Exploring the potential of halophilic bacteria from oil

International Biodeterioration & Biodegradation 126 (2018) 231e242

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Exploring the potential of halophilic bacteria from oil terminal environments for biosurfactant production and hydrocarbon degradation under high-salinity conditions M.B. Gomes a, *, E.E. Gonzales-Limache a, S.T.P. Sousa a, B.M. Dellagnezze a, A. Sartoratto b, L.C.F. Silva c, L.M. Gieg d, E. Valoni e, R.S. Souza e, A.P.R. Torres e, M.P. Sousa e, S.O. De Paula c, C.C. Silva c, V.M. Oliveira a a Microbial Resources Division, Research Center for Chemistry, Biology and Agriculture (CPQBA), Campinas University - UNICAMP, CP 6171, CEP 13081-970, Campinas, SP, Brazil b Organic Chemistry and Pharmaceutical Division, Research Center for Chemistry, Biology and Agriculture (CPQBA), Campinas University e UNICAMP, CP 6171, CEP 13083-970, Campinas, SP, Brazil c General Biology Department, Federal University of Viçosa e UFV, CP 36570-000, Viçosa, MG, Brazil d Petroleum Microbiology Research Group, Department of Biological Sciences, University of Calgary, 2500 University Drive NW, Calgary, Alberta, Canada e cio Macedo, 950, Expansa ~o, Ala C, 21941-915, Ilha do PETROBRAS Research and Development Center (CENPES), Biotechnology Management, Av. Hora ~o, Rio de Janeiro, RJ, Brazil Funda

a r t i c l e i n f o

a b s t r a c t

Article history: Received 5 January 2016 Received in revised form 5 July 2016 Accepted 25 August 2016 Available online 10 September 2016

The wastewater from oil production can be exceptionally saline and contain a complex mixture of hydrocarbons, many of which are highly toxic. This study aimed to identify and characterize 141 halophilic bacteria isolated from production water and activated sludge from Marine Terminal Almirante Barroso (Brazil) and evaluate their potential for biosurfactant production and biodegradation of distinct petroleum hydrocarbons. Sequencing and phylogenetic analysis of the 16S rRNA gene revealed that the halophilic bacteria retrieved are distributed among 20 genera and four phyla (Proteobacteria, Firmicutes, Actinobacteria and Flavobacteria). RAPD fingerprinting was used to differentiate isolates at the infraspecific level, revealing 79 different genetic profiles. GC-MS analysis carried out with eight strains confirmed their ability to efficiently degrade alkanes and aromatic compounds under halophilic conditions, with preference for aromatic degradation. Eleven strains showed significant ability for reduction of the surface tension (from 72 to 40 mN/m) and for emulsification (up to 71%) of four different types of oils (mineral, soybean, diesel and kerosene). Results gathered in this study demonstrate a high taxonomic and genetic diversity of the halophilic bacterial strains isolated from the oil terminal samples and an outstanding potential for further use in biotechnological processes such as biosurfactant production or bioremediation. © 2016 Elsevier Ltd. All rights reserved.

Keywords: Biodegradation Emulsification Halophile Produced water Surface tension reduction Bacterial diversity

1. Introduction Oil is a complex mixture of organic and inorganic compounds and has as its most important constituent hydrocarbons, which can reach up to 98% of the total composition (Bícego, 1988). Oil composition varies significantly depending on the source reservoir and may contain sulfur, nitrogen, oxygen and metal compounds (Freedman et al., 1995; Marques et al., 2009). Polycyclic aromatic

* Corresponding author. E-mail address: [email protected] (M.B. Gomes). http://dx.doi.org/10.1016/j.ibiod.2016.08.014 0964-8305/© 2016 Elsevier Ltd. All rights reserved.

hydrocarbons (PAHs) found in oil can enter the environment through human and natural activities and are considered as environmental contaminants (Johnsen et al., 2005). PAHs are an increasing concern because of their toxic, mutagenic, and carcinogenic properties (Tang et al., 2005). Biodegradation of hydrocarbons is a widely known metabolic process that has been reported for many bacterial genera (Seo et al., 2009), including Halomonas (Wang et al., 2007), Alcanivorax (Yakimov et al., 1998), Marinobacter (Gauthier et al., 1992), Dietzia (Borzenkov et al., 2006), Bacillus (Kumar et al., 2007; Sass et al., 2008), Oleiphilus (Golyshin et al., 2002), Oleispira (Yakimov et al.,

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M.B. Gomes et al. / International Biodeterioration & Biodegradation 126 (2018) 231e242

2003) and Geobacillus (Chamkha et al., 2008), among others. The technology commonly used for hydrocarbon remediation includes mechanical, burying, evaporation, dispersion and washing procedures. The mechanisms employed for hydrocarbon removal from the environment rely on expensive, slow and inefficient methodologies (Mandri and Lin, 2007). Biological treatment or bioremediation is a desirable alternative due to its low cost and high efficiency (Stallwood et al., 2005; Karhu et al., 2009). Bioremediation uses microorganisms to degrade toxic pollutants and convert them into less toxic or harmless products, offering an environmentally safe and cost-effective technique (Kumar et al., 2011). In order to enhance the solubility and bioavailability of these hydrophobic compounds, many bacteria produce biosurfactants, thus facilitating their uptake and biodegradation (Bodour and Maier, 2002). Biosurfactants are distributed into various categories such as glycolipids, lipopeptides, polysaccharideeprotein complexes, phospholipids, fatty acids and neutral lipids (Cappello et al., 2012). The biosurfactants are excellent agents for emulsification, detergency, dispersion, microbial growth enhancement and metal sequestering (Pacwa-Plociniczak et al., 2011). These important characteristics make them suitable for use in oil recovery and bioremediation, with great potential for future replacement of the chemical surfactants (Ghanavati et al., 2008). Information on microorganisms involved with PAH biodegradation in moderate to high salinity environments in the last two decades is extensive. There is, however, little information on the ability of biosurfactant production in highly saline environments. In the present study, isolation, taxonomic identification and genetic characterization of halophilic bacteria from production water and activated sludge originated from a petroleum terminal were performed giving further assessment of their potential to biodegrade PAHs and produce biosurfactants under high salinity conditions. 2. Material and methods 2.1. Sampling, acclimation and bacterial isolation procedure The bacterial strains used in the present work were isolated from activated sludge of wastewater treatment and production water from Marine Terminal Almirante Barroso (TEBAR, S~ ao ~o, SP, Brazil). This site is located in the north coast of S~ Sebastia ao Paulo state and receives oil mixed with high salinity production water originated from offshore platforms. Sampling was performed by the technical staff of PETROBRAS. Two liters of activated sludge and production water were collected using sterilized plastic flasks and stored at 4 C for transportation to the laboratory in the Federal University of Viçosa (UFV), MG, Brazil. After arrival, bacterial isolation procedures were immediately performed. Aliquots of 1 mL of sludge and production water were serially diluted (101 to 108) in saline solution and 100 mL of each dilution were inoculated onto Petri dishes containing one of the four culture media: R2A (Difco) (Reasoner and Geldreich, 1985), MOD (Rohban et al., 2009), SAL (saline) (Pagaling et al., 2009) and MCAT (casamino acid) (Litchfield et al., 2009). Sodium chloride (4%) was added to all culture media in order to mimic the reactor condition to which microorganisms were already adapted to. Petri dishes were incubated at 28 C and monitored for colony growth. For sludge acclimation to high salinity condition, the media R2A (Difco) (Reasoner and Geldreich, 1985) and Moderate (MOD) (Rohban et al., 2009) were employed, since they allowed the recovery of a higher bacterial diversity in the isolation procedure. Five milliliters-aliquots of activated sludge were inoculated into 250 mL-Erlenmeyer flasks containing 50 mL of liquid medium

added of 6e20% NaCl. Flasks were incubated in rotational shaker at 28 C and 150 rpm. Every 6 days, acclimation cultures were transferred to fresh medium containing 1% higher salt concentration, until salt concentration reached 20%. During acclimation process, 100 mL-aliquots of each salt concentration culture were plated on the surface of solid MOD and R2A media. After growth, colonies with different morphologies were selected and submitted to streaking onto individual Petri dishes and incubation at 28 C until the appearance of new colonies. Each culture was submitted to stereomicroscope analysis, followed by Gram staining to verify purity of cultures. 2.2. Bacterial identification Genomic DNA extraction from all bacterial isolates was performed according to the protocol described by Soolinger et al. (1993). 16S rRNA gene was partially amplified from genomic DNA by PCR using the primer set 10f (50 GAG TTT GAT CCT GGC TCA G 30 ) and 1100r (50 AGG GTT GCG CTC GTT G 30 ) (Lane, 1991). PCR was performed in 25 mL-reaction mixtures containing 0.5 mM each primer, 0.2 mM dNTPs (Invitrogen), 1.5 mM MgCl2 (Invitrogen), 2.0 U Taq polymerase (Invitrogen) and 1.0x reaction buffer (Invitrogen) and 50e100 ng genomic DNA. Amplification was conducted using an Eppendorf Mastercycler Gradient (Eppendorf Scientific, New York, USA) and the program consisted of 1 cycle for denaturation at 95 C for 1 min, followed by 30 cycles of 1 min at 94 C for denaturation, 1 min at 55 C for annealing, and 3 min at 72 C for extension, with a final extension step of 3 min at 72 C. PCR products were purified using Illustra PCR DNA and Gel Band Purification Kit (GE Healthcare), and used as template for sequencing in the ABI3500XL (Applied Biosystem) sequencer with ABI Genetic analyzer BigDye Terminator cycle sequencing Kit (Applied Biosystems), buffer BigDye Terminator v1.1 (5 sequencing Buffer-Life Technologies) (0.1 M Tris-HCl, 0.5 mM MgCl2) and 3.2 pmol each primer (10f and 1100r). Partial 16S rRNA gene sequences obtained from isolates were assembled in a contig using the phred/Phrap/CONSED software (Ewing et al., 1998; Gordon et al., 1998). Sequences were compared against sequences of closely related type strains retrieved from the GenBank (http://www.ncbi.nlm.nih.gov) (Benson et al., 2013) database and RDP (Ribosomal Database Project, Wisconsin, USA, http://rdp.cme.msu.edu), using BLASTn and SequenceMatch routines, respectively. Sequences were aligned using the CLUSTAL X program (Thompson et al., 1997) and analysed with MEGA software v 6.06 (Tamura et al., 2013). Phylogenetic reconstruction was done with the neighbor-joining method (Saitou and Nei, 1987) and Kimura's two-parameter model (Kimura, 1980), with bootstrap values calculated from 1000 replicate runs. 2.3. RAPD fingerprinting The molecular technique RAPD (Random Amplified Polymorphic DNA) was employed in order to genetically differentiate the bacterial isolates belonging to the same species. Three primers of Set 100/1 (University of British Columbia, Vancouver, Canada) were used for typing the bacterial strains in RAPD independent reactions (Burgess et al., 2005), as follows: UBC # 2 (50 - CCT GGG CTT G e 30 ), # 4 (50 - CCT GGG CTG G e 30 ) and UBC # 25 (50 - ACA GGG CTC A - 30 ) for Proteobacteria, and UBC # 4 (50 - CCT GGG CTG G e 30 ), UBC # 18 (50 - GGG CCG TTT Ae 30 ) and # 25 (50 - ACA GGG CTC A - 30 ) for Actinobacteria and Flavobacteria. Twenty five microliterreaction mixtures contained 5 ng of genomic DNA, 2 U of Taq DNA polymerase (Invitrogen), 1X Taq buffer, 1.5 mM MgCl2, 0.2 mM of


dNTP mix (GE Healthcare) and 1 mM primer.The amplification program consisted of one cycle at 95 C for 2 min, 30 cycles of 30 s at 94 C, 30 s at 36 C and 1 min at 72 C and final extension cycle at 72 C for 3 min. The primers used were selected among those producing the most polymorphic profiles in a preliminary screening. The RAPD products were subjected to electrophoresis at 100 V for 2 h 30 min in 1.5% agarose gel stained with ethidium bromide. 2.4. Screening for microbial growth on hydrocarbons Screening assays for hydrocarbon utilization were developed according to the methodology described by Johnsen et al. (2002) based on the evaluation of bacterial respiration. Hexadecane (1%) (Vasconcellos et al., 2010); Phenol (0.02%) (Kafilzadeh et al., 2010); naphthalene (0.01%), phenanthrene (0.01%) and pyrene (0.01%) (Zhou et al., 2008); benzopyrene (0.005%) (Juhasz et al., 2000), were chosen as model compounds, according to previous literature, to evaluate the ability of bacteria to grow using them as sole carbon source. Bacterial isolates were pre-cultured in microplates (96 wells) containing 150 mL R2A broth and incubated in a rotational shaker at 30 C and 150 rpm for 24 h. After growth, aliquots (10 mL) of the bacterial cultures were transferred to another microplate containing 150 mL BH medium (Bushnell and Haas, 1940) added of the hydrocarbons described above. The bacterial strains E. coli EPI 300 (CBMAI 636) and Pseudomonas putida CBMAI 994 were used as negative and positive controls, respectively. After 48 h of incubation at 30 C and 150 rpm, 30 mL MTT [3-(4,5-dimethyl-2-thiazolyl)-2,5diphenyl-2 H-tetrazolium bromide] (Merck) solution (1 mg/mL) were added to each well to assess microbial respiration and consumption of hydrocarbons as sole carbon source. After 1 h of incubation at 30 C, cellular activity was indicated by the purple color of the culture medium (Bicalho et al., 2003; Vasconcellos et al., 2010). 2.5. Detection of catabolic genes PCR assays targeting the catabolic genes alkane monooxygenase (alk) and a subunit of aromatic ring hydroxylating dioxygenases (aARDHs) were performed for all bacterial strains showed to be genetically different by RAPD fingerprinting. Amplification reactions were carried out using two sets of degenerate primers (Table 1), in independent reactions, according to Kuhn (2007). Twenty five microliter-reaction mixtures contained 50e100 ng of genomic DNA, 2 U of Taq DNA polymerase (Invitrogen), 1X Taq buffer, 1.2 mM MgCl2, 0.2 mM of dNTP mix (GE Healthcare) and 1.2 mM each primer. PCR amplification was conducted using an Eppendorf Mastercycler Gradient (Eppendorf Scientific, New York, USA) and the amplification program consisted of one cycle at 97 C for 3 min, followed by 30 cycles of 1 min at 94 C, 1 min at 51 C for alk gene or 58 C for ARDH gene and 1 min at 72 C, with a final extension cycle of 5 min at 72 C. DNA from Rhodococcus erythropolis DSM 43066 and Pseudomonas putida CBMAI 994 was used as positive control for the alk and ARDH gene, respectively. The amplification products were checked by electrophoresis in 1.2% (wt vol-1) agarose gels.

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2.6. GC-MS analysis of hydrocarbon biodegradation In order to standardize the amount of inoculum to be used, bacterial isolates with positive growth using different hydrocarbons (as described in section 2.4) were pre-cultured individually in Erlenmeyer flasks containing 100 mL of R2A medium added of 9% NaCl, in triplicate, and incubated at 28 C, 150 rpm for 12 h. Spectrometry reading (l ¼ 600) was performed every 3 h (Cerqueira et al., 2011). In the biodegradation assay, 4 mL-aliquots of the bacterial growth were inoculated individually to a final concentration of 4 106 CFU/mL in Erlenmeyer flasks (125 mL) containing 36 mL of BH medium added of 9% NaCl. As the carbon source, hydrocarbons were added all together, as a cocktail, in the initial concentrations of 7.73 mg/mL hexadecane, 0.1 mg/mL naphthalene, phenanthrene and pyrene, 0.2 mg/mL phenol, and 0.05 mg/mL benzopyrene to a final volume of 45 mL. Assays were performed in triplicate and monitored for 20 days using GC-MS. Samples (triplicate) were taken every 5 days, totaling 5 sampling times (time zero, 5, 10, 15 and 20th day) for each bacterial isolate examined. Dimethylformamide solution (0.5 mg/mL) was used as internal standard. GC-MS analysis followed the conditions described by Passarini et al. (2011). 2.7. Screening for biosurfactant production Screening of biosurfactant-producing isolates was carried out using quantitative methods based on the reduction of the surface tension and emulsification assays. All methods were performed in triplicate using positive and negative controls (Mnif and Ghribi, 2015). Bacteria were cultured in R2A medium for 72 h at 28 C and 150 rpm in a rotational shaker (Innova® 44R, New Brunswick, USA). After growth, culture medium was centrifuged and the supernatant discarded. The resulting pellet was then diluted with distilled water and quantified by spectrophotometry to standardize the cell density (OD600) to approximately 0.100 ± 0.030. To stimulate biosurfactant production, an aliquot of 10% (v/v) of bacterial growth was transferred to Erlenmeyer flasks (50 mL) containing 27 mL of mineral medium and sucrose as carbon source (Paulo et al., 2012). The flasks were incubated at 28 C in a rotational shaker at 150 rpm for 7 days. Supernatant of bacterial cultures was then submitted to emulsification index (E24) and tensiometry assays by Du-Noüy-Ring Method (Walter et al., 2010). 2.8. Tensiometry assay The evaluation of the reduction of surface tension was performed over a period of 7 days. After growth, the bacterial cultures were centrifuged for 15 min at 13.000g (Eppendorf 5810R) and only the supernatant was evaluated for the reduction of surface tension values by Du-Noüy-Ring Method using a bench tensiometer Krüss K-20 EasyDine (Satpute et al., 2010). 2.9. Emulsification assay The emulsifying activity was determined using the methods described by Cooper and Goldenberg (1987) and Willumsen and

Table 1 Oligonucleotide primers used for PCR assays and target genes. Primer

Gene target

Sequence (50 / 30 )

Direction

Expected amplicon size (bp)

Reference

alkF alkR ARHD2 F ARHD2 R

Alkane monooxigenase (alk)

GCICAIGARITIRKICAYAA GCITGITGITCISWRTGICGYTG TTYRYITGYAIITAYCAYGGITGGG AAITKYTCIGCIGSIRMYTTCCA

forward reverse forward reverse

524

Kuhn et al. (2009)

329

Kuhn (2007)

a

ARHDs, a subunit (groups I, II, III and IVa) ARHDs, a subunit (groups I, II, III and IVa)

Classification of ARHDs according to Nam et al., 2001.

234


Karlson (1997), with modifications. To determine the emulsifying activity of bacterial isolate, 1 mL of the culture supernatant was added into the test tubes containing 1 mL of kerosene, diesel, mineral oil or soybean oil. After vortexing for 2 min, tubes were kept at rest. The emulsion stability was determined after 24 h, and emulsifying index (EI-24) was calculated from the ratio between the height of the emulsion layer by the total height of the liquid and multiplied by 100, according to the formula below. The emulsion was defined as stable when EI-24 50% or higher, as described by Plaza et al. (2006).

E24 ¼ h ðemulsionÞ=h ðtotalÞ 100 ¼ E24 %

2.10. Statistical analysis All analyses were performed in triplicate and the mean and standard deviation were calculated. The results were analysed statistically using the ASSISTAT software (version 7.7 beta, 2016) (http://www.assistat.com). The means were compared using oneway ANOVA and the Scott Knott test to indicate any significant difference among parameters and the variables. Results were considered significant when p < 0.01. 3. Results 3.1. Identification of bacterial isolates A total of 141 bacterial isolates from petroleum terminal e TEBAR (30 bacteria from production water, 33 from activated sludge and 78 from saline acclimation process) were identified by 16S rRNA gene sequencing. Phylogenetic analysis showed that 82% of bacterial isolates were affiliated to the phylum Proteobacteria, distributed among the classes Gammaproteobacteria (72%), Alphaproteobacteria (9%) and Betaproteobacteria (1%). The remaining isolates belonged to the phyla Actinobacteria (9%), Firmicutes (6%) and Flavobacteria (3%) (Fig. 1). Phylogenetic analysis revealed that the bacteria under study are related to 20 different genera and 30 different OTUs (operational taxonomic units defined at 97% sequence similarity) (Figs. 2 and 3). Many of the isolates were recovered in tight clusters with the type strain of the most related species (97% sequence similarity), supported by high bootstrap values (70%), and thus could be confidently identified at the species level, as summarized in Table 2. On the other hand, bacterial isolates that were recovered in tight clusters with more than one type strain, or grouped with related species with low bootstrap values (