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Creating omega-3 fatty acids-enriched chicken using de-fatted green microalgal biomass Stephanie Gatrell, JongGun Kim, Theodore Derksen, Eleanore O'Neil, and Xingen Lei J. Agric. Food Chem., Just Accepted Manuscript • Publication Date (Web): 22 Sep 2015 Downloaded from http://pubs.acs.org on October 16, 2015

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Journal of Agricultural and Food Chemistry

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Creating omega-3 fatty acids-enriched chicken using de-fatted green microalgal biomass

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Stephanie K. Gatrell, JongGun Kim, Theodore J. Derksen, Eleanore V. O’Neil, and Xin Gen.

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Lei*

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Department of Animal Science, Cornell University, Ithaca NY 14853

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⃰ Corresponding author; Tel: 607-254-4703; Fax: 607-255-9829; Email: [email protected]

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Running Title: Omega-3 enrichment with defatted microalgae

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Abstract

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This study was to create an omega-3 (n-3) fatty acids-enriched chicken product by using defatted

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green microalgae (DGA, Nannochloropsis oceanica) biomass out of biofuel research. Hatching

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Ross broiler chicks were fed a corn-soybean meal diet containing 0 (control), 2, 4, 8 or 16%

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DGA for 6 wk (n = 6 cages/diet). The DGA inclusion resulted in a linear (P < 0.001) increase in

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total n-3 fatty acids, eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) in plasma,

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liver, breast, and thigh at wk 3 and 6. The increase in the breast EPA + DHA by the 16% DGA

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diet reached 60-fold (P < 0.0001) over the control. The 8 and 4% DGA diets elevated (P < 0.05)

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liver mRNA levels of ∆-9 (88%) and ∆-6 (96) desaturases. In conclusion, 8-16% of the DGA

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can be added in diets for broilers to produce an n-3 fatty acids-enriched chicken meat.

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KEY WORDS: broiler chicken, defatted microalgal biomass, docosahexaenoic acid,

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eicosapentaenoic acid, gene expression

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Introduction Consuming diets high in long chain polyunsaturated fatty acids (PUFAs), in particular

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omega-3 (n-3) fatty acids, has been linked to a decreased risk of cardiovascular disease, diabetes,

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arthritis, and cancer1,2,3,4,5. However, current Western diets contain an unbalanced ratio of the

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“pro-inflammatory” omega-6 (n-6) and “anti-inflammatory” n-3 PUFAs. These diets have an

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average n-6:n-3 fatty acid ratio of 20-30:1, as opposed to the traditional range of 1-2:16.

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Increasing public awareness of the health benefits of n-3 fatty acids has led researchers

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attempting to improve the fatty acid profile of commonly consumed animal products. Because

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the average American consumes approximately 40 kg of chicken annually7, this type of meat is a

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promising candidate for n-3 fatty acids enrichment.

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Fatty acid profiles of chicken breast, thigh, and skin are well-known to be highly

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dependent on dietary fatty acid composition8. Previously, marine sources, mainly fish oil and

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fish meal9,10,11 were used to enhance n-3 fatty acid incorporation into poultry meat. However,

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recent increases in cost of and demand for fishmeal have led to the investigation of alternative n-

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3 fatty acids rich sources for a more sustainable industry. Indeed, dietary inclusion of plant-

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based α-linolenic acid (ALA)-rich flaxseed12,13,14 or canola oil15 could elevate the n-3 fatty acid

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content of broiler meat. However, that enhancement resulted from an increased deposition of

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ALA, not of the longer chain docosahexaenoic acid (DHA) or eicosapentaenoic acid (EPA).

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Metabolically, ALA needs to be converted to DHA and(or) EPA to be functionally meaningful in

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the human body. But, the efficiency of that conversion in vivo seems to be limited16.

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Microalgae contain a superior fatty acid profile with high abundance of EPA and(or)

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DHA to traditional animal feed protein supplements17,18,19. They also contain relatively high

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amounts of crude protein20, essential amino acids21, and carotenoids22. Supplementation of

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microalgae to poultry diets has been shown to improve the overall n-3 fatty acid status in egg

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yolk17,23 and breast muscle16. Our laboratory has been specifically interested in defatted

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microalgal biomass, the byproduct of biofuel research, as a partial replacement of conventional

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poultry feed ingredients. In broiler chicks, we have demonstrated that moderate levels of

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defatted microalgal biomass inclusion exhibited no adverse effects on growth performance or

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health24. However, whether the defatted microalgae, with the residual fatty acids left in the

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biomass, could enhance the deposition of n-3 fatty acids in the tissues of chicks were not

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explored.

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Consuming diets high in PUFAs ultimately enriches cell membranes in these fatty acids,

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subsequently altering signaling molecules involved in carbohydrate and lipid metabolism25.

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Enzymes involved in de novo fatty acid synthesis, such as malic enzyme (ME) and fatty acid

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synthase (FASN) are known to be affected by dietary manipulation and feeding status26,27,28,29.

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Furthermore, desaturase enzymes that introduce double bonds into fatty acids, including ∆-6 and

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∆-9 desaturases, are also affected by nutritional status30,31. Thus, it is fascinating to determine

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how the defatted microalgal biomass affects the expression of these genes and how that effect is

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related to the enrichment of n-3 fatty acids in the chick tissues.

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Therefore, the objectives of this experiment were: 1) to determine the feasibility in

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producing an n-3 fatty acids-enriched chicken by feeding broiler chicks increasing levels of a

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defatted green microalgal biomass (DGA, Nannochloropsis oceanica) (Cellana, Kailua-Kona,

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HI), and 2) to reveal the biomass-mediated lipid metabolism gene expression mechanism related

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to the enrichment of n-3 fatty acids in the tissues of chicks.

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Materials and Methods

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Animals, diets, and management: All protocols of this experiment were approved by

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the Institutional Animal Care and Use Committee of Cornell University (IACUC approval

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number 2010-0106). Male hatchling Ross broiler chicks were obtained from a commercial

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hatchery and housed in a temperature-controlled unit at the Cornell University Poultry Research

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Farm. The broiler chicks were housed in thermostatically-controlled cage batteries for the first 3

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wk (6 chicks per cage); and 4 chicks per cage (2 killed for sample collection at wk 3) were then

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transferred to grower cages at room temperature from wk 3 to 6. Chicks had free access to feed

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and water and received a lighting schedule of 22 h light and 2 h dark. Birds were fed one of five

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diets (n = 6 cages/diet) containing 0% (control), 2%, 4%, 8% or 16% DGA, on an ‘as is’ basis,

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replacing a mixture of corn and soybean meal. Nutrient composition of DGA is shown in Table

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S1. Starter (wk 0 to 3, Table S2) and grower (wk 3-6, Table S3) diets were formulated to be

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isoenergetic and to meet the requirements for all essential nutrients for each phase of growth32.

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The fatty acid profiles of each starter and grower diet are given in Table 1. At wk 3 and 6, 2

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birds per cage were euthanized via asphyxiation with CO2 and blood was drawn from heart

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puncture by using heparinized needles. After blood was kept on ice and centrifuged at 3000 g

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for 15 min, plasma was stored at -20oC until analysis. Liver, breast muscle, and legs were

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removed and a portion of each was snap frozen in liquid nitrogen and stored at -80oC until

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analysis. Whole skinless breast and legs were sealed in plastic bags and frozen for lipid analysis.

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Fatty acid extraction: For fatty acid extraction, experimental diets (~100 grams) were

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ground to a fine powder. Tissue samples were taken from the liver, the core of the whole breast

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(pectoralis major) and thigh (bicep femoris). Total lipids from the diet (100 mg), plasma (100

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µL), and tissue samples (1 g) were extracted, with some modifications, according to Folsh et al33,

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and were methylated with methanolic-KOH according to Ichihara et al34, by using tridecanoic

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acid (Sigma-Aldrich Co., St Louis, MO) as an internal standard. Each fatty acid was identified

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by comparing its retention time and peak area with the individual fatty acid methyl ester

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standards (Sigma-Aldrich Co., St Louis, MO). Methyl esters of fatty acids were analyzed by

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using a gas chromatography instrument (Agilent 6890N, Agilent Technologies, Santa Clara, CA)

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fitted with a flame-ionization detector and used a fused-silica capillary column coated with CP-

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SIL 88 (100 m × 0.25 mm i.d., 0.2 mm film thickness; Varian Inc, Lake Forest, CA). Oven

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temperature was programmed to be held for 4 min at 140ºC, increased by 4ºC per min to 220ºC,

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and then held for 5 min. Carrier gas was N2 with constant flow rate of 2 ml/sec and injector

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temperature was 230ºC and detector temperature was 280ºC. Fatty acid content is expressed as

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either a weight percentage of the sum of all of the fatty acids analyzed, or calculated as mg of

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fatty acid per 100g of wet tissue.

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Gene expression: Real time RT-PCR was performed on the snap frozen liver samples to

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estimate the abundance of mRNA by using β-actin as a reference gene. Target genes included

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ME, FASN, ∆-6 desaturase, and ∆-9 desaturase. After the RNA was isolated and its quality was

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verified by agarose gel and spectrometry (A260/A280), it was transcribed by using a

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commercially available kit (Applied Biosystems, Grand Island, NY). The resulting cDNA (0.3

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µg) was added to a 10 µL total reaction that included Power SYBR Green PCR mater mix

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(Applied Biosystems) and 0.625 μM forward and reverse primers (Table 2). Real-time PCR

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analysis was performed by using a 7900HT Fast Real-Time PCR System (Applied Biosystems).

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The PCR conditions consisted of an initial 2 minute 50oC step and a “hot start” step at 95oC for

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10 min, followed by 40 cycles of a 95oC denaturing step for 15 sec and a 60oC annealing step

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for 60 sec. A melting curve was analyzed for all primers to ensure the quality of the

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amplification product. Each sample was analyzed in duplicate for both the target and reference

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genes. Relative mRNA abundance was determined by using the ∆ cycle threshold (∆Ct) method.

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For each sample, the Ct difference between the target and reference gene was calculated (∆Ct =

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Cttarget – Ctreference). The ∆Ct values were then converted to fold differences by raising 2 to the

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power – ∆Ct (2-∆Ct).

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Statistical analyses: Data were pooled within cage for an experimental unit of 6 per

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treatment, and were analyzed by ANOVA and(or) linear regression models by using PC-SAS

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9.2. Mean differences between dietary groups were separated by using Duncan’s multiple range

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test. For the gene expression data, selected treatment effects were directly compared with the

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control group by using the t-test. Data are expressed as mean ± SEM, and the treatment effects

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were deemed significant at P < 0.05, and a trend at P < 0.10.

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Results and Discussion The animal performance and biochemical analyses, including plasma uric acid, inorganic

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phosphorus and protein, for this experiment were reported elswhere35. Moderate levels (8%) of

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DGA were tolerated by chicks for 6 wk without affecting growth performance or plasma

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biochemistry. However, the higher level of inclusion (16%) led to a reduction in indices of

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growth performance.

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The fatty acid composition of the defatted microalgal biomass and experimental diet is

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shown in Table 1. Containing 3.6% total lipid, the DGA was rich in EPA (representing 16.5%

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of total fatty acids by weight), but without DHA. The fatty acid profile (as a weight percentage

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of total fatty acids) of wk 6 plasma is shown in Table 3 (wk 3 is shown on Table S4). The main

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fatty acid found in all dietary treatments at both time points was linoleic acid (C18:2 n6),

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followed by palmitic acid (C16:0) and stearic acid (C18:0), which are similar. There was no

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effect of DGA inclusion on saturated fatty acids (SFA), monounsaturated fatty acids (MUFA), or

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PUFAs, regardless of time. In contrast, the DGA inclusion resulted in a linear increase in plasma

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n-3 fatty acids, and the increases were 5- and 15-fold by the 16% DGA diet over the control at

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wk 3 (P < 0.0001, R2 = 0.93) and 6 (P < 0.0001, R2 = 0.75), respectively. The increases were

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manifested with elevated EPA (C20:5 n3) and DHA (C22:6 n3), indicating that the birds were

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able to convert the dietary EPA into DHA. At wk 6, there was a linear reduction in n-6 fatty

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acids (P < 0.05), resulting in a favorable decrease in the ratio of n-6 to n-3 fatty acids (P