Skip to main content
Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Enhanced eicosapentaenoic acid production by a new deep-sea marine bacterium Shewanella electrodiphila MAR441T

  • Jinwei Zhang,

    Roles Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation, Writing – original draft, Writing – review & editing

    Affiliations Institute of Biomedical and Clinical Sciences, University of Exeter Medical School, Hatherly Laboratory, Exeter, United Kingdom, School of Natural and Environmental Sciences, Newcastle University, Newcastle upon Tyne, United Kingdom

  • J. Grant Burgess

    Roles Conceptualization, Funding acquisition, Supervision

    Affiliation School of Natural and Environmental Sciences, Newcastle University, Newcastle upon Tyne, United Kingdom


Omega-3 fatty acids are products of secondary metabolism, essential for growth and important for human health. Although there are numerous reports of bacterial production of omega-3 fatty acids, less information is available on the biotechnological production of these compounds from bacteria. The production of eicosapentaenoic acid (EPA, 20:5ω3) by a new species of marine bacteria Shewanella electrodiphila MAR441T was investigated under different fermentation conditions. This strain produced a high percentage (up to 26%) of total fatty acids and high yields (mg / g of biomass) of EPA at or below the optimal growth temperature. At higher growth temperatures these values decreased greatly. The amount of EPA produced was affected by the carbon source, which also influenced fatty acid composition. This strain required Na+ for growth and EPA synthesis and cells harvested at late exponential or early stationary phase had a higher EPA content. Both the highest amounts (20 mg g-1) and highest percent EPA content (18%) occurred with growth on L-proline and (NH4)2SO4. The addition of cerulenin further enhanced EPA production to 30 mg g-1. Chemical mutagenesis using NTG allowed the isolation of mutants with improved levels of EPA content (from 9.7 to 15.8 mg g-1) when grown at 15°C. Thus, the yields of EPA could be substantially enhanced without the need for recombinant DNA technology, often a commercial requirement for food supplement manufacture.


Omega-3 polyunsaturated fatty acids (PUFAs), such as eicosapentaenoic acid (EPA, 20:5ω3) and docosahexaenoic acid (DHA, 22:6ω3), are important to human health. Nutritional insufficiencies of omega-3 PUFAs may have adverse effects on brain development and neurodevelopmental outcomes [1]. Omega-3 PUFAs are considered as therapeutic options which may reduce secondary neuronal damage initiated by traumatic brain injury [2]. They also have a protective role in age-related macular degeneration, and can prevent the harmful effects of chronic stress [3], and may contribute to the prevention of cognitive decline [4]. In addition, benefits for bone health and turnover have been reported [5]. Omega-3 fatty acids also play an important role in the modulation and prevention of heart failure and cardiovascular disease [6], and reduce the risk of development or progression of Alzheimer's disease [7].

Currently, the most prominent dietary sources of EPA and DHA are fish oil, supplies of which are unable to meet increasing global demand. In addition, fish may contain mercury, polychlorinated biphenyls, and other contaminants that have adverse effects on humans, particularly the developing foetus [8]. Furthermore, there are other potential problems associated with fish oils as a source of PUFA, such as: taste, odor, chemical stability as well as coextracted contaminants [9]. Nevertheless, the main problem of fish oil as a source of omega-3 fatty acids is its sustainability due to the worldwide decline of fish stocks [10]. Alternatively, natural production of omega-3 fatty acids by marine microbes is a potential alternative source, which would also be suitable for vegetarians. Furthermore, most bacteria can be grown on waste nutrients from other agricultural or industrial processes. Therefore, dietary sources of EPA and DHA from microbial biomass appear to be particularly promising and an area in which further research is warranted.

So far, bacterially derived PUFAs were reported mainly from Gram-negative bacteria [11], from two bacterial phyla: the Gammaproteobacteria (e.g., Shewanella, Moritella, Colwellia, Alteromonas, and Photobacterium) and the Bacteroidetes (e.g. Flexibacter and Psychroserpens) [12]. These EPA/DHA producing bacteria include psychrophiles and piezophiles, and were isolated from polar regions and the deep sea [1319], as well as mesophiles isolated from a temperate estuary [20] and from shallow seawater samples [2,2123]. However, it is unclear why these bacteria produce omega-3 fatty acids. Production of EPA by some bacteria increases as temperature decreases, leading to the hypothesis that these molecules may be important for growth at low temperatures [11,14,24]. Cells must cope with decreases in temperature by modulating the lipid composition of their membrane, which can crystallize or enter nonbilayer phases at low temperatures [25]. EPA was not required for low-temperature growth in the deep-sea bacterium Photobacterium profundum [26], but it may be required for low temperature growth in Shewanella [11,18,27]. Bacteria adjust their membrane lipid composition by modifying certain types of fatty acids to maintain membrane viscosity in response to environmental changes, such as temperature, pressure or salt concentration.

[28]. A number of studies are available on the physiology of bacterial PUFA production under varying culture conditions [13,28,29]. These studies indicate that PUFA biosynthesis could be manipulated by changing environmental conditions. Metabolic engineering of bacterial PUFA gene clusters into E. coli as a host, has also been extensively studied for the production of PUFA, however, the yield of EPA/DHA reported was low (e.g. 2.2mg/g EPA and 4.1% TFA) [4,30]. Thus, continued research utilizing a variety of bacterial strains is warranted to more fundamentally understand PUFA biosynthesis and to continue to explore non-recombinant natural sources of microbially produced omega-3 fatty acids.

We recently identified a psychrophilic strain Shewanella electrodiphila MAR441T that was isolated from sediment from a Mid-Atlantic Ridge (MAR) non-vent site at a depth of 2,500 m. This species was capable of producing relatively high yields of EPA (up to 24 mg g-1) when grow at all growth temperatures from 4 to 25°C [31]. In the current study, we have extended this work by significantly improving production of PUFA by varying the carbon source and using artificial sea water with different concentrations of Na+, time of culture and temperature. In addition, cerulenin treatment and chemical mutation by N-methyl-N′-nitro-N-nitrosoguanidine (NTG) were used in order to enhance secondary metabolite production of PUFAs. The acyl chain speciation of the major phospholipid classes and non-esterified fatty acid (NEFA) fraction produced by strain MAR441T are also described.


Cultivation conditions for biomass

For biomass production, strains were inoculated into 10 ml of Zobell’s broth (ZB) [32], and incubated at 15°C until turbidity was apparent by optical density (OD) 600 nM. The 10 ml cultures were then used to inoculate 90 ml volumes of marine broth (MB) contained in 500 ml conical flasks pre-rinsed with chloroform. Flasks were incubated at 15°C (or indicated temperatures) with agitation provided by a magnetic stirrer or orbital shaker (180 rpm) for 24–48 h until sufficient mass of estimated late-log phase cells were present for harvest. Cell mass from broth cultures was collected by centrifugation at 4500 g for 20 min. Cell pellets were resuspended in 100 ml M9 solution (22mM KH2PO4, 22mM Na2HPO4, 85mM NaCl, 1mM MgSO4) and recentrifuged followed by rinsing with 0.1% ammonium acetate and frozen. The washed cell pellets were suspended in 2.0 ml saline and lyophilised in pre-weighed containers prior to lipid extraction as descripted previously [31]. The collected samples were then stored at -20°C followed by -80°C, for fatty acids analysis.

The effect of temperature

The temperature-growth response (4–30°C) of strain MAR441T was studied by growing in marine broth (MB). Growth was observed and samples collected every day for five days, centrifuged and washed with sterile M9 solution. The generation time was used to evaluate the cell growth in the exponential phase according to previous methods [33].

Effect of carbon and nitrogen source

Strain MAR441T was grown on various sole carbon sources (L-alanine, L-leucine, L-proline, L-serine, propionic acid, glucose, glycerol, Tween 80, 60 and 40) and nitrogen sources (Urea and (NH4)2SO4) in triplicate or duplicate (see Supplementary S2 Table). Negative Control (NC) medium contained 0.01% (w/v) yeast extract in seawater. Modified media for sole carbon/nitrogen source cultures was 0.5% (w/v) carbon/nitrogen source and 0.01% (w/v) yeast extract in 0.22 µm filtered and sterilized natural seawater [34]. Optimal Medium (OM) contains 0.5% (w/v) Proline and 0.5% (w/v) (NH4)2SO4) in NC; Cultures were incubated at 15°C, in triplicate in 50-ml ZB contained within 200-ml flasks pre-rinsed in chloroform with shaking (180 rpm) for 36–48 h until sufficient mass of estimated late-log phase cells was present for harvest.

Cultivation conditions with treatment of cerulenin

Ten ml seed cultures of MAR441T were used to inoculate 90 ml volumes of marine broth contained in 500 ml conical flasks, where the antibiotic cerulenin (Merck), in 50% (v/v) ethanol (1 mg ml-1) was added at various concentrations (0, 0.5, 1, 2.5, 5 and 7.5 µg ml-1) prior to cultivation. Flasks were incubated at 4 and 15°C respectively. The growth of cells was monitored turbidometrically at 600 nm.

Induction of mutations with nitrosoguanidine (NTG)

N-methyl-N’-nitro-N-nitrosoguanidine (NTG) purchased from Tokyo Kasei Kogyo Co., Ltd. Japan, was used as chemical mutagen. Chemical mutagenesis was carried out according to the literature (Kotchoni et al. 2003; Liu et al. 2004). Exponentially growing cells of strain MAR441T were harvested from 3 ml of fermentation broth by centrifuging at 1600 g for 10 min at room temperature. The pellet was washed twice with 0.85% NaCl solution and then resuspended in 5 ml 0.1 M phosphate buffer, producing a suspension containing ~105 cfu l−1 (colony forming units l broth−1). The cells were then exposed to two NTG concentrations (300 µg l−1 and 500 µg l−1) for 12 h at 4 and 15°C by adding appropriate volumes of NTG stock (720 µg l−1 NTG in 0.1 M phosphate buffer, pH 6.5) to the cell suspensions. 1 ml samples of the serially-diluted culture were then spread on agar plates for mutant discovery. The agar plates were incubated for 2 days at 4 and 15°C respectively, and the resulting colonies (more than 1000 colonies) were taken off the agar plates by random selection based on morphology (e.g. size and colour), 50 colonies were further chosen for evaluation of their ability to produce PUFAs. The transformation experiments involving the selected mutants were conducted using the same culture methods used for the parent strain.

Scanning electron microscopy

Cells were processed for scanning electron microscopy (SEM) (Cambridge Stereoscan 240). The samples were fixed in 2% glutaraldehyde in 0.2 M Sorenson’s phosphate buffer (pH 6.8) for 12 h, then rinsed in Sorenson’s phosphate buffer twice for 15 mins and dried in an alcohol series up to 100%. The samples were then CO2-critical point dried by a Samdri 780 Critical Point Dryer, mounted on an aluminium stub with Achesons Silver ElectroDag and coated with 15 nm gold/palladium (40/60) using a polaron SEM coating unit and then observed using SEM.

Lipid extraction, preparation of fatty acid methyl esters and analysis

Cell samples that prepared as mentioned above in “Cultivation conditions for biomass were harvested by centrifugation (4500 g, 4°C) and frozen at -20°C followed by -80°C, before freeze-drying. Freeze dried biomass was accurately weighed; an internal standard (2-Terthiophene triheneicosanoin, n-21:0, Sigma) was added. Fatty-acyl methyl esters were prepared by using sulfuric-acid-catalysed trans-esterification [35,36]. After the transmethylation, fatty acid methyl esters (FAMEs) were extracted with n-hexane, concentrated under a stream of oxygen-free dry nitrogen at 37°C, to give a total lipid extract (TLE). Fractionation of phospholipids from the TLE was accomplished by thin-layer chromatography (TLC). Portions of sample TLEs were applied to silica gel plates (Silica gel 60 F254, Merck) that had been activated at 100°C for 1h. Plates were developed in CHCl3/MeOH/CH3COOH/H2O (85:15:10:3.5, v/v/v/v). Samples were visualised by iodine vapour and identified by comparison with known standards which were identified with rhodamine-6-G, ninhydrin and Dragendorff stains [37]. The lipid classes were separated by silica gel (1:30 w/w of lipid) column chromatography by successive elution with chloroform (1:10 m/v of lipid), acetone-methanol (9:1 v/v;1:15 w/v of lipids) and methanol (1:10 w/v of lipid) to get neutral-(NL), glyco-(GL) and phospho-(PL) lipids respectively. All fractions along with total lipids were transmethylated using sodium methoxide (0.5 M) to obtain the FAMEs and analyzed with a modified method published previously [38] Analyses of the FAME preparations were performed with a Hewlett-Packard model 7890A GC (Varian CP-3800, Varian, Inc. 2700 Mitchell Drive Walnut Creek, CA 94598-1675/USA) equipped with type DB225 capillary column (BPX70, 10 m x 0.1 mm, 0.2 µm; J & W Scientific, Folsom, Ca, USA) with programmed temperature of 170°C–220°C, a linear increase at 5°C min-1, injection and detection temperature maintained at 250 and 260°C, respectively, and helium as the carrier gas. GC/MS analysis was carried out with Agilent 5975 GC/MS (Agilent Technologies Co., Ltd., Palo Alto, USA) equipped with HP-5ms Capillary GC-MS Column (Agilent 19091S-433, 30 m x 0.25 mm, 0.25 µm), temperature programme 120°C for 1 min, increased at 8°C min-1 to 260°C, which was maintained for 10 min with He as the carrier gas. MS operating conditions were as follows: electron multiplier, 2,000 V; transfer line, 250°C; electron impact energy, 70 eV; scan threshold, 50; 1.3 scans s21 with a mass range of 50 to 500 atomic mass units; and solvent delay, 2.35 min. Compounds were identified by comparison of their retention times with those of known standards, and sample mass spectra data were compared to the mass spectra data of 275, 000 compounds in the Wiley 275 spectra library.


Fatty acid composition of phospholipid classes

The amount of total lipid was 12.5% of dry cell mass from 10°C MB cultures and the content of phospholipids and neutral lipids were about 72% and 28% of total lipid, respectively (Table 1). As identified by TLC from fractionation of TLEs, phosphatidyl ethanolamine (PE) was the dominant lipid class in phospholipids (50%) followed by phosphatidyl glycerol (PG) (40%). About 5% of diphosphoglyceride (DPG) and 3% of lysophosphatidylethanolamine (LPE) were also detected with some unidentified phospholipids (2%). The fatty acid compositions of TLEs and their derived PE, PG and DPG fractions are shown in Table 2. There was a near equal proportion of SCFAs within the PE and PG fractions. However, PE contained a higher proportion of BCFAs components (25% PE versus 15% PG) mainly due to a higher percentage of i-13:0 (9% PE versus 6% PG) and i-15:0 (15% PE versus 8% PG). In contrast, PG contained a greater proportion of MUFAs (28% versus 19%) which was due to slightly higher proportions of all monounsaturated acyl species. PUFA were present in both phospholipid classes, although PG contained a higher percentage of total PUFAs (17% versus 14%) and EPA (14% versus 10%). Free fatty acids (FFA) or non-esterified fatty acids (NEFA) were recovered from the TLC plates (S1 Fig), and found with high content of MUFAs and PUFAs, especially the high content of EPA and n-16:1ω7c (18.7% and 26.5%, respectively) and with less SCFAs and BCFAs. Whereas, DPG contained higher SCFAs and BCFAs, especially n-13:0 and i-15:0 (28.4% and 15.7%, respectively), and less MUFAs and PUFAs, from which 6.5% of EPA was also detected. Thus, a mass balance calculation of the percentage of EPA, based on a qualitative assessment of probable phospholipid class distribution did balance (Table 2).

Table 1. Lipid composition of strain MAR441T when grown in marine broth at 10°C.a.

Table 2. Distribution of major fatty acid in total and different lipid classes in strain MAR441T when grown in marine broth at 10°C.

The effect of growth temperature on the percentage composition of individual fatty acids in MAR441T grown between 4 and 25°C is as described in our recent publication (Supplementary S1 Table [39]) and further analysed as Fig 1B. Growth at temperatures 4°C and 10°C which are below the optimal growth temperature range of 15–20°C, resulted in a higher percentage of EPA and n-16:1ω7c, and a lower percentage of n-13:0 and n-16:0 compared to growth at 15°C or above 20°C, the optimum growth temperature range. Growth within the optimal region achieved the highest percentage of n-15:0 and n-16:0, while iso-tridecanoic acid (i-13:0) and the sum of monounsaturated fatty acids were at their lowest levels. At growth temperatures above the optimal region, the percentage of iso-pentadecanoic acid (i-15:0), n-17:0 and n-17:1ω8 were maximal, while the percentage of n-13:0 increased with increasing growth temperature. With increasing temperature from 4 ºC to 25 ºC, the proportion of monounsaturated fatty acids (MUFAs) and polyunsaturated fatty acids (PUFAs) decreased, with an increase in the proportion of straight chain fatty acids (SCFAs), whereas branched chain fatty acids (BCFAs) were at their lowest concentrations at 15°C. The values of average chain length (ACL, calculated after reference [40]), are 16.29–14.67 and quantitative levels of EPA decreased with increasing growth temperature (24–0.2 mg g-1 cells dry weight) at all growth temperatures from 4–25°C (Fig 1B).

Fig 1.

(A) Growth curves of strain MAR441T with shaking at various temperatures under atmospheric pressure in marine broth medium; (B) Change in average chain length (ACL, open diamonds) and relative proportion of whole cell FAs in strain MAR441T grown at 4, 10, 15, 20 and 25°C in marine broth medium. Straight chain fatty acids (SCFAs, filled circles), branched chain fatty acids (BCFAs, filled boxes), monounsaturated fatty acids (MUFAs, open boxes); eicosapentaenoic acid (EPA, open circles). The experiments were carried out in triplicate and values are means of three samples based on our recent published data (Supplementary S1 Table -).

Time course of cell growth and FA production

Fatty acids are intracellular products in strain MAR441T which shows a typical time course of EPA production at 15°C (Fig 2 and S1 Table). The increase in lipid and EPA content parallels that of cell growth (Fig 2A). After 36 h of cultivation, the cells entered the late exponential or early stationary phase and the content of TFAs, PUFAs and EPA reached their maximum of 103.1 mg g-1, 17.7 mg g-1and 15.48 mg g-1 respectively, when EPA is at its highest content of 15% of TFA. In this experiment a 5% culture inoculum was used, and the maximum TFA concentration reached 113 mg g-1 after 60 h culture, with ca. 16.8 mg g-1 of PUFA and 13.4 mg g-1 of EPA. The TFAs and EPA were produced with a stable quantitative yield of 90–120 mg g-1 and 11–15 mg g-1 for 60 hours from 24 h to 84 h time period culture, during which the percentage composition of SCFAs, BCFAs, MUFAs, PUFAs and EPA was kept at a relative stable levels of 41–46%, 19–22%, 18–20%, 13–17% and 10–15% respectively, and the average chain length remained more or less constant from 15.21–15.73. However, an increase in cell growth during the exponential phase led to a decrease in SCFAs from a high level of 60–43% in favour of BCFAs, MUFAs and PUFAs, due to the great decrease of n-13:0 with a corresponding increase of n-16:0, i-C15:0, n-16:1ω7 and EPA, which contributing the increase of average chain length. Interestingly, the cell death during the late stationary phase and death phase of cell culture led to a great increase in SCFAs from 50–73% at the expense of BCFAs, MUFAs and PUFAs, due to a surprising rise of n-13:0 from 29–63%, which was accompanied by a decrease of n-16:0, i-15:0, n-16:1ω7 and EPA, as well as the average chain length.

Fig 2.

(A) Time period of cell growth (filled circles) and quantitative yield of whole cell TFAs and EPA in strain MAR441T grown in marine broth medium at 15°C during the time period. Total fatty acids (TFAs, filled boxes); eicosapentaenoic acid (EPA, open circles); quantitative yield of biomass (open boxes). (B) Change in average chain length (ACL, open diamonds) and relative proportion of whole cell FAs in strain MAR441T grown in marine broth medium at 15°C during the time period. Straight chain fatty acids (SCFAs, filled circles), branched chain fatty acids (BCFAs, filled boxes), monounsaturated fatty acids (MUFAs, open boxes); eicosapentaenoic acid (EPA, open circles). The experiments were carried out in triplicate and values are means of three samples based on S1 Table.

Effect of Na+ on cell growth and PUFA production

This bacterium required Na+ for growth; growth occurred at Na+ concentrations ranging from 0.051 M (0.3%) to 1.197 M (7%) with an optimum between 0.051 and 0.513 M (3%), which is consistent with the sea-water environment containing about 19.45 g l-1 (0.85 M) Na+; the growth was inhibited at a Na+ concentration less of 0.3% or above 7%. Different concentrations of sodium chloride were supplemented to the regular no salt contained ZB medium to study their effects in biomass formation, fatty acid biosynthesis and desaturation reactions were investigated (Fig 3 and S2 Table). A significant difference in biomass formation was observed between cultures within optimum Na+ concentrations and cultures without Na+ or with high Na+ concentrations. 1.7–2.2 g l-1 biomass and 2–10 mg g-1 EPA were achieved when the Na+ in the ZB medium were controlled between 0.051–0.684 M, while only 0.4–1.0 g l-1 biomass and 0–0.18 mg g-1 EPA were produced when the medium were supplemented without or with higher concentration of Na+ (>0.85 M). A low percentage composition (0.2%) of EPA was produced by MAR441T cells, with high levels of SCFAs (64.5%), when the culture medium contained no metal ions. When the medium was supplemented with a low concentration of Na+ (0.3%), a decrease of SCFAs in favour of BCFAs, MUFAs and PUFAs was observed, the decrease of n-13:0 corresponded to the increase of n-14:0, n-16:0, i-13:0, n-16:1ω7, n-18:1ω7 and EPA, contributing to the increase of average chain length from 14.27 to 15.32. With 0.3% Na+ there was a significant increase in the productivity of EPA from 0.4% to 8.6%. EPA production reached its maximum at 15.4% of TFAs when the concentration of Na+ was maintained at 0.5%. However, increasing Na+, led to a decrease in the proportion of EPA from 15.4% to 0.4% when 7% of Na+ was added. Nevertheless, at 0.5–3% Na+, the percentage composition of SCFAs, BCFAs, MUFAs and PUFAs in MAR441T cells was stable. Whereas, growth on 4–7% Na+ increased the proportion of SCFAs, which was accompanied by a decrease in the percentages of BCFAs, MUFAs and PUFAs, and this was mirrored by a corresponding decrease of average chain length.

Fig 3.

(A) Change in average chain length (ACL, open diamonds) and relative proportion of whole cell FAs in strain MAR441T grown in ZB medium at 15°C with various concentration of NaCl. Straight chain fatty acids (SCFAs, filled circles), branched chain fatty acids (BCFAs, filled boxes), monounsaturated fatty acids (MUFAs, open boxes); eicosapentaenoic acid (EPA, open circles). (B) Quantitative yield of biomass (open boxes), TFA and EPA in strain MAR441T grown in ZB medium at 15°C with various concentration of NaCl. Total fatty acids (TFAs, filled boxes); eicosapentaenoic acid (EPA, open circles). The experiments were carried out in triplicate and values are means of three samples based on S2 Table.

Effect of sole carbon and nitrogen source on FA production

To elucidate global adaptation mechanisms of membrane fatty acids in strain MAR441T in response to growth at a low temperature on several different carbon sources, the degree of saturation (i.e. saturated/unsaturated ratio), the degree of polyunsaturation (i.e. PUFA/MUFA ratio) and average chain length (ACL) of fatty acids were determined and compared. The changes in fatty acid composition of MAR441T grown on various sole carbon/nitrogen sources at 15°C are shown in S3 Table.

Growth on L-leucine, L-alanine, L-serine, L-proline, glucose, glycerol, pyruvate, Tween 80, Tween 60, Tween 40, urea and (NH4)2SO4 produced a fatty acid composition similar to that obtained in complex media MB or ZB. Only some differences in percentage composition in quantitative levels of TFA were apparent.

Growth on complex media MB or ZB achieved high production of n-16:1ω7 (14%) and EPA (13–15%). However, growth on MB lowered production of SCFAs, such as n-13:0 (23% versus 36%), which was mirrored by a corresponding increase in BCFAs due to the higher amount of i-13:0 (7% versus 3% for ZB) and i-15:0 (10% versus 3%), as well as by a higher quantitative yield of EPA (15.5 mg g-1 versus 14.2 mg g-1) but with similar levels of ACL (15.57 versus 15.49). PUFA levels produced by strain MAR441T in MB and ZB (0.91 and 1.11 of polyunsaturation degree) were substantially higher compared with other organic substrates, where almost equal quantities of PUFA have been observed (0.1–0.86 of polyunsaturation degree).

Growth on L-alanine and L-serine produced a very similar fatty acid percentage composition as well as ACL (15.39% versus 15.38%), with a very low proportion of n-13:0 (24–26% versus 32–51% for other carbon sources excluding glycerol). This was accompanied by a rise in the percentages of other fatty acids with different levels, especially on n-16:1ω7 (11–15%) and EPA (11–13%). However, growth on L-alanine led to a higher quantitative yield of EPA (12.57 mg g-1 versus 11.13 mg g-1).

Growth on L-leucine resulted in the highest production of n-13:0 (45.7% versus 21–41% for other carbon sources) and i-15:0 (6.5%) at the expense of MUFAs and PUFAs, such as n-16:1ω7, n-18:1ω7c and EPA. This was mirrored by a corresponding decrease in the quantitative yield EPA (5.68 mg g-1) and ACL of 14.73.

Growth on L-proline decreased the proportion of BCFAs, such as i-13:0 (4% versus 5–9% for other carbon/nitogen sources excluding glycerol and Tween medium). This was accompanied by a rise in the percentages of n-18:1ω7 (7.1% versus 2–6% excluding Tween 60) and EPA (15.6% versus 4–12%). Furthermore, the increased proportion of EPA was mirrored by a corresponding increase in the quantitative yield (15.23 mg g-1 versus 4–11 mg g-1 for other carbon/nitrogen sources), which was comparable to MB cultures (15.5 mg g-1), and the ACL was in higher value (15.58).

The fatty acid composition of glucose cultured cells was very similar to that of ZB cultured cells. The only difference was that the former had a higher content of BCFAs (12.3% versus 7.7% for ZB) and lower proportion of PUFAs (12% versus 19%).

Growth on glycerol led to lower SCFA and BCFA content due to the lowest proportion of n-13:0 (21% versus 24–51% for other carbon/nitrogen sources) and i-13:0 (3.4% versus 4–9%, excluding Tween medium), whereas n-16:1ω7 at its highest point (29% versus 6–15%) which contributed a high level of MUFAs (34.7% versus 12–27%), in TFAs (saturation degree of 0.92), and percentage of EPA production was in a reasonable level (10%) as well as the quantitative yield (10.66 mg g-1).

Growth on Tween 80, 60 and 40 exhibited a marked alteration in fatty acid composition and the quantitative yield of EPA (4–7 mg g-1 versus 10–12 mg g-1 for other carbon sources) and cells (1.5–1.7 g l-1 versus 1.8–2.3 g l-1). The cultivation of strain MAR441 cells with Tween 80 (35% content of n-18:1ω9c oleic acid) caused the increase of MUFAs up to 48% in cellular lipids (saturation degree of 0.7) while inhibiting the production of PUFAs (polyunsaturation degree of 0.113). Growth on Tween 60 led to 10% of n-16:0 and 33% of n-18:0 causing the increase of SCFAs up to 77% in TFAs (saturation degree of 4.0) and therefore inhibited the BCFAs, MUFAs and PUFAs production. Growth on Tween 40 resulted in a high content of n-18:0 (15%) and n-16:1ω7c (25%), causing an increase of MUFAs up to 32% in TFAs, the degree of saturation and polyunsaturation were 1.2 and 0.29 respectively. However, the average chain length from these tween media were among the highest levels, 15.79–16.16 versus 14.73–15.62 for other carbon sources.

The percentage of EPA from the L-leucine, L-serine and L-glucose cultures was comparable to that obtained when grown in ZB at 15°C (10.2% versus 9–11%). These higher proportions of EPA corresponded with the higher quantitative yield (1–3.9 mg g-1) and higher level of TFA (4.8–9.9 mg g-1).

However, when MAR441T was grown on a single nitrogen source with urea or (NH4)2SO4, this led to a low proportion of MUFAs and PUFAs in favour of SCFAs, due to an increase of n-13:0 (42.5% for urea and 51.1% for (NH4)2SO4 with corresponding decrease of n-16:1ω7, n-18:1ω7 and EPA, and less average chain length (14.79 for urea and 14.91 for (NH4)2SO4 in that the deficiency of carbon sources used for developing backbone.

Thus, ZB medium (ZB1) supplemented with L-proline and (NH4)2SO4 for growing MAR441T cells led to a higher production of MUFAs and PUFAs with the proportion of 20.7% and 25.3% respectively due to the rise of n-16:1ω7 (20.4%) and EPA (17.6%), which was mirrored by the quantitative yield of TFA (115.2 mg g-1) and EPA (20.28 mg g-1), and higher ACL of 15.91.

Effect of cerulenin treatment on the amount of biomass, lipids and EPA in MAR441T cells

Biomass production, lipid and EPA yields obtained from MAR441T cells grown in the medium containing cerulenin up to 7.5 μg ml-1 at 4°C and 15°C are summarized in Fig 4 and S4 Table. The cell biomass was around 3.2 g dry cells l-1 when cells were grown in the medium containing cerulenin at 0.5, 1, 2.5, 5 and 7.5 μg ml-1 respectively. The value was 20% increased as that of non-treated cells. The lipid yield (g l-1) was not changed by the concentration of cerulenin in the range from 0 to 7.5 μg ml-1. However, the biomass was slightly influenced by increasing the concentration of cerulenin to the medium up to 7.5 μg ml-1, indicating some effects of cerulenin on the growth of MAR441T cells. The highest EPA yield at 81.5 μg ml-1 was obtained from cells treated with cerulenin at 0.5 μg ml-1 at 4°C. This yield was 53% increased over that obtained from non-treated cells. However, EPA production was 93% increased at 15°C when the cells were treated with 1 μg ml-1 cerulenin, whereas the percentage of PUFAs was 201% increased due to a marked increase of both n-18:2ω6t and n-18:3ω3 All these results clearly show that cerulenin treatment enhances EPA or PUFAs production as well as the short chain fatty acids, such as n-13:0 and n-15:0. mMost of the middle chain FAs, such as n-16:0, n-18:0, i-15:0, n-15:1ω6, n-16:1ω7, n-17:1ω8 and n-18:1ω7c acids were inhibited in MAR441T cells when various concentration of cerulenin were included in the medium.

Fig 4.

Change in average chain length (ACL, open diamonds) and relative proportion of whole cell FAs in strain MAR441T grown in marine broth medium at 4°C (A) and 15°C (B) with various concentration of cerulenin. Straight chain fatty acids (SCFAs, filled circles), branched chain fatty acids (BCFAs, filled boxes), monounsaturated fatty acids (MUFAs, open boxes); polyunsaturated fatty acids (PUFAs, upside down triangles); eicosapentaenoic acid (EPA, open circles). The experiments were carried out in triplicate and values are means of three samples based on S4 Table.

Fatty acid production by NTG-mutant strains (A4 and A13)

Two mutants A4 and A13 were morphologically different from the wild type strain MAR441T. Colonies of strain MAR441T on marine agar plates were 2–4 mm in diameter, tan-pigmented, butyrous in consistency, circular and convex in shape with an smooth edge, and the central rough area was adherent to or embeds into the agar and was not easy to emulsify. Whereas, colonies of NTG-mutant strains A4 and A13 on marine agar plates were 3–5 mm in diameter, tan-pigmented, opaque, dull, with dentate margin or undulate edge, and the central rough area was attached to the agar loosely, and this was easy to be pushed/moved away by pipette tips (S2A Fig). Under scanning electron microscopy (SEM) cells of A4 and A13 were found to lack fimbriae after the NTG mutation (S2B Fig). The effect of growth temperature on the percentage composition of individual fatty acids in MAR441T and its NTG-mutants (A4 and A13) grown at 4°C, 15°C and 25°C is shown in Fig 5 and S5 Table. Comparing to the fatty acid compositions of strain MAR441T, the mutants were found with lower levels of SCFAs and higher percentage of BCFAs at 15°C and 25°C, and lower levels of EPA at 4°C. However, the percentage of EPA in mutant A13 was higher than that in wild type strain at 15°C, and the levels of EPA were decreased at 4°C and 25°C. By increasing temperatures from 4°C to 25°C, the values of ACL in these mutants decreased. The quantitative level of lipid content of the mutants were similar to that of MAR441T at 4°C and 15°C, while increasing levels of EPA were found at 25°C. Mutant A13 could reach relatively higher level of EPA of 15.8 mg g-1 cells dry weight at 15°C (S5 Table).

Fig 5. Change in average chain length (ACL, open diamonds) and relative proportion of whole cell FAs in strain MAR441T and its NTG mutants (A4 and A13) grown at 4, 15 and 25°C.

Straight chain fatty acids (SCFAs, filled circles), branched chain fatty acids (BCFAs, filled boxes), monounsaturated fatty acids (MUFAs, open boxes), polyunsaturated fatty acids (PUFAs, upside down triangles) and eicosapentaenoic acid (EPA, open circles). The experiments were carried out in triplicate and values are means of three samples based on S5 Table.


Environmental adaptation

The fatty acid composition of MAR441T exhibited changes in response to growth temperature and sole carbon/nitrogen source, as has been reported for some Shewanella PUFAs producers [15,28], and non-PUFA producers, such as Cobetia marina [41]. Both the percentage and the quantitative level of EPA markedly changed at different growth temperatures, indicating that PUFA may play a critical role in the modulation of membrane fluidity [28,42]. The precise reason why these bacteria produce omega-3 fatty acids is still unclear, although many PUFA synthase genes responsible for EPA/DHA synthesis have been cloned and sequenced [43,44], and been successfully expressed in E. coli [5,11]. As growth temperature increased, MAR441T also demonstrated a novel adaptive response by increasing the percentage of n-13:0 and i-15:0 with corresponding decrease of n-16:1ω7 and n-18:1ω7, which might be due to the role of fatty acid precursor selection in this bacterium as an adaptive response [45]. This finding is also supported by some other strains possessing similar fatty acid compositions, which share similar adaptive responses, such as Shewanella gelidimarina ACAM 456T, which exhibited an increased proportion of n-15:0 and a decrease of n-16:1ω7 and EPA with increasing culture temperature [28], S. olleyana ACEM 9T showed an increase in the percentage of n-16:0 and i-15:0 with a corresponding decrease of n-16:1ω7, n-18:1ω7 and EPA when the growth temperature increased [20]. These observations indicate that PUFAs may play a key role in the homeostatic adaptation of cellular membrane viscosity and the modulation of membrane fluidity in these marine isolates.

Growth on the sole carbon sources L-proline, Tween 80, Tween 60 or Tween 40 demonstrated that in this bacterium the fatty acid composition can be manipulated by the provision of potential acyl chain primers [28]. There was a large increase in the observed percentage of EPA and lower levels of i-13:0 and i-15:0 in the L-proline cultures. While an observed increase of i-13:0 and to a lesser extent i-15:0 from L-alanine and L-leucine cultures that there is a preference for 4–5 cycles of chain elongation from the alanine- or leucine-derived primer molecule [46]. This is similar to S. gelidimarina ACAM 456T, which exhibited a similar change in fatty acid composition when it was grown in L-leucine medium [28]. The degree of acyl chain elongation may therefore be primer-specific. L-alanine and L-proline also increased the level of TFA, and therefore EPA, suggesting an increase in the level of total lipid resulted from growth on these substrates. In contrast, FA composition patterns were dominated by the n-18:1ω9 in Tween 80 cultures, while mainly n-18:0 in Tween 60 cultures and with rise of n-16:1ω7 when grown in Tween 40, which greatly decrease the percentage of PUFAs or polyunsaturation degree. These data are in agreement with the results obtained for EPA-producing Shewanella strain GA-22, the Tween 80-grown cells showed an increase of monounsaturated fatty acids, up to 78% in cellular lipids and an inhibition of the PUFA production [15]. These observations were a consequence of the nature of the added substrates, because Tween 80 contains oleic (n-18:1) acid, Tween 60 has stearic acids (n-18:0) and Tween 40 is composed of n-16:0. Although Tween 80, 60 and 40 usually serve as surfactants, they can also be used as carbon sources.

However, the strain preferred growing in the complex media, such as marine broth and ZoBell broth, from which the higher levels of biomass, lipid or TFA and EPA were produced. This corroborates previous reports that production of PUFA at low temperatures was enhanced more than two-fold and reached 5% of total fatty acids in the strain GA-22 cells grown on marine broth [15]. In our study, L-proline and (NH4)2SO4 were selected as the most suitable carbon and nitrogen sources used in combination for preparing ZB medium, which improved the production of biomass two-fold and EPA 1.5-fold.

MAR441T required Na+ for growth and EPA synthesis and this is the case with other marine species such as S. halifaxensis, S. sediminis, S. pealeana and S. woodyi, which also preferred low temperatures for growth and are thus considered cold-adapted obligate species of Shewanella [47,48]. Studies on protein coding sequences from two obligate marine S. halifaxensis and S. sediminis, found that many genes coding Na+-dependent nutrient transporters were recruited to use the high Na+ content as an energy source. For example, many unique Na+-dependent nutrient symporters and Na+/nutrient symporters are used for transport of L-glutamine acid, L-proline, dicarboxylate and amino acids [49]. Based on genome annotations of Shewanella species, L-glutamine acid was predicted to be an essential precursor for biosynthesis of heme, nucleobase (purine, pyrimidine), peptidoglycan, aminosugar and fatty acids[50]. The requirement of Na+ as a motive force for transport of these essential growth substrates is consistent with the nature of strain MAR441T being an obligate marine bacterium.

Cerulenin specifically blocks the activity of β-keto acyl thioester synthetase, which may account for the inhibition of fatty acid synthesis [51]. However, EPA production was greatly improved in MAR441T treated by cerulenin, and middle-chain fatty acids were almost absent. These results suggest that cerulenin inhibited the de novo synthesis of middle-chain fatty acids, but not the synthesis of EPA and short-chain fatty acids. Therefore, strain MAR441T may employ two fatty acid-biosynthetic systems and independently synthesize middle-chain fatty acids and LC-PUFAs using a common starting material as a FA precursor [24].

NTG mutagenesis is particularly useful for chemical mutagenesis of a variety of Gram-negative bacteria [52]. However, NTG mutagenesis has not previously been reported as a means for increasing PUFA production from environmental isolates of bacteria, although S. putrefaciens strain 2738 has been treated by NTG to get cold-sensitive EPA-requiring mutants at low temperatures [11]. Therefore, by treating MAR441T with NTG at higher temperatures, we were able to obtain less temperature sensitive mutants with improved levels of lipid and EPA content when grown at higher temperatures.

EPA yield

The yield of EPA from strain MAR441T ranged from 2 to 20 mg g-1 (30 mg g-1 from the cerulenin-treated cultures) (cells dry weight) or 6 to 63 mg l-1 depending on culture conditions, and showed the highest quantitative yield of 11% TFA of dry cell weight and 20 mg l-1day-1 of EPA production in marine broth at 15°C for 1.5 days, values which compare favourably with literature reports for other EPA-producing bacteria. A Shewanella putrefaciens-like strain, SCRC-8132 produced 4–15 mg g-1 (cells dry weight) of EPA and 2% TFA of dry cell weight, although it was reported with high percentage of EPA (24–40%) [53,54]. Under various temperatures and utilizing different carbon sources, Shewanella gelidimarina ACAM 456T produced 1–16 mg g-1 (cells dry weight) of EPA [28], whereas the S. putrefaciens-like strain SCRC-2738, is presently identified as S. pneumatophori SCRC-2738 [55], which produced 4–11 mg g-1 of EPA [56], the production was further enhanced to 17 mg g-1 [57].

Many wild-type strains of autophotrophic microalgae produce similar levels of EPA to the bacteria mentioned above, such as P. tricornutum and Monodus subterraneus [58]. Continuous culture of the heterotrophic diatom Nitzschia laevis, gave EPA productivities of 73 mg l-1 day-1 using a glucose feed [59]. In a high cell density system maximum cell dry weight and EPA yields were 22.1 g l-1 and 695 mg l-1, respectively, in a 14-day incubation [60]. While, the productivity of P. tricornutum by using batch culture was 25 mg l-1 day-1 [58], and improved up to 40 mg l-1 day-1 using a photobioreactor [61], N. alba ATCC 40775 gave an enhanced EPA production of 100–300 mg l-1 day-1 (40–50 g l-1 biomass) [62], Thraustochytrid strains were reported with an EPA productivity of 47 mg L-1 day-1 in a 50 l tubular photobioreactor [63], Mortierella alpina 1S-4 and M. subterraneus were reported with EPA production of 30 and 50 mg l-1 day-1 respectively [63], M. alpine was with high biomass production of 43 g l-1 and EPA productivity of 60 mg l-1 day-1 [64], and Nannochloropsis oceanica CY2 gave an EPA content and biomass concentration of 2.4% (per dry cell weight) and 1.5 g/l and productivity of 13.2 mg l-1 day-1 [65]. However, the majority of these algal species generally require strictly controlled growth conditions in terms of nutrients, light quantity and quality, oxygenation and carbon dioxide levels; these factors can result in considerable expense. In contrast, strain MAR441T has a high EPA productivity of 20 mg l-1day-1 (2 g l-1 of biomass) when grown in marine broth at 15°C and could provide a cost effective and reliable source of PUFA. However, biomass yield per unit volume and culture time may also affect production economics. Nevertheless, the strain could be used as a feedstock for organisms, such as rotifers, as a way of introducing them into a marine food web for producing PUFA-rich oils by aquaculture.

EPA in phospholipid and non-esterified fatty acids

The occurrence of non-esterified fatty acids (NEFA), or free fatty acids (FFA) in MAR441T was identified using TLC and GC, accounting for 15% of the total lipid and containing 18.7% EPA. Phospholipids were found as the main component of total lipid (72% versus 28% for neutral lipid). The major phospholipid classes found were phosphatidyl ethanolamine (PE) and phosphatidyl glycerol (PG) as the main components (50% versus 40%), appreciable levels of EPA were detected from both fractions (9.5% versus 14.2% for PG) and 6.5% EPA in diphosphoglyceride (DPG), which is accounted for 5% of total lipid. The presence of non-esterified fatty acids within a PUFA-producing Vibrio sp. was previously reported to be 13.3% of total lipid, and EPA accounted for 13% of NEFA [66]. PE (with 5.5% of EPA) and PG (with 10.6% of EPA) accounted for 61% and 19% of phospholipids respectively in Aeromonas sp. 3010, and a high content of EPA and palmitoleic acid (19.7 and 50%, respectively) was found in the free fatty acid fraction [67]. For Shewanella sp. strain ACAM 456, PG contained a higher percentage of total PUFA (14% versus 9% for PG) and EPA (13% versus 9%), and NEFA accounted for 9% of total lipid and contained 22% EPA [28]. By employing FAB-MS-MS, acyl chains, such as i-13:0/13:0 and i-14:0/14:0 appeared to be associated with EPA in PE phospholipid species only, whereas the association of n-17:1 and acyl-18:0 chains with EPA was specific to PG in strain ACAM 456 [28]. Yazawa also reported the presence of NEFA in the PUFA producing strain SCRC-2738, with 5–10% of total EPA in the non-esterified form. This may be explained by the contribution of most EPA-producing bacterial genera possessing PG as their major phospholipid type, MUFAs and EPA were mostly concentrated in the PG component, while the proportion of branched-chain fatty acids was elevated in PE. Furthermore, the above mentioned five PUFA-producing bacteria all contain appreciable amounts of NEFA or FFA, of which EPA appears as a particular component of NEFA in PUFA metabolism. This remains an area for further investigation.

Genetically modified organisms (GMOs) as a source of ingredients have been used in our foods for over 20 years—the major argument for using them is that they offer "increased yield, drought tolerance, enhanced nutrition" and other consumer benefits [68]. However, the traditional approach of GMO risk assessment has been questioned [10], there is also an argument against GMOs due to the cause of health problems, environmental damage and violation of farmers and consumers rights [69]. Furthermore, consumers become more health conscious and knowledgeable about what they eat and where it comes from and if the food they eat contains GMOs, and they do try to avoid this [70]. Thus, GMOs as a source of dietary supplements is not recommended as a way to support better nourishment. Nevertheless, the number of companies which are publicly taking a stand against genetically modified foods is growing. For example, smaller natural foods companies are removing foods sourced from GMOs from their products. For example Popcorn Indiana's Fit Popcorn, Trader Joes' health and beauty products and Clif Bars [71]. Our approach to producing improved yields of omega-3 fatty acids using wild-type microorganisms, without using recombinant DNA techniques is an approach which is often preferred by an increasing number of ingredients manufacturers.


This study reveals that temperature, medium composition, Na+ concentration, carbon and nitrogen sources, and the addition of cerulenin all play crucial roles in affecting bacterial growth and EPA accumulation in Shewanella electrodiphila MAR441T. The most efficient EPA production occurred when the strain was grown in modified Marine medium with L-proline and (NH4)2SO4 at a concentration of 1.50 g/l. The optimal EPA content (20–30 mg/g) and EPA productivity (20 mg/l/d) is among the highest levels yet reported, indicating the potential of using this system for the commercial production of EPA using this new wild-type marine bacterium.

Supporting information

S1 Fig. Lipid profiles from strain MAR441T whole-cell lipid extracts separated on TLC plates.


S2 Fig.

(A) Colonies of strain MAR441T and its NTG mutants (A4 and A13) grown on marine agar plates at 15 ºC for 3 days; (B) scanning electron microscopy (Right, Bar 500 nm) of a negatively-stained cell of strain MAR441T and its NTG mutants (A4 and A13).


S1 Table. Fatty acid composition of strain MAR441T during a time course of cell growth in marine broth medium at 15°C.


S2 Table. Fatty acid composition of strain MAR441T grown on various concentrations of NaCl in ZB liquid medium at 15°C.


S3 Table. Fatty acid composition of strain MAR441T grown on various sole carbon/nitrogen sources at 15°C.


S4 Table. Fatty acid composition of strain MAR441T grown on various concentrations of cerulenin in marine broth medium at 4°C and 15°C.


S5 Table. Fatty acid composition of strain MAR441T and its NTG mutants (A4 and A13) grown on marine broth at 15°C.



The authors acknowledge the officers and crew of RRS James Cook and Dr W. D. K. Reid and Dr B. Wigham for the provision of data and sediment samples made available for this study.


  1. 1. Schantz SL, Widholm JJ, Rice DC (2003) Effects of PCB exposure on neuropsychological function in children. Environ Health Perspect 111: 357–576. pmid:12611666
  2. 2. Dailey FE, McGraw JE, Jensen BJ, Bishop SS, Lokken JP, Dorff KJ, et al. (2015) The Microbiota of Freshwater Fish and Freshwater Niches Contain Omega-3 Fatty Acid-Producing Shewanella Species. Appl Environ Microbiol 82: 218–231. pmid:26497452
  3. 3. Amiri-Jami M, Abdelhamid AG, Hazaa M, Kakuda Y, Griffths MW (2015) Recombinant production of omega-3 fatty acids by probiotic Escherichia coli Nissle 1917. FEMS Microbiol Lett 362.
  4. 4. Gong Y, Wan X, Jiang M, Hu C, Hu H, et al. (2014) Metabolic engineering of microorganisms to produce omega-3 very long-chain polyunsaturated fatty acids. Prog Lipid Res 56: 19–35. pmid:25107699
  5. 5. Amiri-Jami M, Griffiths MW (2010) Recombinant production of omega-3 fatty acids in Escherichia coli using a gene cluster isolated from Shewanella baltica MAC1. J Appl Microbiol 109: 1897–1905. pmid:20666868
  6. 6. Balogun KA (2016) Lipid metabolism and the risk factors of cardiovascular disease: implication of dietary omega-3 polyunsaturated fatty acids. Appl Physiol Nutr Metab: 1.
  7. 7. Kramkowska M, Grzelak T, Czyzewska K (2013) Benefits and risks associated with genetically modified food products. Annals of Agricultural and Environmental Medicine 20: 413–419. pmid:24069841
  8. 8. Schantz SL, Gardiner JC, Aguiar A, Tang X, Gasior DM, Sweeney AM, et al. (2010) Contaminant profiles in Southeast Asian immigrants consuming fish from polluted waters in northeastern Wisconsin. Environ Res 110: 33–39. pmid:19811781
  9. 9. Arts M, Ackman R, Holub B (2001) “Essential fatty acids” in aquatic ecosystems: a crucial link between diet and human health and evolution. Can J Fish Aquat Sci 58: 122–137.
  10. 10. Rajan SR, Letourneau DK (2012) What Risk Assessments of Genetically Modified Organisms Can Learn from Institutional Analyses of Public Health Risks. Journal of Biomedicine and Biotechnology.
  11. 11. Valentine RC, Valentine DL (2004) Omega-3 fatty acids in cellular membranes: a unified concept. Prog Lipid Res 43: 383–402. pmid:15458813
  12. 12. Nichols DS, McMeekin TA (2002) Biomarker techniques to screen for bacteria that produce polyunsaturated fatty acids. J Microbiol Methods 48: 161–170. pmid:11777566
  13. 13. Bowman JP, McCammon SA, Nichols DS, Skerratt JH, Rea SM, Nichols PD, et al. (1997) Shewanella gelidimarina sp. nov. and Shewanella frigidimarina sp. nov., novel Antarctic species with the ability to produce eicosapentaenoic acid (20:5 omega 3) and grow anaerobically by dissimilatory Fe(III) reduction. Int J Syst Bacteriol 47: 1040–1047. pmid:9336903
  14. 14. Delong EF, Yayanos AA (1986) Biochemical function and ecological significance of novel bacterial lipids in deep-sea procaryotes. Appl Environ Microbiol 51: 730–737. pmid:16347037
  15. 15. Gentile G, Bonasera V, Amico C, Giuliano L, Yakimov MM (2003) Shewanella sp. GA-22, a psychrophilic hydrocarbonoclastic antarctic bacterium producing polyunsaturated fatty acids. J Appl Microbiol 95: 1124–1133. pmid:14633042
  16. 16. Nichols D, Bowman J, Sanderson K, Nichols CM, Lewis T, McMeekin T, et al. (1999) Developments with antarctic microorganisms: culture collections, bioactivity screening, taxonomy, PUFA production and cold-adapted enzymes. Curr Opin Biotechnol 10: 240–246. pmid:10361072
  17. 17. Nogi Y, Masui N, Kato C (1998) Photobacterium profundum sp. nov., a new, moderately barophilic bacterial species isolated from a deep-sea sediment. Extremophiles 2: 1–7. pmid:9676237
  18. 18. Wang F, Xiao X, Ou HY, Gai Y, Wang F (2009) Role and regulation of fatty acid biosynthesis in the response of Shewanella piezotolerans WP3 to different temperatures and pressures. J Bacteriol 191: 2574–2584. pmid:19201790
  19. 19. Kato C, Nogi Y (2001) Correlation between phylogenetic structure and function: examples from deep-sea Shewanella. FEMS Microbiol Ecol 35: 223–230. pmid:11311432
  20. 20. Skerratt JH, Bowman JP, Nichols PD (2002) Shewanella olleyana sp. nov., a marine species isolated from a temperate estuary which produces high levels of polyunsaturated fatty acids. Int J Syst Evol Microbiol 52: 2101–2106. pmid:12508875
  21. 21. Frolova GM, Pavel KG, Shparteeva AA, Nedashkovskaia OI, Gorshkova NM, Ivanova EP, et al. (2005) Lipid composition of novel Shewanella species isolated from far Eastern seas. Mikrobiologiia 74: 766–771. pmid:16400986
  22. 22. Ivanova EP, Sawabe T, Gorshkova NM, Svetashev VI, Mikhailov VV, Nicolau DV, et al. (2001) Shewanella japonica sp. nov. Int J Syst Evol Microbiol 51: 1027–1033. pmid:11411670
  23. 23. Ivanova EP, Sawabe T, Hayashi K, Gorshkova NM, Zhukova NV, Nedashkovskaya OI, et al. (2003) Shewanella fidelis sp. nov., isolated from sediments and sea water. Int J Syst Evol Microbiol 53: 577–582. pmid:12710629
  24. 24. Amiri-Jami M, Wang H, Kakuda Y, Griffiths MW (2006) Enhancement of polyunsaturated fatty acid production by Tn5 transposon in Shewanella baltica. Biotechnol Lett 28: 1187–1192. pmid:16799757
  25. 25. Russell NJ, Nichols DS (1999) Polyunsaturated fatty acids in marine bacteria—a dogma rewritten. Microbiology 145 (Pt 4): 767–779.
  26. 26. Allen EE, Facciotti D, Bartlett DH (1999) Monounsaturated but not polyunsaturated fatty acids are required for growth of the deep-sea bacterium Photobacterium profundum SS9 at high pressure and low temperature. Appl Environ Microbiol 65: 1710–1720. pmid:10103272
  27. 27. Sato S, Kurihara T, Kawamoto J, Hosokawa M, Sato SB, Esaki N. (2008) Cold adaptation of eicosapentaenoic acid-less mutant of Shewanella livingstonensis Ac10 involving uptake and remodeling of synthetic phospholipids containing various polyunsaturated fatty acids. Extremophiles 12: 753–761. pmid:18668196
  28. 28. Nichols DS, Nichols PD, Russell NJ, Davies NW, McMeekin TA (1997) Polyunsaturated fatty acids in the psychrophilic bacterium Shewanella gelidimarina ACAM 456T: molecular species analysis of major phospholipids and biosynthesis of eicosapentaenoic acid. Biochim Biophys Acta 1347: 164–176. pmid:9295160
  29. 29. Nichols D, McMeekin T, Nichols P (1994) Manipulation of polyunsaturated, branched-chain and trans-fatty acid production in Shewanella putrefaciens strain ACAM 342. Microbiology 140: 577–584
  30. 30. Peng YF, Chen WC, Xiao K, Xu L, Wang L, Wan X. (2016) DHA Production in Escherichia coli by Expressing Reconstituted Key Genes of Polyketide Synthase Pathway from Marine Bacteria. PLoS One 11: e0162861. pmid:27649078
  31. 31. Zhang J, Burgess JG (2015) Shewanella electrodiphila sp. nov., a psychrotolerant bacterium isolated from Mid-Atlantic Ridge deep-sea sediments. Int J Syst Evol Microbiol 65: 2882–2889. pmid:25999594
  32. 32. ZoBell C (1946) Marine Microbiology. Waltham, Massachusetts, USA: Chronica Botanica Company. pp. 240.
  33. 33. Zhang JW, Zeng RY (2007) Psychrotrophic amylolytic bacteria from deep sea sediment of Prydz Bay, Antarctic: diversity and characterization of amylases W J Microbiol Biotechnol 23: 1551–1557.
  34. 34. Dobson SJ, James SR, Franzmann PD, McMeekin TA (1991) A numerical taxonomic study of some pigmented bacteria isolated from Organic Lake, an Antarctic hypersaline lake. Archives of Microbiology 156: 56–61.
  35. 35. Komagata K, Suzuki K (1987) Lipid and cell-wall analysis in bacterial systematics. In Methods in Microbiology. London: Academic Press. pp. 161–208.
  36. 36. Christie WW (1989) Gas Chromatography and Lipids. A Practical Guide. Ayr, UK: The Oily Press. pp. 64–84.
  37. 37. Kates M (1986) Techniques of lipidology: Isolation, analysis and identification of lipids. In: Burdon R, van Kippenberg P, editors. Laboratory techniques in biochemistry and molecular biology. New York (NY) Elsevier. pp. 1–464.
  38. 38. Opperman M, Marais D, Benade AJS (2011) Analysis of omega-3 fatty acid content of South African fish oil supplements. Cardiovascular Journal of Africa 22: 324–329. pmid:22159321
  39. 39. Zhang J, Burgess JG (2015) Shewanella electrodiphila sp. nov., a psychrotolerant bacterium isolated from Mid-Atlantic Ridge deep-sea sediments. Int J Syst Evol Microbiol.
  40. 40. White SW, Zheng J, Zhang YM, Rock (2005) The structural biology of type II fatty acid biosynthesis. Annu Rev Biochem 74: 791–831. pmid:15952903
  41. 41. Yumoto I, Hirota K, Iwata H, Akutsu M, Kusumoto K, Morita N, et al. (2004) Temperature and nutrient availability control growth rate and fatty acid composition of facultatively psychrophilic Cobetia marina strain L-2. Arch Microbiol 181: 345–351. pmid:15067498
  42. 42. Nichols D, Russell N (1996) Fatty acid adaptation in an Antarctic bacterium changes in primer utilization. Microbiology 142: 747–754.
  43. 43. Allen EE, Bartlett DH (2002) Structure and regulation of the omega-3 polyunsaturated fatty acid synthase genes from the deep-sea bacterium Photobacterium profundum strain SS9. Microbiology 148: 1903–1913. pmid:12055309
  44. 44. Okuyama H, Orikasa Y, Nishida T, Watanabe K, Morita N (2007) Bacterial genes responsible for the biosynthesis of eicosapentaenoic and docosahexaenoic acids and their heterologous expression. Appl Environ Microbiol 73: 665–670. pmid:17122401
  45. 45. Jostensen JP, Landfald B (1996) Influence of growth conditions on fatty acid composition of a polyunsaturated-fatty-acid-producing Vibrio species. Arch Microbiol 165: 306–310. pmid:8661921
  46. 46. Kaneda T (1991) Iso- and anteiso-fatty acids in bacteria: biosynthesis, function, and taxonomic significance. Microbiol Rev 55: 288–302. pmid:1886522
  47. 47. Zhao JS, Manno D, Beaulieu C, Paquet L, Hawari J (2005) Shewanella sediminis sp. nov., a novel Na+-requiring and hexahydro-1,3,5-trinitro-1,3,5-triazine-degrading bacterium from marine sediment. Int J Syst Evol Microbiol 55: 1511–1520. pmid:16014474
  48. 48. Zhao JS, Manno D, Leggiadro C, O'Neil D, Hawari J (2006) Shewanella halifaxensis sp. nov., a novel obligately respiratory and denitrifying psychrophile. Int J Syst Evol Microbiol 56: 205–212. pmid:16403888
  49. 49. Zhao JS, Deng Y, Manno D, Hawari J (2010) Shewanella spp. genomic evolution for a cold marine lifestyle and in-situ explosive biodegradation. PLoS One 5: e9109. pmid:20174598
  50. 50. Makemson JC, Hastings JW (1979) Glutamate functions in osmoregulation in a marine bacterium. Appl Environ Microbiol 38: 178–180. pmid:16345412
  51. 51. Vance D, Goldberg I, Mitsuhashi O, Bloch K (1972) Inhibition of fatty acid synthetases by the antibiotic cerulenin. Biochem Biophys Res Commun 48: 649–656. pmid:4625866
  52. 52. Lewenza S, Falsafi RK, Winsor G, Gooderham WJ, McPhee JB, Brinkman FS, et al. (2005) Construction of a mini-Tn5-luxCDABE mutant library in Pseudomonas aeruginosa PAO1: a tool for identifying differentially regulated genes. Genome Res 15: 583–589. pmid:15805499
  53. 53. Yazawa K (1996) Production of eicosapentaenoic acid from marine bacteria. Lipids 31 Suppl: S297–300.
  54. 54. Yazawa K, Araki K, Okazaki N, Watanabe K, Ishikawa C, Inoue A, et al. (1988) Production of eicosapentaenoic acid by marine bacteria. J Biochem 103: 5–7. pmid:2834356
  55. 55. Hirota K, Nodasaka Y, Orikasa Y, Okuyama H, Yumoto I (2005) Shewanella pneumatophori sp. nov., an eicosapentaenoic acid-producing marine bacterium isolated from the intestines of Pacific mackerel (Pneumatophorus japonicus). Int J Syst Evol Microbiol 55: 2355–2359. pmid:16280496
  56. 56. Yazawa K, Araki K, Watanabe K, Ishikawa C, Inoue A, Kondo K, et al. (1988) Eicosapentaenoic acid productivity of the bacteria isolated from fish intestines. Nipp Suis Gakk 54: 1835–1838.
  57. 57. Akimoto M, Ishii T, Yamagaki K, Ohtaguchi K, Koide K, Yazawa K. (1990) Production of eicosapentaenoic acid by a bacterium isolated from Mackerel intestines. J Am Oil Chem Soc 67: 911–915.
  58. 58. Yongmanitchai W, Ward OP (1991) Growth of and omega-3 fatty acid production by Phaeodactylum tricornutum under different culture conditions. Appl Environ Microbiol 57: 419–425. pmid:2014989
  59. 59. Wen ZY, Chen F (2003) Heterotrophic production of eicosapentaenoic acid by microalgae. Biotechnol Adv 21: 273–294. pmid:14499126
  60. 60. Wen Z-Y, Jiang Y, Chen F (2002) High cell density culture of the diatom Nitzschia laevis for eicosapentaenoic acid production: fed-batch development. Proc Biochem 37: 1447–1453.
  61. 61. Miron A, Gomez A, Camacho F, Grima E, Chisti Y (1999) Comparative evaluation of compact bioreactors for large-scale monoculture of microalgae. J Biotechnol 70: 249–270.
  62. 62. Boswell K, Gladue R, Prima B, Kyle D (1992) SCO production by fermentative microalgae. In: Kyle D, Ratledge C, editors. Industrial applications of single cell oils. Champaign, Illinois: American Oil Chemists`Society. pp. 274–286.
  63. 63. Grima E, Perez J, Camacho F, Fernandez F, Sevilla J, Sanz F. (1994) Effect of dilution rate on eicosapentaenoic acid productivity of Phaeodactylum tricornutum UTEX-640 in outdoor chemostat culture. Biotech Lett 16: 1035–1040.
  64. 64. Bajpai P, Ward O (1991) Effects of ageing Mortierella mycelium on production of arachidinic acid and eicosapentaenoic acid. J Am Oil Chemists Soc 68: 775–780.
  65. 65. Chen CY, Chen YC, Huang HC, Ho SH, Chang JS (2015) Enhancing the production of eicosapentaenoic acid (EPA) from Nannochloropsis oceanica CY2 using innovative photobioreactors with optimal light source arrangements. Bioresource Technology 191: 407–413. pmid:25777066
  66. 66. Henderson RJ, Millar RM, Sargent JR, Jostensen JP (1993) Trans-monoenoic and polyunsaturated fatty acids in phospholipids of a Vibrio species of bacterium in relation to growth conditions. Lipids 28: 389–396. pmid:8316045
  67. 67. Cho KW, Mo SJ (1999) Screening and characterization of eicosapentaenoic acid-producing marine bacteria. Biotechnol Lett 21: 215–218.
  68. 68. Uzogara SG (2000) The impact of genetic modification of human foods in the 21st century: A review. Biotechnology Advances 18: 179–206. pmid:14538107
  69. 69. Meyer H (2011) Systemic risks of genetically modified crops: the need for new approaches to risk assessment. Environmental Sciences Europe 23: 7.
  70. 70. Wunderlich S, Gatto KA (2015) Consumer Perception of Genetically Modified Organisms and Sources of Information. Advances in Nutrition 6: 842–851. pmid:26567205
  71. 71. Kulikowski L (2014) 5 companies against genetically modified foods. 5 companies against genetically modified foods, https://wwwthestreetcom/story/12530112/1/5-companies-against-genetically-modified-foodshtml. New York: