A rapid method for the isolation of eicosapentaenoic acid-producing marine bacteria

A rapid method for the isolation of eicosapentaenoic acid-producing marine bacteria

Journal of Microbiological Methods 82 (2010) 49–53 Contents lists available at ScienceDirect Journal of Microbiological Methods j o u r n a l h o m ...

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Journal of Microbiological Methods 82 (2010) 49–53

Contents lists available at ScienceDirect

Journal of Microbiological Methods j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j m i c m e t h

A rapid method for the isolation of eicosapentaenoic acid-producing marine bacteria Jason Ryan a,b,⁎, Hannah Farr a,b, Sandra Visnovsky c, Mikhail Vyssotski a, Gabriel Visnovsky b a b c

Integrated Bioactive Technologies, Industrial Research Limited, PO Box 31-310, Lower Hutt 5040, New Zealand Chemical and Process Engineering, University of Canterbury, Private Bag 4800, Christchurch 8140, New Zealand The New Zealand Institute for Plant & Food Research Limited, Private Bag 4704, Christchurch 8140, New Zealand

a r t i c l e

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Article history: Received 10 December 2009 Received in revised form 7 April 2010 Accepted 7 April 2010 Available online 14 April 2010 Keywords: Eicosapentaenoic acid Marine bacteria Polyunsaturated fatty acids Screening Tetrazolium salts Triphenyltetrazolium chloride

a b s t r a c t Bacterial production of long chain polyunsaturated fatty acids (LC-PUFAs) is a promising biotechnological approach for the mass production of these valuable compounds, but extensive screening is currently needed to select a strain that meets industrial requirements. A method was developed for the rapid screening and isolation of eicosapentaenoic acid (EPA)-producing marine bacteria from mixed cultures using the dye 2,3,5-triphenyltetrazolium chloride (TTC). The method was first validated using two bacteria from the Shewanella genus, S. gelidimarina (known to contain EPA) and S. fidelis (known not to contain EPA), and subsequently applied to a range of bacterial samples collected from seven randomly selected New Zealand fish species. By incorporating TTC in both solid and liquid state fermentation treatments, a clear association between the reduction of TTC to the red-coloured triphenyl formazan (TF) and the presence of EPA within Gram-negative bacteria was confirmed. Incubation in 0.1% w/v TTC was optimal for colour response and cell growth in agar plates and liquid cultures. Bacteria that produce EPA reduced TTC to TF, but a number of non-EPA-producing bacteria also showed this capacity. By conducting a subsequent Gram staining, all EPA-producing strains were revealed to be G (−) rod bacteria while the non-producing ones were all G (+) cocci. The fatty acid methyl esters of the isolated bacteria that reduced TTC to TF were analysed using gas chromatography–mass spectrometry and the content of EPA was confirmed by gas chromatography. From a pool of 2.0 × 108 CFU/ml, this method allowed the rapid isolation of 16 bacteria capable of producing EPA. This new approach significantly reduces the number of samples submitted for GC analysis and therefore the time, effort and cost of screening and isolating strains of EPA-producing marine bacteria. © 2010 Elsevier B.V. All rights reserved.

1. Introduction Polyunsaturated fatty acids (PUFAs), such as eicosapentaenoic acid (20:5ω3, EPA), are important for human health. They are the precursors of many essential regulatory molecules in the body as well as being important lipid components of brain and retina cell membranes. They also help to prevent cardiovascular disease (Das, 2008; Shin et al., 2007). The primary source of these supplements has been fish oil. However, fish oil sources have many disadvantages: they can be contaminated by heavy metals, retain a “fishy” odour in the final product, and are processed from the world's declining fish stocks (Hinzpeter et al., 2006; Tocher, 2003). Gram-negative marine bacteria, particularly those found in the gut flora of fish in deep, low temperature waters, produce these fatty acids (Kawamoto et al., 2009; Russell and Nichols, 1999; Valentine and Valentine, 2004), and its growth in large-scale fermentation processes

⁎ Corresponding author. Integrated Bioactive Technologies, Industrial Research Limited, PO Box 31-310, Lower Hutt 5040, New Zealand. Fax: + 64 4 9313 055. E-mail address: [email protected] (J. Ryan). 0167-7012/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2010.04.001

presents a renewable, sustainable, and contaminant-free source of PUFAs for use in human and animal dietary supplements (Berge and Barnathan, 2005). Sources for the isolation of EPA-producing bacteria use either environmental samples, such as free sea water, sea ice and sea sediments or marine organisms (Bowman et al 1997; Cho and Mo, 1999; Gentile et al., 2003; Ivanova et al., 2003b; Yazawa, 1996) and the screening of specific fish such as blue-backed fish or deep sea fish, has been suggested as the most promising approach to obtain high yields of these particular bacteria strains (Kawamoto et al., 2009; Valentine and Valentine, 2004; Yazawa, 1996). Methods that search for bacteria that produce EPA have been conducted using standard techniques, consisting in culturing samples on marine agar plates, isolating single clones and regenerating from them biomass for EPA content analysis (Bowman et al 1997; Gentile et al., 2003; Ivanova et al., 2003b). A range of methods for the screening of EPA-producing bacteria have been used once the bacteria has been isolated, such as thin layer chromatography (Cho and Mo, 1999), gas chromatography (GC) (Yazawa, 1996) and gas chromatography–mass spectrophotometry (GC–MS) (Gentile et al., 2003). However, and while the methods are accurate, they require significant

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time and resources since each bacteria isolated must be treated individually. Other methods, using enzyme functionality and growth temperatures, have been used to identify fungal PUFA contents. Fungi are a known source of PUFAs, mainly the omega 6 fatty acid, arachidonic acid (20:4ω6, AA) (Zhu et al., 2004). One of these methods consists in the addition of O-acetylsalicylic acid to agar media to produce the inhibition of the growth of fungi that produce AA (Botha et al., 1999; Eroshin et al., 1996), but the approach requires a control system for positive selection. The incubation of environmental samples at low temperature (0–5 °C) has been used to select fungi that produce AA (Carreiro and Koske, 1992; Zhu et al., 2004) however, the low temperatures required lead to significant growth periods (8–15 days). Tetrazolium salts have been used as indicators of metabolic activity of living cells (Beloti et al., 1999; Matalon and Sandine, 1986) in a range of applications, including the detection of hydrocarbon-oxidizing bacteria in oil-contaminated water and soil specimens (Petrović et al., 2008), the measurement of bacterial growth in clinical diagnostic (Tengerdy et al., 1967), and the identification of Gram-negative pathogenic bacteria (Velegraki and Logotheti, 1998). When exposed to mild reduction conditions the water soluble tetrazolium salts are reduced to water insoluble, strongly coloured formazans (Marshall et al., 1995). The tetrazolium salt, 2,3,5-triphenyltetrazolium chloride (TTC) is colourless, but is reduced to the bright triphenyl red formazan (TF) (Zhivich et al., 1991). Low concentrations of TTC may not provide sufficient colour development and high concentrations may inhibit bacterial growth (Matalon and Sandine, 1986). Mitochondrial dehydrogenase has been strongly associated with this mechanism. However, Marshall et al. (1995) suggested that other intracellular systems are just as able to donate electrons to tetrazolium salts. Zhu et al. (2004) developed a method that combined low temperature and TTC to rapidly identify increased production of AA in the fungi Mortierella alpina, finding a positive correlation between the TF formation and whole cell AA content. The underlying reason for this correlation is unknown, but by comparing the fatty acids in two Mortierella species (M. alpina and M. mucor) and their ability to reduce TTC, Zhu suggested that the enzyme Δ5-desaturase, part of the fatty acid synthase pathway and a form of dehydrogenase responsible for the reduction of dihomo-γlinoleic acid to AA, could be also responsible for reducing TTC to TF (Zhu et al., 2004). Gram-negative marine bacteria, particularly those found in the gut flora of fish in deep, low temperature waters, produce fatty acids using similar de novo biosynthetic pathways, such as fatty acid synthase pathways for the synthesis of monoenoic fatty acids (Valentine and Valentine, 2004; Watanabe et al., 1997) and polyketide synthase pathways for the synthesis of PUFAs (Kawamoto et al., 2009; Ratledge, 2001; Russell and Nichols, 1999; Valentine and Valentine, 2004). As the literature shows, no methodical approach for the primary selection of EPA-producing bacteria has been developed yet. Based on Ratledge and Zhu's findings (Ratledge, 2001; Zhu et al., 2004), we report here on studies to develop a systematic method using TTC for the primary screening and rapid isolation of EPA-producing marine bacteria. 2. Materials and methods 2.1. Bacterial strains, primary screen, and culture conditions Two Gram-negative rod-shaped bacteria, Shewanella gelidimarina (ATCC 700752) and Shewanella fidelis (BAA-318), used as EPA (+) and EPA (−) controls, respectively, were obtained from American Type Culture Collection (ATCC). S. gelidimarina was originally isolated from Antarctica and contains EPA concentrations from 11 to 17% of total fatty acids (Bowman et al., 1997), while S. fidelis was originally isolated from the South China Sea and does not contain EPA (Ivanova

et al., 2003a). EPA-producing bacteria were isolated from bluenose (Hyperoglyphe antarctica), black cardinal (Epigonus telescopus), snapper (Pagrus auratus), tarakihi (Nemadactylus macropterus), jack mackerel (Tracherus declivis), scorpionfish (Helicolenus barathri) and blue warehou (Seriolla brama). All the isolated strains, and EPA (+) and (−) controls, were maintained in 30% glycerol at −80 °C in Industrial Research Ltd's (IRL) culture collection. All fish specimens were captured in New Zealand sea waters directly or obtained from a local supplier, and processed immediately after their capture. To isolate the EPA-producing bacteria, the stomach and intestines were first excised from the fish into sections no longer than 5 cm. Each separate section was then placed in a 250 ml Erlenmeyer flask containing 100 ml of PYM medium and incubated for 1 day at 15 °C. This temperature has been proven optimal for the growth of EPAproducing bacteria (Yazawa, 1996). A three-fold dilution was obtained from the sample and 10 μl was streaked on to marine agar containing 0.1% w/v TTC, and incubated at 15 °C for 2–4 days. Bacterial colonies were isolated from these plates and re-cultured on to marine and nutrient agar plates to check the purity of the strains and for fatty acid analysis. Gram stain tests were conducted on all colonies that reduced TTC to TF. Gram stain tests were performed as described by Simbert and Kreig (1994). All colonies tested were submitted to GC to confirm the presence or absence of EPA. Isolated EPA-producing bacteria were named consecutively IRL 544 to 552 and IRL 563 to 569, and stored in IRL's culture collection. The composition of the peptone-yeast extract-meat extract (PYM) medium used was 10 g yeast extract, 2.5 g meat peptone, 5 g bactopeptone, 20 g glucose and 15 g sea salt in 1 l of RO water (Yazawa, 1996). The Luria Bertani (LB) medium used was 10 g bacto-tryptone, 5 g bacto-yeast extract and 10 g sodium chloride in 1 l of RO water. The marine and nutrient agars used were Diffco 2105 and Diffco 297801, respectively, and both were prepared following the supplier's instructions. Marine agar with TTC was prepared as per standard marine agar, with the addition of sterile filtered TTC to a final concentration of 0.1% w/v before pouring the agar plates. 2,3,5triphenyltetrazolium chloride was obtained from Sigma-Aldrich. TTC reduction to TF was also checked in liquid culture using PYM medium. Two sterile 50 ml polystyrene tubes (Corning) were filled with the medium and seeded with a single colony of S. gelidimarina and S. fidelis. The cultures were incubated at 15 °C for 3 days, and then added with TTC to a final concentration of 0.1% w/v. The cultures were then incubated for an additional hour at the same temperature and the results evaluated. 2.2. Analysis of EPA content Fatty acid methylation was conducted as described by Svetashev et al. (1995). Five of the sixteen isolated EPA-producing marine bacteria were randomly selected and their EPA content determined. S. gelidimarina and S. fidelis were used as EPA (+) and EPA (−) controls, respectively. A sample of biomass from each plate was removed and placed in a pre-weighed vial with the help of a sterile 10 μl plastic loop. The vial was then reweighed and the biomass to be extracted calculated. To this, 10 μl of a 1 μg/μl solution of 23:0 (used as an internal standard for methylation and concentration) and 0.5 ml of 1% sodium base reagent were added, and incubated at 80 °C for 30 min. After cooling to room temperature, 0.5 ml of 5% methanolic HCl was added to the vial, repeating the incubating cycle. The fatty acid methyl esters (FAME) were then extracted into 1 ml of hexane. Hexane was evaporated under argon gas and then the FAME residue was redissolved into 50 μl chloroform, obtaining a sample ready for gas chromatography (GC) analysis (Svetashev et al., 1995). GC analysis was performed on an Agilent Technologies 6890N Network GC system equipped with a flame ionization detector. The gas chromatography capillary column was BP-20 (WAX) (30 m length,

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with an internal diameter of 0.32 mm, and 0.25 μm film phase) (SGE International Pty Ltd, Australia). A 1 µl sample was injected into the GC using an Agilent 7683 series autosampler. The injector and the flame ionization detector temperature were both 250 °C, and the column temperature was held at 195 °C for 45 min. The carrier gas (hydrogen) pressure was 0.379 MPa, with a flow rate of 46.6 ml/min. The split ratio was 30:1. The hydrogen and air flow rates were 40.0 ml/min and 450 ml/min, respectively, and the make-up flow rate (nitrogen) was 20.0 ml/min. The chromatograph was integrated using Agilent Technologies Enhanced ChemStation software. Individual peaks of FAME were identified by comparison with standards of FAME and by equivalent chain length (ECL) values (Stransky et al., 1997). The 23:0 standard was used to determine total lipid mass, and the relative concentrations of each fatty acid was determined by comparing peak areas. The tests were conducted in triplicate. Gas chromatography–mass spectrometry (GC–MS) experiments were performed on a Hewlett-Packard HP-6890 chromatograph linked to a HP-5973 mass spectrometer using a HP5-MS column (29 m × 0.25 mm × 0.25 µm). The carrier gas was helium. The temperatures of injector and source were held at 260 °C, and the temperature of the GC–MS interface was held at 290 °C. The column temperature was programmed from 160 to 300 °C at 3 °C/min. The chromatograph was integrated using Agilent Technologies Enhanced ChemStation software. 2.3. Determination of the optimum TTC concentration Two of the EPA-producing marine bacterial strains isolated with known EPA content, IRL 551 and IRL 569, and S. gelidimarina and S. fidelis as EPA (+) and EPA (−) controls, respectively, were used to determine the optimum TTC concentration for both solid and liquid cultures. The bacteria were grown in standard PYM medium at 15 °C for 24 h and then the broth removed to generate a 1 ml solution at an optical density (OD) ≤1.000. OD was determined using a UV 160A spectrophotometer (Shimadzu) and the wave length calibrated at 600 nm. A 1% w/v stock solution was used to provide a range of TTC concentrations (0.0025, 0.005, 0.01, 0.05, 0.1, and 0.25% w/v) in the fermentation broth, and the solutions were incubated for 30 min at room temperature. After incubation, 1 ml of dimethyl sulfoxide (DMSO) and 8 ml of ethyl acetate were added to the biomass and vortexed for 1 min. The mixture was then centrifuged at 2576 g for 30 s, and the top organic layer containing the extracted TF removed and its absorbance measured. The degree of staining was quantified by measuring the absorbance of TF in ethyl acetate at 485 nm. Tests were conducted in triplicate. 2.4. DNA isolation and PCR on selected isolates DNA isolation and PCR analysis were only conducted on the five EPA-producing bacteria where EPA content was determined. Bacterial isolates were selected from marine agar plates and incubated in LB broth at 21 °C for 20 h. Aliquots of 1.5–3 ml of culture were centrifuged at 4500 g for DNA extraction. Genomic DNA was isolated using the Qiagen DNeasy Blood and Tissue kit for Gram-negative bacteria following the manufacturer's instructions. Amplification of the 16S rRNA region was performed by PCR using the universal primer pair U16A and U16B (Wang and Wang, 1996), giving an amplicon of 1.5 kb. Each reaction was carried out in a total volume of 25 μl containing 1 μl of each primer (5 μM), 2 μl of dNTPs (2 mM), 2.5 μl of 10 × buffer, 1 μl of 50 mM MgCl2, 0.10 μl of Taq polymerase (5 U/μl Invitrogen and 1 μl of template DNA 10–20 ng/µl) in sterile water. Amplification was performed in a GeneAmp-PCR system 9700 (Applied Biosystems) thermocycler using the following conditions: 2 min for initial denaturation at 94 °C followed by 30 cycles of 94 °C for 30 s, 55 °C for 30 s and 72 °C for 1 min, and a final extension step of 72 °C for 7 min. PCR products were separated in a 1% agarose gel by

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electrophoresis and visualized using a Multi Doc-It Digital Imaging System, UVP (Bio-Strategy). PCR products were purified using a Qiagen PCR purification kit following the manufacturer's instructions. Sequencing of the 16S rRNA region was performed by Macrogen Ltd (Seoul, Korea) using the primers named above. DNA sequence output was edited with Sequencher (Gene Codes) and compared to the GenBank nucleotide database using BLASTN (Altschul et al., 1990). All bacteria shared 99% 16S rRNA gene nucleotide sequence similarity to sequences of the genera Photobacterium, Vibrio or Shewanella present in GenBank. Sequences were submitted to GenBank and were granted Accession Numbers (Table 1).

3. Results and discussion 3.1. Development of the screening method for selecting EPA-producing bacteria To observe the effects of TTC on cell growth, S. gelidimarina (EPA positive control) and S. fidelis (EPA negative control) were incubated on nutrient agar and marine agar plates containing 0.1% w/v TTC (Velegraki and Logotheti, 1998). S. gelidimarina grew well on both nutrient agar and TTC marine agar. S. fidelis grew moderately well on nutrient agar, but no growth was observed on TTC marine agar (Fig. 1). The description of S. gelidimarina grown on nutrient and marine agar was consistent with published data (Bowman et al., 1997). A positive association was observed between the ability to grow on agar plates containing TTC, the ability to reduce TTC to TF, and the ability of bacteria to produce EPA. While there are no published data reporting the growth of S. gelidimarina in a culture medium containing TTC, the development of the red colour on the colonies confirms the reduction of TTC to TF and correlates well with the findings of Zhu et al. (2004). They observed a similar result when identifying the increased production of a different PUFA, arachidonic acid, in the fungi Mortierella alpina, and suggested that the enzyme Δ5-desaturase could be the responsible for the reduction of the colourless TTC to the bright TF. Some marine bacteria use the same enzyme in their metabolic pathway to produce ω3 PUFAs, like EPA, converting eicosatetraenoic acid (ETA) to EPA with Δ5-desaturase (Ratledge, 2001). To confirm whether TTC is also reduced to TF in liquid phase, S. gelidimarina and S. fidelis were cultivated in PYM medium for 3 days, and then TTC was added to a final concentration of 0.1% w/v TTC. The culture medium in the tube containing the EPAproducing strain (S. gelidimarina) developed an intense red colour within the first 10 min of incubation while the one in which the nonEPA-producing bacteria (S. fidelis) was cultivated remained unchanged after 60 min of incubation (Fig. 2). This result confirmed the trend observed previously when the tetrazolium salt was incorporated into agar plates and used as a base to optimise the TTC concentration. Table 1 Strain identification for isolated strains used to investigate EPA content. Strain

IRL number

GenBank accesion number

Source organism

Photobacterium sp.

IRL 544

GQ414574

Shewanella sp.

IRL 551

GQ414575

Vibrio sp.

IRL 552

GQ414576

Shewanella halifaxensis Shewanella marintestina

IRL 566

GQ414577

IRL 569

GQ414578

Tarakihi (Nemadactylus macropterus) Jack mackerel (Trachurus novaezelandiae) Scorpionfish (Scorpaena cardinalis) Blue warehou (Seriolella brama) Blue warehou (Seriolella brama)

All sourced species used were New Zealand salt water fish.

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producing bacteria are Gram negative (Kawamoto et al., 2009; Russell and Nichols, 1999). The EPA-producing strains were named consecutively IRL 544 to 552 and IRL 563 to 569, and stored in IRL's culture collection. In addition, 5 of the 16 EPA-producing marine bacteria isolated were randomly selected and their EPA content analysed by chromatography–mass spectrometry. Results confirmed the presence of EPA on the analysed isolates, which ranged from 1.6 to 7.0% of total fatty acids (Table 2). 3.3. Determination of the optimum TTC concentration

Fig. 1. S. fidelis (nutrient agar bottom left, TTC marine agar top left) and S. gelidimarina (nutrient agar bottom right, TTC marine agar top right) grown at 15 °C for 1 week.

3.2. Screening of intestinal marine bacteria from fish A single specimen of bluenose, cardinal, snapper, tarakihi, jack mackerel, blue warehou and scorpionfish was used to confirm the results obtained with the TTC screening method. After processing each fish, marine agar plates with and without TTC were seeded and incubated as described in the methods section. From a pool of 2.0 × 108 CFU/ml of mixed flora, only 36 bacteria were capable to reduce TTC to TC, and only 16 of them showed the ability to produce EPA by gas chromatography (GC). By conducting a subsequent Gram staining, all EPA-producing strains were revealed to be G (−) rod bacteria, while the non-producing ones were all G (+) cocci. These data correlated well with other published data that states all EPA-

The initial concentration of TTC used to screen EPA-producing bacteria was 0.1% w/v. This concentration was selected based on the work of Velegraki and Logotheti (1998) who used TTC to identify G (−) pathogenic bacteria. To confirm that this concentration was optimal, two of the EPA-producing marine bacteria isolated previously with the lowest (IRL 559) and highest (IRL 569) EPA contents (Table 2), and S. gelidimarina and S. fidelis as EPA (+) and (−) negative controls, respectively, were incubated in PYM media containing TTC concentrations ranging from 0 to 0.25% w/v and the values of absorbance were read after 30 min of incubation. A maximum absorbance indicating the TTC to TF reduction response was observed in all samples at 0.01%, followed by 0.05% TTC (Fig. 3). However, when 0.01% TTC was added to agar plates to screen EPA-producing bacteria, the number of non-reducing, white bacterial colonies increased significantly. A similar result was observed when 0.05% TTC was incorporated into agar plates. When colonies were transferred from either 0.01 or 0.05% TTC to 0.1% TTC, only pink–red colonies that showed TF formation grew well (EPA positive), while those that did not produce TF (white colonies, EPA negative) did not grow. A higher concentration of TTC (0.25%) resulted in poor reduction response (low absorbance values) and cell growth inhibition. When TTC 0.1 and 0.25% concentrations on marine agar plates were compared using the same source material, there were significantly more TTC-reducing colonies on 0.1% than on 0.25% TTC. Based on these results, 0.1% TTC was selected as the optimal screening concentration. In preliminary experiments conducted using the same EPAproducing isolated and the EPA (+) and (−) controls, there was no correlation between the reduction of TTC to TF and the relative EPA content of the bacteria based on absorbance at 485 nm (data no shown). Bacteria with high EPA contents did not have a higher absorbance than strains producing lower amounts of EPA when incubated with the same TTC concentration and temperature conditions. It seems, therefore, that this technique can only be used to distinguish between EPA-producing and non-producing bacteria, and cannot be used to compare the relative EPA content of two distinct bacteria through the intensity of the red colour developed. There may be a positive correlation between EPA content and colour development, but only when considering one isolate independently. Therefore, a quantitative approach such as that used by Zhu et al. (2004) cannot be used as a primary screen. The effect of tetrazolium

Table 2 EPA content from 5 of 16 EPA-producing marine bacteria isolated.

Fig. 2. S. fidelis (left) and S. gelidimarina (right) after 1 h incubation at 15 °C in PYM medium added with 0.1% w/v TTC. Cultures were previously grown in PYM for 3 days at the same temperature.

Strain

EPA content (%)

IRL 551 (Shewanella sp.) IRL 552 (Vibrio sp.) IRL 544 (Photobacterium sp.) IRL 566 (Shewanella sp.) IRL 569 (Shewanella sp.) S. gelidimarina S. fidelis

1.6 ± 0.3 2.3 ± 0.3 4.6 ± 0.7 5.5 ± 0.2 7.0 ± 0.2 7.3 ± 0.2 0.0

S. gelidimarina and S. fidelis were used as EPA positive and negative controls, respectively. Tests were conducted in triplicate. EPA content is represented as a percentage of total fatty acids.

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Fig. 3. Determination of the optimum TTC concentration. Actively growing samples of Shewanella sp. (IRL551) ( ), Shewanella marintestina (IRL569) (□), Shewanella gelidimarina (ATCC 700752) ( ),and Shewanella fidelis (BAA-318) ( ) were incubated at various TTC concentrations for 30 min. The measured absorbance relates to the relative presence of TF in each bacterial sample. Each point represents the average of three readings and the error bars the standard deviation.

dyes on different genera/species is difficult to predict as dye reduction is known to differ with cell type (Berridge et al., 2005). The correlation discovered by Zhu et al. (2004) may only hold true for strains of the fungi M. alpina and not for the many bacteria screened by our method. However, further research needs to be done over a wider range of EPA-producing bacteria to corroborate this. 4. Conclusions Using TTC as the primary test for EPA-producing marine bacteria resulted in the development of a rapid method for screening and isolating these bacteria. A direct association between the ability to grow on agar plates containing TTC, the ability to reduce TTC to TF, and the ability of bacteria to produce EPA was found. EPA-producing bacteria were isolated without the need for GC analysis from a large pool of microorganisms. Identification of some of the isolated bacteria showed that they belonged to the Shewanella, Photobacterium, and Vibrio genera, indicating that the screening method was not specific to a single genus. This new approach has the advantages of significantly reducing the number of samples submitted for GC analysis and therefore reducing the time, effort and cost involved in the screening and isolation of EPA-producing marine bacteria strains. Further improvements to this method, such as the addition of compounds to the culture medium to avoid the growth of Gram positive bacteria, and the optimization of the incubation temperature to improve the reaction kinetics, could be explored to improve its selectivity. Acknowledgements The authors are grateful to Dr. Eduard Nekrasov and Dr. Andrew MacKenzie (Industrial Research Limited, New Zealand) for assistance with GC techniques, and Fanny Dheilly and Laurent Chastanet (Polytech Clermont-Ferrand, France) for their assistance with bacterial isolation. We also thank the valuable contribution of Prof. Ken Morison (University of Canterbury, New Zealand) and Dr. Andrew Pitman (Plant & Food Research, New Zealand) for reviewing the manuscript. This research was supported by the Foundation for Research, Science and Technology (C08X0709). References Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403–410.

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