 25 cycles of 1 min at 94 °C, 1 min at 56 °C (bacteria and Alphaproteobacteria)/61 °C (Betaproteobacteria)/63 °C (Actinomycetales and Pseudomonas), and 2 min at 72 °C, followed by a final extension at 72 °C for 10 min. The PCR products (2 μL) obtained was used for a PCR-denaturing gradient gel electrophoresis (DGGE) using general bacterial primers with a 40-bp GC clamp added to the 5′-end of the forward primer (Table 1). PCR mixture was the same used for previous general bacteria amplification. This PCR was run using the following conditions: initial denaturation of the template DNA at 94 °C for 5 min, 94 °C for 1 min, 60 °C for 1 min, and 72 °C for 2 min and a final elongation step at 72 °C for 10 min. Negative control reaction mixtures containing no template were included in all amplifications performed. PCR products were visualized by ethidium bromide staining after standard agarose gel (1.5 %) electrophoresis. One and 10-Kb ladders (Bioline and Promega, respectively) were used as a size standard, according with the size of the products obtained. PCR products were then
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quantified with the Qubit fluorometer (Invitrogen) and standardized amount of the mixed template product (800 ng) was resolved by DGGE. DGGE DGGE was performed in a CBS Scientific DGGE system (Del Mar, CA), essentially following Gomes et al. (2001, 2005). PCR product (800 ng) from each replicate sample was loaded on a 6.5 % polyacrylamide gel containing a gradient of denaturant (urea and formamide) from 40 to 70 %. PCR reactions containing DNA from one of the samples were used as DGGE standard by running it four times at each DGGE gel. Gels were run for 17 h at a constant voltage of 67/68 V in 1× TAE buffer at 60 °C. Gels were stained using a solution of Syber Green at 1× (Lonza) and DGGE profiles visualized in a molecular image Gel Doc XR system (Bio-Rad). Data analysis Statistically significant differences among samples for 5 % level of significance were evaluated through ANOVA tests using Statistica software and Tukey pairwise comparisons. Composite DGGE profiles were converted to a densitometry scan and aligned using the image analysis software Quantity One 1-D Analysis Software (Bio-Rad). The presence or absence of DGGE bands in each sample was used as input variables to evaluate differences in general bacteria, Actinomycetales, Betaproteobacteria, Alphaproteobacteria, and genus Pseudomonas assemblage composition. Samples were then analyzed using the Bray–Curtis similarity method and clustered in the on group average linking mode with the default parameters (5 % significance; mean number of permutations, 1,000; number of simulations, 999) to generate a drendrogram based on percent similarity. Significance of groupings in the cluster analysis was tested using similarity profile test (SIMPROF). PRIMER 6 Software package (version 6.1.11) was used for both CLUSTER and SIMPROF analysis.
Results Hydrocarbon concentrations and degradation TPH levels in sediments submitted to a new oil contamination (M + O) and supplemented with different nitrogen concentrations decreased by 61–85 % (Fig. 1). Addition of 20 mM N (M + O + N) enhanced significantly (p 0.05) differences between treatments with 20 mM N (M + O + N) and
3638
Environ Sci Pollut Res (2014) 21:3634–3645
Table 1 Oligonucleotide probes used in amplification of the gene encoding the 16S rRNA fragment Target group
Primers
Sequence (5′–3′)
Size (bp)
Reference
Bacteria
F27 R1492 F984 β R1494 F203α R1494 F243 R1378 F311Ps R1459Ps F984-GC R1378 GC-Clamp
AGAGTTTGATC(A/C)TGGCTCAG TACGG(C/T)TACCTTGTTACGACTT CGCACAAGCGGTGGATGA CTACGG(T/C)TACCTTGTTACGAC CCGCATACGCCCTACGGGGGAAAGATTTAT CTACGG(T/C)TACCTTGTTACGAC GGATGAGCCCGCGGCCTA CGGTGTGTACAAGGCCCGGGAACG CTGGTCTGAGAGGATGATCAGT AATCACTCCGTGGTAACCGT AACGCGAAGAACCTTAC CGGTGTGTACAAGGCCCGGGAACG CGCCCGGGGCGCGCCCCGGGCGGGGCGGGGGCACGGGGGG
1,465
Weisburg et al. 1991 Heuer and Smalla 1997 Heuer and Smalla 1997 Weisburg et al. 1991 Gomes et al. 2001 Weisburg et al. 1991 Heuer and Smalla 1997 Heuer and Smalla 1997 Widmer et al. 1998 Milling et al. 2004 Heuer and Smalla 1997 Heuer and Smalla 1997 Nübel et al. 1996
Betaproteobacteria Alfaproteobacteria Actinomycetales Pseudomonas Universal bacteria (DGGE-PCR)
40 mM N (M + O + 2N) were found. For the sterile sediments, no significant differences were observed between initial and final levels of TPHs, indicating that no significant hydrocarbons degradation occurred due to abiotic factors (results not shown). Data from the saturated hydrocarbons (a smaller range than that covered by FTIR determination) corroborate the data from the TPHs.In fact, the chemical profiles observed in the chromatograms (Fig. 2) showed a degradation of n -alkanes after the 15 days of incubation being it much more pronounced in the treatments with extra 20 mM (M + O + N) and 40 mM of nitrogen (M + O + 2N). At the end of the incubation, the total PAHs concentration (sum of the concentration of the eight PAHs detected) decreased about 38 % (M + O) (Fig. 1). Supplementation with 20 mM N (M + O + N) increased PAHs degradation to 46 % and the maximum degradation (70 %) was obtained at the highest nitrogen addition (M + O + 2N). Of the 16 PAHs analyzed, only eight were detected in the samples (Fig. 3), presenting anthracene and phenanthrene the highest concentrations. For these two compounds, degradation increased with nitrogen supplementation reaching, respectively, 70 and 72 % with the highest nitrogen addition. The same was observed for chrysene with a maximum of 61 % of degradation. For fluorene, no significant (p >0.05) differences were registered among treatments since degradation was total after 15 days. Complete degradation was also observed for pyrene, but only at the highest nitrogen addition. For naphthalene, a small but significant (p
RESEARCH ARTICLE
Bioremediation potential of microorganisms from a sandy beach affected by a major oil spill Izabela Reis & C. Marisa R. Almeida & Catarina M. Magalhães & Jaqueline Cochofel & Paula Guedes & M. Clara P. Basto & Adriano A. Bordalo & Ana P. Mucha
Received: 11 August 2013 / Accepted: 11 November 2013 / Published online: 24 November 2013 # Springer-Verlag Berlin Heidelberg 2013
Abstract The aim of this work was to evaluate the bioremediation potential of microorganisms from intertidal sediments of a sandy beach affected by a major oil spill 7 years before and subject to chronic petroleum contamination since then. For that, the response of microorganisms to a new oil contamination was assessed in terms of community structure, abundance, and capacity to degrade hydrocarbons. Experiments were carried out under laboratory-controlled conditions by mixing sediment with crude oil with three different nitrogen supplementations in 50 ml serum bottles under constant shake for 15 days. Autochthonous microorganisms were able to respond to the new oil contamination by increasing their abundance (quantified by DAPI) and changing the community structure (evaluated by DGGE). This response was particularly clear for some specific bacterial groups such as Pseudomonas, Actinomycetales, and Betaproteobacteria. These communities presented an important potential for hydrocarbon degradation (up to 85 % for TPHs and 70 % for total PAHs), being the biodegradation stimulated by addition of an appropriate amount of nitrogen.
Responsible editor: Robert Duran I. Reis : C. M. R. Almeida (*) : C. M. Magalhães : J. Cochofel : P. Guedes : A. A. Bordalo : A. P. Mucha CIMAR/CIIMAR—Centro Interdisciplinar de Investigação Marinha e Ambiental, Universidade do Porto, Rua dos Bragas, 289, 4050-123 Porto, Portugal e-mail: [email protected] I. Reis : J. Cochofel : P. Guedes : M. C. P. Basto CIMAR/CIIMAR and Faculdade de Ciências, Universidade do Porto, Rua do Campo Alegre, 4169-007 Porto, Portugal A. A. Bordalo Laboratório de Hidrobiologia, Instituto de Ciências Biomédicas, Universidade do Porto (ICBAS-UP), Rua Viterbo Ferreira, 228, 4050-313 Porto, Portugal
Keywords Oil contamination . Hydrocarbons . Microbial communities . Hydrocarbon degraders . DGGE
Introduction During the past decade, bioremediation of petroleumcontaminated soils has been a hot issue in environmental research, and many bioremediation strategies have been developed and improved to clean up soils/sediments polluted with petroleum and its derivatives (Xu and Lu 2010). Bioremediation involves highly heterogeneous and complex processes, being the success of oil bioremediation dependent on having the appropriate microorganisms in place under suitable environmental conditions (Zhu et al. 2001). Bacteria are considered to represent the predominant agents of hydrocarbon degradation in the environment (Röling et al. 2002). All hydrocarbon biological remediation processes have a shared principle, a technology based in specialized hydrocarbon-degrading microorganisms (HD). It is well known that HD are ubiquitous in the environment and have been found in habitats ranging from polar soils to marine environments, including salt marshes (Daane et al. 2001). Their distribution is also related to the historical exposure of a particular environment to hydrocarbons. In fact, while these degraders are generally found at much lower concentrations in pristine environments (Margesin et al. 2003), those environments with a recent or with a chronic oil contamination tend to have a higher percentage of hydrocarbon degraders than unpolluted areas (Zhu et al. 2001). It has been shown that when an environment is contaminated with petroleum, the proportion of hydrocarbon-degrading microorganisms increases rapidly, so the levels of these microorganisms generally reflect the degree of contamination of the ecosystem (Atlas 1995; Leahy and Colwell 1990; Vidali 2001). Nonetheless, it is
Environ Sci Pollut Res (2014) 21:3634–3645
important to test microbial potential to effectively degrade the hydrocarbons. Although several studies are available on the use of bioremediation on post-hydrocarbon contamination soil and beaches, the response of microorganisms from oil-impacted beaches to a new oil contamination skipped the attention of scientists. The aim of this work was to evaluate the bioremediation potential of microorganisms from intertidal sediments of a sandy beach 7 years after a major oil spill, a beach subject to chronic petroleum contamination since then. For that, the response of microorganisms to a new oil contamination was assessed in terms of community structure, abundance, and capacity to degrade hydrocarbons. Experiments were carried out under laboratory-controlled conditions in order to isolate microbial activity from physical–chemical factors. To our knowledge, no data is found on hydrocarbon degradation potential of beach microbial communities taken under laboratory-controlled conditions. The beach selected was the sandy beach of O Rostro in the northwestern Spanish coast. In November 2002, more than 800 km of this coast was affected by the wreck of the Prestige oil tanker that occurred near the Galician coast (Díez et al. 2005). At that time, Prestige was carrying 77,000 t of heavy fuel (Alonso-Gutiérrez et al. 2008), with a composition of 22 % of aliphatic hydrocarbons, 50 % aromatics hydrocarbons, and 28 % of resins and asphaltenes (Albaigés et al. 2006). Large spills of heavy fuel oil are barely dispersed in the water column, being mainly stranded on the shoreline or sediments in the form of patches or tar aggregates (AlonsoGutiérrez et al. 2008). Although a huge process of clean up was launched immediately after the spill several years after the accident, oil still remains in the affected beaches (Bernabeu et al. 2009). In fact, the surveys carried out in the O Rostro beach showed still the presence of oil sparsely dispersed throughout the beach 7 years after the incident either in the form of tar balls or in the form of gray sands, indicating a chronic petroleum contamination at this site (Bernabeu et al., unpublished results). In addition, the Atlantic coast of the Iberian Peninsula, including the Galician coast, is one of the main routes of oil cargo, and the possibilities of another occurring significant oil spill cannot be excluded. So, it is important to access if the indigenous microbial communities have effectively the potential to degrade hydrocarbons to a new contamination. Nutrients are required to support soil microbial activity and, therefore, are a key issue in any bioremediation process. Due to the high carbon content of oil, the levels of other nutrients such as nitrogen and phosphorus, essential for microbial growth, can be low, limiting the rate and extent of degradation (Prince 1997). For example, treating petroleumcontaminated soil with nitrogen can increase cell growth rates, decrease the microbial lag phase, help to maintain microbial
3635
populations at high activity levels, and therefore increase the rate of hydrocarbon degradation (Walworth et al. 2007). Therefore, in the present work, experiments were carried out with three different nitrogen supplementations.
Materials and methods Study area and sampling procedure and sediment characteristics In October 2009, sediment samples were collected from the intertidal area of the sandy beach O Rostro (42.960° N; 9.270° W). The beach is 2,000 m long and is very exposed to dominant and most frequent waves. It is characterized by an oxic medium-coarse sand (97 % of total grain size was between 0.25 and 1.0 mm), with a very low organic matter content (0.14±0.01 %). Nutrient concentrations in interstitial water were 28 μM NO3−, 0.16 μM NO2−, 1.4 μM NH4+, and 0.56 μM PO4−. In November 2002, this beach was directly affected by the Prestige oil spill, and since then, it has been monitoring, being constantly found superficial tar bars spread randomly through the beach (Bernabeu et al. 2009). Sand from the middle intertidal zone at low tide, without visual oil presence (without detected hydrocarbons concentrations), was collected from a 5 to 50 cm depth layer and thoroughly homogenized to avoid small-scale variations that could influence the experiment. Samples were collected in sterile plastic bags and transported to the laboratory in refrigerated ice chests. In the laboratory, the sediment was separated, and the portion which was in contact with the plastic bags was discarded. Laboratory experiments For the laboratorial evaluation of the biodegradation potential, experimental work involving three different nitrogen supplementations was performed in aerobic conditions, as no oxygen limitation exists in the environment were the sediments were collected. At the laboratory, the collected sediment was immediately used for the experiments, being distributed among the different glass vessels and doped with crude oil. The experimental design adopted was briefly 10 ml (volume) of sediment samples (S) were placed in 50 ml flasks, with 20 ml Bushnell–Haas (BH) medium supplemented with 2 % NaCl (M) and 0.5 ml of Arabian light crude oil (O) (supplied by an oil refinery). The crude oil had ca 50 % saturated hydrocarbons, 40 % aromatic hydrocarbons, and around 10 % resins, asphaltenes, and waxes. The BH medium already contains nitrogen (10 mM of KNO3 plus 9 mM of NH4NO3), being this the first treatment performed (M + O). Nitrogen was then extra supplemented into solution using two different concentrations, 20 mM of N (N) and 40 mM of N (2N) (both as
3636
KNO3), being these the second and third treatment tested (M + O + N and M + O + 2N, respectively). Initial triplicate flasks with sediment samples were collected for analysis and considered as T0 samples. The remaining flasks, with triplicate sediment samples per treatment, were incubated at room temperature in the dark under constant stirring at 100 rpm for 15 days. The flasks were also manually shaken once every day to improve blending between crude oil and sediment. Abiotic losses were assessed in flaks with sterile sediment spiked with oil. Sterilization was done by autoclaving the sediment at 121 °C. This sterilization process was not 100 % effective but reduced microbial abundance in four orders of magnitude (from 107 to 103 TCC g−1wet). Simultaneously, sediment samples, not spiked with crude oil, were also incubated in the abovementioned conditions to verify the effect of the media on the microbial community under different nitrogen supplementation. After 15 days of incubation, the samples were centrifuged, and the sediment samples were removed and considered as T15 samples. Samples (including T0) were frozen at −20 °C for further analysis of hydrocarbons content (store in aluminum foil) and DNA extraction. It was assumed that due to centrifugation (and to the relatively high grain size of the sediment), particulate sediment was deposit and that no significant amount would remain in solution. Therefore, all measurements were carried in the sediments. Hydrocarbon analysis For total petroleum hydrocarbon (TPH) measurements, a previous optimized method was used (Couto et al. 2011). Sediment, after thawing, was ultrasonic (Elma, Transsonic 460/H model) extracted with tetrachloroethylene (≥99 % spectrophotometric grade from Sigma-Aldrich) [1:10 (w/v)], being the extracts analyzed by Fourier transform infrared spectrophotometry (Jasco FT/IR-460 Plus) using a quartz cell of 10 or 40 mm path length (Infrasil I, Starna Scientific). Although no significant differences were observed in TPHs concentrations in dry and wet samples (Guedes 2010), analysis was carried out in wet samples to prevent any possible loss of volatile compounds during drying. Considering that the same type and amount of sediment and medium was used in the entire experiment, differences among sample moistures were considered negligible. To have a chemical profile of the saturated hydrocarbons (a smaller range than that covered by FTIR determination), analyses by gas chromatography with flame ionization detection (GC-FID) of dry sediments were also carried out. Sediments were firstly extracted with a mixture of n -hexane (95 %, UV-IR-HPLC, PAI-ACS, Panreac) and acetone (≥99.9 %, CHROMASOLV® plus, HPLC, Sigma-Aldrich) [1:1 (v /v )] in the abovementioned ultrasonic device for 30 min. The extracts were cleaned with Florisil® (60–100
Environ Sci Pollut Res (2014) 21:3634–3645
mesh, Fluka) and analyzed in a GC-FID (Varian 3800) with a column 30 m×0.25 mm×0.25 μm (Varian Factor Four, VF5ht), following the conditions described in Saari et al. (2007). The hydrocarbon range window measured was between nonane (C9) and tetracontane (C40). Polycyclic aromatic hydrocarbon (PAH) concentrations [the 16 considered a priority by the Environmental Protection Agency of the USA (USEPA and US Army Corps of Engineers 1991)] were measured as described in Rocha et al. (2011), after extraction from sediments in a microwave system [Panasonic NE-1037, with Parr reactor bombs (model 4782)] with acetone, followed by cleaning of the extract with florisil (adsorbent for chromatography, 60–100 mesh, Fluka), and analysis in a Varian CP-3800 gas chromatographer (GC) provided with a Saturn 2200 ion trap mass spectrometric detection system and a solid-phase micro-extraction device. Enumeration of microorganisms Total cell counts (TCC), expressed as in per gram wet sediment, were obtained by 4′,6′-diamidino-2-phenylindole (DAPI) direct count method (Porter and Feig 1980; Kepner and Pratt 1994). Triplicate sediment samples were immediately fixed with formaldehyde (0.2 μm filtered) to reach a final concentration of 4 % (v/v). For that, 0.1 g of homogenized samples were added to 2.5 mL of saline solution (0.2 μm filtered, 9 g L−1 NaCl), 200 μL of Tween 80 [0.2 μm filtered, 12.5 % (v/v)], and fixed with 1 mL of formaldehyde [0.2 μm filtered, 4 % (v /v )]. Samples were stirred at 150 rpm for 15 min followed by sonication for 20–30 s at low intensity (0.5 cycles, 20 % amplitude). Subsamples of fixed solution were then stained with DAPI and incubated in the dark for 12 min. Samples were filtered onto black Nuclepore polycarbonate filters (0.2 μm pore size, 25 mm diameter, Whatman, UK) under gentle vacuum and washed with autoclaved 0.2 μm-filtered distilled water. Membranes were mounted in glass slides and cells counted at ×1,875 using an epifluorescence microscope (Labphot, Nikon, Japan). Enumeration of hydrocarbon-degrading microbial populations Most probable numbers of culturable hydrocarbon degraders (MPN) were estimated according to the protocol developed by Wrenn and Venosa (1996) in 96-well microtiter plates. Prefiltered (0.2 μm) Arabian light crude oil was the selective substrate for determination of total hydrocarbon degraders. BH medium supplemented with 2 % NaCl was used as the growth medium for MPN procedures (180 μL BH/well). The crude oil was added to 96well microtiter plates (10 μL/well) after the plates were filled with growth medium. For each sample, 1 g of sediment was mixed in 1 mL BH, and the supernatant
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was diluted in a saline buffer solution containing 0.1 % sodium pyrophosphate (pH 7.5) and 2 % NaCl. Tenfold serial dilution was performed, and the plates were inoculated by adding 20 μL of each dilution to five rows of 11 wells. Five wells remained uninoculated to serve as sterile control. MPN plates were incubated for 2 weeks at room temperature. After incubation, 50 μL of filter sterilized Iodonitrotetrazolium violet (3 g L−1) was added to each well. Positive wells were scored after overnight incubation at room temperature. A computer program (Mike Curiale Calculator) was used to calculate the MPN. DNA extraction and PCR amplification The DNA from three independent replicates (1 g of homogenized sediments) was extracted from all treatments except those with nutrients only (M + N and M + 2N) using a modification of the CTAB (bromide–polyvinylpyrrolidone– βmercaptoethanol) extraction protocol (Coyne et al. 2001) described by Barrett et al. (2006). Each replicate of extracted DNA was amplified in 50 μL volumes containing general bacteria, Actinomycetales, Pseudomonas , Beta, and Alphaproteobacteria specific primers (Table 1; STABVIDA, Lisbon, Portugal) according to Gomes et al. (2001) and Milling et al. (2004). All polymerase chain reactions (PCR) were performed on a Mastercycler gradient (Eppendorf) thermal cycle. PCR mixtures for all amplifications contained 5– 20 ng of DNA, 100 μM of both primers (STABVIDA, Lisbon, Portugal), 200 μM dNTPs (Finnzymes), 1× Taq PCR buffer, 1 U (Beta and Alphaproteobacteria), 2.5 U (bacteria and Pseudomonas), and 1.25 U (Actinomycetales) of High Purity Taq DNA Polymerase (Citomed), 3.75 mM (bacteria, Beta, and Alphaproteobacteria), and 2.5 mM (Actinomycetales and Pseudomonas) of MgCl2 (Citomed). The PCR mixture was held 5 min at 94 °C, followed by 25 cycles of 1 min at 94 °C, 1 min at 56 °C (bacteria and Alphaproteobacteria)/61 °C (Betaproteobacteria)/63 °C (Actinomycetales and Pseudomonas), and 2 min at 72 °C, followed by a final extension at 72 °C for 10 min. The PCR products (2 μL) obtained was used for a PCR-denaturing gradient gel electrophoresis (DGGE) using general bacterial primers with a 40-bp GC clamp added to the 5′-end of the forward primer (Table 1). PCR mixture was the same used for previous general bacteria amplification. This PCR was run using the following conditions: initial denaturation of the template DNA at 94 °C for 5 min, 94 °C for 1 min, 60 °C for 1 min, and 72 °C for 2 min and a final elongation step at 72 °C for 10 min. Negative control reaction mixtures containing no template were included in all amplifications performed. PCR products were visualized by ethidium bromide staining after standard agarose gel (1.5 %) electrophoresis. One and 10-Kb ladders (Bioline and Promega, respectively) were used as a size standard, according with the size of the products obtained. PCR products were then
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quantified with the Qubit fluorometer (Invitrogen) and standardized amount of the mixed template product (800 ng) was resolved by DGGE. DGGE DGGE was performed in a CBS Scientific DGGE system (Del Mar, CA), essentially following Gomes et al. (2001, 2005). PCR product (800 ng) from each replicate sample was loaded on a 6.5 % polyacrylamide gel containing a gradient of denaturant (urea and formamide) from 40 to 70 %. PCR reactions containing DNA from one of the samples were used as DGGE standard by running it four times at each DGGE gel. Gels were run for 17 h at a constant voltage of 67/68 V in 1× TAE buffer at 60 °C. Gels were stained using a solution of Syber Green at 1× (Lonza) and DGGE profiles visualized in a molecular image Gel Doc XR system (Bio-Rad). Data analysis Statistically significant differences among samples for 5 % level of significance were evaluated through ANOVA tests using Statistica software and Tukey pairwise comparisons. Composite DGGE profiles were converted to a densitometry scan and aligned using the image analysis software Quantity One 1-D Analysis Software (Bio-Rad). The presence or absence of DGGE bands in each sample was used as input variables to evaluate differences in general bacteria, Actinomycetales, Betaproteobacteria, Alphaproteobacteria, and genus Pseudomonas assemblage composition. Samples were then analyzed using the Bray–Curtis similarity method and clustered in the on group average linking mode with the default parameters (5 % significance; mean number of permutations, 1,000; number of simulations, 999) to generate a drendrogram based on percent similarity. Significance of groupings in the cluster analysis was tested using similarity profile test (SIMPROF). PRIMER 6 Software package (version 6.1.11) was used for both CLUSTER and SIMPROF analysis.
Results Hydrocarbon concentrations and degradation TPH levels in sediments submitted to a new oil contamination (M + O) and supplemented with different nitrogen concentrations decreased by 61–85 % (Fig. 1). Addition of 20 mM N (M + O + N) enhanced significantly (p 0.05) differences between treatments with 20 mM N (M + O + N) and
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Table 1 Oligonucleotide probes used in amplification of the gene encoding the 16S rRNA fragment Target group
Primers
Sequence (5′–3′)
Size (bp)
Reference
Bacteria
F27 R1492 F984 β R1494 F203α R1494 F243 R1378 F311Ps R1459Ps F984-GC R1378 GC-Clamp
AGAGTTTGATC(A/C)TGGCTCAG TACGG(C/T)TACCTTGTTACGACTT CGCACAAGCGGTGGATGA CTACGG(T/C)TACCTTGTTACGAC CCGCATACGCCCTACGGGGGAAAGATTTAT CTACGG(T/C)TACCTTGTTACGAC GGATGAGCCCGCGGCCTA CGGTGTGTACAAGGCCCGGGAACG CTGGTCTGAGAGGATGATCAGT AATCACTCCGTGGTAACCGT AACGCGAAGAACCTTAC CGGTGTGTACAAGGCCCGGGAACG CGCCCGGGGCGCGCCCCGGGCGGGGCGGGGGCACGGGGGG
1,465
Weisburg et al. 1991 Heuer and Smalla 1997 Heuer and Smalla 1997 Weisburg et al. 1991 Gomes et al. 2001 Weisburg et al. 1991 Heuer and Smalla 1997 Heuer and Smalla 1997 Widmer et al. 1998 Milling et al. 2004 Heuer and Smalla 1997 Heuer and Smalla 1997 Nübel et al. 1996
Betaproteobacteria Alfaproteobacteria Actinomycetales Pseudomonas Universal bacteria (DGGE-PCR)
40 mM N (M + O + 2N) were found. For the sterile sediments, no significant differences were observed between initial and final levels of TPHs, indicating that no significant hydrocarbons degradation occurred due to abiotic factors (results not shown). Data from the saturated hydrocarbons (a smaller range than that covered by FTIR determination) corroborate the data from the TPHs.In fact, the chemical profiles observed in the chromatograms (Fig. 2) showed a degradation of n -alkanes after the 15 days of incubation being it much more pronounced in the treatments with extra 20 mM (M + O + N) and 40 mM of nitrogen (M + O + 2N). At the end of the incubation, the total PAHs concentration (sum of the concentration of the eight PAHs detected) decreased about 38 % (M + O) (Fig. 1). Supplementation with 20 mM N (M + O + N) increased PAHs degradation to 46 % and the maximum degradation (70 %) was obtained at the highest nitrogen addition (M + O + 2N). Of the 16 PAHs analyzed, only eight were detected in the samples (Fig. 3), presenting anthracene and phenanthrene the highest concentrations. For these two compounds, degradation increased with nitrogen supplementation reaching, respectively, 70 and 72 % with the highest nitrogen addition. The same was observed for chrysene with a maximum of 61 % of degradation. For fluorene, no significant (p >0.05) differences were registered among treatments since degradation was total after 15 days. Complete degradation was also observed for pyrene, but only at the highest nitrogen addition. For naphthalene, a small but significant (p
