3. Sources of Long-chain Omega-3

## **Readily Available Sources of Long-Chain Omega-3 Oils: Is Farmed Australian Seafood a Better Source of the Good Oil than Wild-Caught Seafood?**

**Peter D. Nichols, Brett Glencross, James R. Petrie and Surinder P. Singh** 

**Abstract:** Seafood consumption enhances intake of omega-3 long-chain (≥C20) polyunsaturated fatty acids (termed LC omega-3 oils). Humans biosynthesize only small amounts of LC-omega-3, so they are considered semi-essential nutrients in our diet. Concern has been raised that farmed fish now contain lower LC omega-3 content than wild-harvested seafood due to the use of oil blending in diets fed to farmed fish. However, we observed that two major Australian farmed finfish species, Atlantic salmon (*Salmo salar*) and barramundi (*Lates calcifer*), have higher oil and LC omega-3 content than the same or other species from the wild, and remain an excellent means to achieve substantial intake of LC omega-3 oils. Notwithstanding, LC omega-3 oil content has decreased in these two farmed species, due largely to replacing dietary fish oil with poultry oil. For Atlantic salmon, LC omega-3 content decreased ~30%–50% between 2002 and 2013, and the omega-3/omega-6 ratio also decreased (>5:1 to <1:1). Australian consumers increasingly seek their LC omega-3 from supplements, therefore a range of supplement products were compared. The development and future application of oilseeds containing LC omega-3 oils and their incorporation in aquafeeds would allow these health-benefitting oils to be maximized in farmed Australian seafood. Such advances can assist with preventative health care, fisheries management, aquaculture nutrition, an innovative feed/food industry and ultimately towards improved consumer health.

Reprinted from *Nutrients*. Cite as: Nichols, P.D.; Glencross, B.; Petrie, J.R.; Singh, S.P. Readily Available Sources of Long-Chain Omega-3 Oils: Is Farmed Australian Seafood a Better Source of the Good Oil than Wild-Caught Seafood? *Nutrients* **2014**, *6*, 1063–1079.

#### **1. Introduction**

The health benefits of omega-3 long-chain (≥C20) polyunsaturated fatty acids (LC-PUFA, also termed LC omega-3 oils) were first documented over three decades ago. Scientists observed that Greenland Eskimos had lower incidence of heart disease than other ethnic groups despite their high fat diet that was rich in the blubber of marine mammals [1]. The main LC omega-3 oils that have been attributed to this health benefit are eicosapentaenoic acid (EPA, 20:5ω3) and docosahexaenoic acid (DHA, 22:6ω3). For brevity, we use the term LC omega-3 here in consideration of both fatty acids, and also that of docosapentaenoic acid (DPA, 22:5ω3). Seafood has traditionally been the major source of these health-benefitting LC omega-3 oils [2]. Over the past decade the supply of farmed seafood has steadily increased, with the contribution of aquaculture to human food supplies now similar in volume to that of the wild catch harvest.

Aquaculture is currently the main user of industrially produced fish oils, with around 90% of global fish oil production used in aquafeeds [3]. Fish oils are used in feeds for most aquaculture species to satisfy both essential fatty acid and energetic demands [4]. However, as aquaculture has expanded, the ability of existing supplies, including the increasing cost, of the wild harvest fish oil resource to meet industry needs has been largely surpassed [5]. Non-marine sources of oil, including vegetable and animal-derived, are therefore now also included in aquafeeds as alternatives to fish oil. However, both alternatives to fish oil have a lower content of LC omega-3 oil, which has the flow on effect of causing a lower concentration of LC omega-3 in farmed seafood products compared to that observed previously [6]. Generally Australian consumers and also, to our knowledge, consumers in most other countries are not fully aware of the lower LC omega-3 content now occurring in many farmed fish species. In addition to the trend of fish oil replacement largely by oil blends, other potential strategies exist, including finishing diets used prior to harvest and novel ingredients such as LC omega-3 precursors [7]. However, to ensure the long-term supply of LC omega-3 oils for aquaculture there is a clear need for new and sustainable sources of these oils from fishery independent sources.

Obtaining information on the composition of farmed seafood products is important for future developments within the aquaculture and associated feed manufacturing industries. This study was initiated to examine the fatty acid content and composition of two major Australian farmed fish species— Atlantic salmon (*Salmo salar*) and barramundi (*Lates calcifer*). Particular emphasis was given to the LC omega-3 oils, and also the comparison of the changes seen in the fatty acid profiles between samples collected over 2010–2013 and earlier data for these same species obtained during 2002 when a largely fish oil diet was in use. As many Australian consumers increasingly seek their LC omega-3 from supplements, a range of supplement products also were examined and compared.

#### **2. Experimental Section**

#### *2.1. Sample Collection*

Norwegian quality cut (NQC) samples from three fresh fillets of farmed Atlantic salmon were obtained twice per year between December 2010 and November 2013. At CSIRO, the skin was removed and each sample was independently blended for two minutes in an OSKAR 400 continuous flow homogenizer. Whole barramundi were supplied to CSIRO in November 2010 by Marine Produce Australia Pty Ltd. (Broome, WA, Australia). The right fillet of each of three fish was used and the NQC analogous cut from that fillet of the barramundi was taken. The NQC sample was then minced, with a sample analysed for moisture content by gravimetric analysis after oven drying at 105 °C for 24 h. The remainder of the sample was frozen, freeze dried, ground in a coffee grinder, and then shipped to Hobart for lipid extraction and analysis.

Fourteen commercially produced omega-3 fish oil capsules were purchased from a local Hobart pharmacy. Each product was given a laboratory code: FO1-14. Oil capsules were cut open with a scalpel and the oil transferred to a glass vial using a glass pipette.

#### *2.2. Lipid Extraction*

NQC samples (typically between 1 and 3 g wet weight for Atlantic salmon, dry weight for barramundi) were quantitatively extracted overnight using a modified Bligh and Dyer [8] single-phase methanol-chloroform-water extraction (2:1:0.8 v/v/v). The phases were separated by addition of chloroform-water (final solvent ratio, 1:1:0.9 v/v/v methanol-chloroform-water). The total solvent extract (TSE) was concentrated using rotary evaporation at 40 °C, and total lipid content was determined gravimetrically.

#### *2.3. Fatty Acid Analysis*

An aliquot of the TSE of the farmed fish samples or the fish oils dissolved in chloroform was trans-methylated to produce FA methyl esters (FAME) using methanol–chloroform–conc. hydrochloric acid (3 mL, 10:1:1, 80 °C, 2 h) [9]. FAME were extracted into hexane–chloroform (4:1, 1.8 mL). The samples were dried on a heat block (40 °C) under a stream of nitrogen gas, and an internal injection standard (C19 or C23 FAME) added.

Samples were analysed by gas chromatography (GC) using an Agilent Technologies 7890A GC (Palo Alto, CA, USA) fitted with a Supelco Equity™-1 fused silica capillary column (15 m × 0.1 mm ID, 0.1 μm film thickness (Bellefont, PA, USA) an FID, a split/splitless injector and an Agilent Technologies 7683B Series auto sampler and injector. Helium was the carrier gas. Samples were injected in splitless mode at an oven temperature of 120 °C. After injection, the oven temperature was raised to 250 °C at 10 °C min−1 and finally to 270 °C at 3 °C min−1. Peaks were quantified with Agilent Technologies ChemStation software (Palo Alto, CA, USA). GC-mass spectrometric (GC-MS) analyses of selected samples were performed on a Finnigan Thermoquest GCQ GC-MS fitted with an on-column injector using Thermoquest Xcalibur software (Austin, TX, USA). The GC was fitted with a capillary column of similar polarity to that described above. Individual components were identified using mass spectral data and by comparing retention time data with those obtained for authentic and laboratory standards.

#### **3. Results**

#### *3.1. Oil Content*

Lipid content of the farmed barramundi samples averaged 10% (WW) in 2002 and 8.5% in 2010 (wet weight, WW). Oil content of the farmed Atlantic salmon samples ranged from 6.7% to 14.7% (WW). The 2010 and 2011 autumn Atlantic salmon samples contained lower oil content than all other samples collected from 2010 to 2013.

#### *3.2. Fatty Acid Composition and Content*

#### 3.2.1. Farmed Fish

Major fatty acids (FA) (as % of the total fatty acids, TFA) in farmed Australian Atlantic salmon harvested in 2002 were in decreasing order of abundance: 16:0 (18%), DHA (17%), 18:1ω9 (oleic acid, OLA; 15%), EPA (10%) and 16:1ω7 (6%) (Table 1). The three LC omega-3 PUFA—EPA + DPA + DHA accounted for 30% of the TFA.

For farmed Atlantic salmon obtained in 2010–2013, the FA profile differed both from the 2002 sample, and also over the 4 year period. Major FA in the 2010–2013 samples in decreasing order of abundance were: OLA, 16:0, 18:2ω6 (linoleic acid, LOA) and 16:1ω7 (Table 1); these four FA accounted for 57% (autumn 2010) rising to 72% of TFA in late spring 2013. In 2002, these four FA had accounted for 42% of TFA in farmed salmon. The next most abundant FA in the 2010–2013 Atlantic salmon samples were: EPA, DHA and 18:0, with all three FA decreasing over this period. LOA increased from around 2.5-fold (autumn 2010) to 4-fold (spring 2013) compared to the 2002 sampling, and the relative proportion of arachidonic acid (ARA) decreased over the 2010–2013 period.

Expressed on an absolute basis (mg/100 g serve, WW), LC omega-3 content in farmed Atlantic salmon was 2010 mg/100 g in 2002, then ranged from as high as 1770 mg/100 g (spring 2010) decreasing to 980 mg/100 g in spring 2013 (Figure 1). The ratio of omega-3 PUFA/omega-6 PUFA was 7.8 in 2002 samples, and through 2010 to 2013 showed a steady decrease from 2.6 (autumn 2010) to 0.8 (spring 2013) (Figure 2).

The FA profiles of farmed and wild caught barramundi are shown in Table 2. Major FA (as % TFA) in farmed samples from 2010 in decreasing order of abundance were: OLA, 16:0, LOA and 16:1ω7; these four FA accounted for approximately 60% of TFA in farmed barramundi. The next most abundant FA in farmed barramundi in 2010 were: EPA, DHA and 18:0. The relative level values for DHA in particular are lower than recorded for the 2002 farmed samples.


**Table 1.** Composition of fatty acids (as percent of total FA) in farmed Atlantic salmon (*n* = 3)—2002 and 2010–2013.

SFA, Saturated fatty acids; MUFA, monounsaturated fatty acids; PUFA, polyunsaturated fatty acids. Prefix a denotes anteiso branching.

**Figure 1.** Content (±SD) of LC omega-3 oils (EPA + DPA + DHA, mg/100 g) in farmed Tasmanian Atlantic salmon sampled in 2002 [10] and 2010–2013.

**Figure 2.** Ratio (±SD) of omega-3 PUFA/omega-6 PUFA in farmed Tasmanian Atlantic salmon, 2002 [10] and 2010–2013.

Major fatty acids in wild caught barramundi were 16:0, OLA, 18:0, DHA and ARA (Table 2). Less abundant components included LOA, 16:1ω7c, EPA, 18:1ω7c and DPA. Freshwater samples had higher relative levels of LOA, OLA, 16:1ω7c, 18:1ω7c, 18:3ω3 (α-linolenic acid, ALA), 14:0 and lower EPA and DHA than the saltwater specimens. The greatest difference between the wild fresh and saltwater samples for any single FA was observed for DHA (5%, freshwater; 22%, saltwater). Saltwater fish contained higher levels of LC omega-3 oils than freshwater fish. In addition to containing considerably higher relative levels of DHA, the saltwater fish contained higher relative levels of total PUFA than freshwater fish (Table 2). The relative level of ω6 PUFA was similar in freshwater barramundi and saltwater fish. In freshwater fish ARA made up 7.1% of TFA, while in the saltwater wild fish it made up 12.2% of TFA. In contrast, in farmed barramundi collected in 2002 and

2010, ARA levels were consistent at 0.6% to 0.7% and an order of magnitude lower. The ratio of ω3 PUFA/ω6 PUFA differs between salt (1.9) and freshwater (0.8) barramundi (Table 2).


**Table 2.** Composition of fatty acids (as percent of total FA) in wild and farmed barramundi.

2002 farmed barramundi [10]. 1998 and 2002 samples analysed using identical methods to 2010 fish. Wild caught barramundi data [11]. Abbreviations as used in Table 1. Prefix a denotes anteiso branching.

Absolute concentration data (mg/100 g wet weight) for the LC omega-3 oils in wild harvest barramundi are shown in Figure 3. Wild saltwater fish contained higher DHA levels. For farmed barramundi, LC omega-3 content (EPA + DPA + DHA) was markedly higher than in the wild harvest samples, in 2002, although the content decreased to 790 mg/100 g in 2010 (Figure 3).

#### 3.2.2. Fish Oil Capsules

The capsule samples were divided into three groups of LC omega-3 containing products. The first group was those products containing elevated EPA + DHA (FO1-5, Table 3); these products contained ≥500 mg of EPA + DHA per capsule, which represents use of enrichment processes to obtain the product. One product reached this grouping largely based on its larger capsule size (Cenovis Fish Oil Plus, FO4, 1500 mg) compared with all other products. The second group of capsules (FO6-9) was the brands containing 180 mg EPA and 120 mg DHA per 1000 mg capsule. The Cenovis Fish Oil Plus (FO4) 1500 mg capsule product, when normalized to 1000 mg, contained the same DHA content as the other group 2 oils purchased, with EPA higher than for the other group 2 products. The third group (FO10-14) contained lower EPA + DHA content than group 1 and 2 oils, ranging from 52 to 160 mg per capsule.

The FA profiles of the fish oil capsule products are shown in Table 4. All fish oil capsule supplements examined generally contained EPA + DHA at levels indicated on the product labels. The five group 1 fish oils contained 41%–78% EPA+DHA. The four group 2 oils—containing 180 mg EPA + 120 mg DHA—were very similar in composition containing around 30% EPA + DHA. The group 3 oils contained 16%–25% EPA + DHA.

Other major FA present in group 2 and 3 oils included the saturated FA (SFA)—16:0, 18:0 and 20:0, the monounsaturated FA (MUFA)—OLA, 18:1ω7c and 16:1ω7c. Several group 3 oils—wild salmon oil and cod-liver oil—both contained elevated levels (~18%) of LC-MUFA—20:1ω11c (salmon oil), 20:1ω9 (cod liver oil) and 22:1ω11c (Table 4)—distinguishing these two oils readily from other group 2 and 3 oils. The group 1 PUFA-enriched oils varied in composition. The Omega Brain oil (FO2) was elevated in DHA, with the other four group 1 products (FO1, FO3, FO4, FO5) each containing EPA > DHA. Products FO3 (Healthy Care Fish Oil One a Day) and FO5 (Natures Own Omega-3 Ultra) showed similar profiles, with 32%–36% EPA and 23% DHA. The omega-3 joint product (FO1) contained the highest EPA+DHA levels (78%), with a number of other PUFA present (18%); this oil contained the highest EPA/DHA ratio (3.4), and represents a highly purified PUFA-containing oil product compared with all other products.


\* EPA + DHA data not supplied on label. Fatty acid data from [11].

**Table 3.** Fish oil capsules—brand and composition details as supplied by manufacturers.


**Table 4.** Fatty acid composition (as % of total fatty acids) of fish oil capsule products.

Sample codes—refer to Table 3. Abbreviations as used in Table 1. Other FA (≤0.3% of total FA): 14:1ω5c, 4,8,12TMTD, i15:0, a15:0, i16:0, MBrFA:1, MBrFA, 17:1, C18PUFA, i18:0, 18:1ω7t, 18:1, 19:1, 20:1ω5c, 22:1ω7, 24:1ω11c.

#### **4. Discussion**

### *4.1. Farmed Fish—Oil Content and Fatty Acid Profiles*

Total oil (lipid) content of farmed Atlantic salmon from 2010 to 2013 was generally similar to the oil content previously observed for Tasmanian Atlantic salmon [10]. Total lipid content in the whole body and fillet of most fish species, and barramundi are included in this regard, tends to increase with fish size [12]. In comparison, wild caught barramundi are considerably lower in lipid content (0.4% to 0.9%, WW; shoulder portion) [11]. Farmed barramundi fed a fish oil-based diet and previously analysed contained various concentrations of lipid according to study, diet and fish size and fillet section analyzed; around 10% (WW) lipid [10], 1.3% to 30% lipid [13] and 8% to 14% lipid [14].

The relative level (as % TFA) values for EPA and in particular DHA for the two farmed fish species sampled in 2010–2013 are lower than those recorded in the 2002 analyses when both species were being fed a fish oil based-diet (barramundi—EPA 6.2%, DHA 10.2%; Tasmanian Atlantic salmon— EPA 10%, DHA 17%) [10]. These 2010–2013 % composition values for EPA and DHA are also generally lower than those observed for most white muscle wild-caught fish species [11].

In addition to the reduction in EPA and DHA, the relative level of omega-6 PUFA was elevated in both farmed species sampled over the 2010–2013 period relative to 2002, due mainly to increased LOA. This leads to a dramatic shift in the ratio omega-3/omega-6 (*i.e.*, from >7:1 in 2002, to ≤1:1 currently for Atlantic salmon). A recent study [15] followed the release of an American Heart Association advisory on omega-6 fatty acids and cardiovascular risk [16] and has indicated—"advice to specifically increase omega-6 intake is unlikely to provide the intended benefits, and may actually increase the risk of CHD and death". A further change in the FA profile of the two farmed fish is a large increase in the relative level of OLA. Collectively, this profile shift is reflecting a change in diet formulation to include higher proportions of terrestrial animal and plant-based oils, with concomitant reduction in fish oil. Similar changes in salmon oil quality occur in overseas markets [17]. Also notable over the 2002–2013 period was the overall decrease in SFA which are regarded as the "least healthy" and are known precursors for endogenous cholesterol synthesis. Therefore at least one element of the current fish oil replacement strategy could be argued as improving the nutritional quality of farmed fish. It is the absolute content of the LC omega-3 that is the most critical issue in terms of the contribution that farmed fish make to the human diet. On an absolute (mg/100 g serve) basis, the LC omega-3 content values observed for farmed Atlantic salmon and barramundi sampled in 2010–2013 are considerably higher than those found in most wild caught fish species [10,11], although the values from both species are lower than previously reported a decade ago in 2002 for these two farmed species fed a fish oil-based diet [10]. Large differences were observed between the autumn and spring/summer samples in 2010 and 2011. Tasmanian waters varied in water temperature through this period, together with the presence and degree of disease and other biological factors; these may be contributing factors to the observed differences.

Wild harvest barramundi contained considerably lower absolute amounts of PUFA than the famed samples (Figure 3). However, a low oil content combined with similar high relative levels of omega-3 in saltwater fish and both ω3 and ω6 PUFA in freshwater fish are nutritionally attractive features of the wild-harvest barramundi. The overall FA profile and lower LC omega-3 content of the wild specimens

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provide no major nutritional advantage in terms of these key components. Both wild specimens had an omega-3/omega-6 ratio similar to or poorer that that found in farmed barramundi.

Until about 10 years ago Australian and New Zealand fish farms were largely using feeds made with high inclusion of fishery products (fish meals and fish oils). Prior to this period, there was evidence that LC omega-3 content in farmed fish products remained high [2,3,10]. Increasing demand for fishery resources, such as fish oil, at a time where there is limited ability or capacity to increase sustainable harvest of wild fish stocks, has resulted in increased use of oils derived from non-traditional sources [7]. Oils now being routinely used include those from land plants (e.g., canola and palm oil) and rendered oils (e.g., poultry oil) [6]. The problem with the use of these oils is that they result in flesh with: (i) lower relative levels of LC omega-3 (as % of TFA); (ii) lower absolute content of the LC omega-3; and (iii) lower omega-3/omega-6 ratio. As the relative levels of these key LC omega-3 oils decrease, LOA and OLA increase. OLA and LOA are derived from the non-marine ingredients that are being increasingly substituted into aquafeeds, although most fish also have significant capacity to synthesis their own OLA from both lipid and non-lipid substrates [4].

Interest has existed in Australia and New Zealand on comparison of the LC omega-3 oil content of salmon farmed in the two locations. In Tasmania, the species farmed is Atlantic salmon (*Salmo salar*), whilst in New Zealand the predominant species is Chinook salmon (*Oncorrhynchus tshawytscha*, also termed king salmon). LC omega-3 content of the two species sampled at a similar time (spring 2012) showed Atlantic salmon containing 1117 ± 117 mg/100 g LC omega-3 with Chinook salmon at 2568 ± 153 mg/100 g. The two species generally receive similar diets [18], and the differences in LC omega-3 content (and similarly for the SFA and omega-6 PUFA) result largely from the higher oil content of the fillet of farmed Chinook salmon (~25% oil content cf 10%–15% in Atlantic salmon).

Atlantic salmon and barramundi have, when fed a FO-containing diet, provided an excellent source of beneficial omega-3 LC-PUFA for human consumption, but reduced concentrations of these nutrients, as occurs through the use of vegetable oil and/or animal fat diets, may reduce their nutritional benefit to consumers. Limited research has been performed to examine this issue. In one study [19], dietary intake of differently fed salmon (100% fish oil (FO), 50/50 FO/rapeseed oil, 100% rapeseed oil) and the influence on markers of human atherosclerosis were compared. Significant differences between the human consumer groups were observed in the serum fatty acid profiles, especially for the levels of total omega-3 PUFA and the omega-3/omega-6 ratio, which were markedly increased in the FO-fed fish consuming group in contrast to the two other groups. In addition, significant reductions of serum TAG and of vascular cell adhesion molecule-1 and interleukin-6 were observed in patients receiving the FO-fed salmon diet when compared with the two other groups. The authors concluded that Atlantic salmon fed the FO-containing diet containing very high concentrations of omega-3 LC-PUFA seemed to produce favourable biochemical changes in patients with coronary heart disease risk factors when compared with ingestion of fillets with intermediate and low levels of the marine omega-3 LC-PUFA, where FO was replaced in part or in full by rapeseed oil [19]. To our knowledge, there have been no consumer trials with fish fed diets containing LOA, ALA and/or SDA rich oils *versus* FO derived EPA + DHA, and looking at the effects on consumers.

A recent study tested whether Atlantic salmon smolt fed a diet with a higher DHA/EPA ratio and a lower content of LC omega-3 oils to that of conventional FO based diets would enhance deposition of LC omega-3 in the liver and muscle [20]. Comparisons were made between fish fed: (1) a FO diet; (2) a blend of 50% rapeseed and 50% tuna oil diet (termed model oil, MO1); (3) a blend of 50% rapeseed, 25% tuna and 25% FO diet (MO2); and (4) a blend of 50% FO and 50% poultry oil diet (FO/PO). The latter diet was representative of commercial diets in use in Australia at the time of the study, with the proportion of chicken fat increasing even further since the study was performed. The dietary DHA/EPA ratio was in the order MO1 > MO2 > FO/PO ~ FO. The LC omega-3 content was approximately 2-fold lower in the MO1, MO2 and FO/PO diets compared to the FO diet, with the relative levels (as % total FA) lowest in the MO1 diet. For the feeding trial, there were comparable contents of LC omega-3 in the muscle of the FO, MO 1 and FO/PO fed fish [20].

A major outcome for the feeding trial was the observation that a higher DHA/EPA ratio than that commonly occurring with FO-only diets used for Atlantic salmon was better suited for more efficient deposition of LC omega-3 in the flesh, in particular DHA. Evidence was therefore apparent for LC omega-3 "sparing" in the Atlantic salmon smolt fed a diet with a high DHA/EPA ratio [20]. The use of a 50% FO and 50% PO blend in aquafeeds for Atlantic salmon, as was in the range commercially practiced in Australia in 2010, resulted in comparable LC omega-3 content in the muscle [20] and liver of juvenile Atlantic salmon to a FO fed fish. It is noteworthy that such an oil blend decreases the inefficient utilization of a 100% FO diet, due to the high loss of EPA in particular, and may be considered as an appropriate current strategy, in terms of LC omega-3 sparing, for present use in aquafeeds for Atlantic salmon [20]. The reduction of FO incorporation in the aquafeeds has also enhanced the sustainability of the industry, although sufficient FO still remains in the feeds used to ensure that farmed Tasmanian Atlantic salmon remains one of the best sources of the LC omega-3 oils available to Australian consumers.

It is important to note that in spite of changes that have occurred in feeding practices, and the resulting lower content of LC omega-3 oils, the scope remains for the potential future use of new alternate sources of LC omega-3 to restore the content of these health-benefitting ingredients to those higher contents previously seen.

Further research is needed to determine the optimum relative and absolute concentrations of dietary EPA and DHA to enhance their deposition in larger-sized commercially farmed Atlantic salmon. The rationale to pursue such studies is supported by recent developments in plant genomics. As this research field has progressed, important breakthrough steps have included: the isolation and characterization of genes from the marine microalgae encoding front-end desaturases involved in DHA biosynthesis, the isolation of highly efficient desaturases and elongases, the use of genes with omega-3 substrate preference and the development and use of a land plant (tobacco) leaf-based assay using interchangeable design principles to rapidly assemble multistep recombinant pathways [21–23]. Progress with research on insertion of microalgal-derived genes leading to DHA production into a range of omega-3 C18 PUFA accumulating land plants has been reviewed [24–26].

Recent developments have resulted in oilseeds containing elevated LC omega-3 oils, including with fish oil-like proportions of DHA and an elevated omega-3/omega-6 ratio (2–5) [22,23,27]. These major breakthroughs can in the future provide the feed and aquaculture industries with an opportunity to both sustainably farm this key protein source for the growing global population, and enhance the nutritional quality of farmed seafood products in terms of the health-benefitting LC omega-3 oils. Alternate feed-grade oils containing the desired FA profiles, in particular including a high DHA/EPA ratio, are not presently available. However, a realistic examination of the future steps required suggests that they can be a commercial reality by the end of this decade [22,23].

#### *4.2. Fish Oil Capsules as Sources of EPA + DHA*

All products generally contained EPA + DHA at levels indicated on the product labels. The cost per 500 mg EPA + DHA (value generally advised for daily consumption by many nutritional and health groups [2]) varied markedly from \$0.05 to \$5.50 (Table 3), or from \$20 to \$2000 per annum (Figure 4). The lowest cost products, based on EPA + DHA levels alone, were those oils containing the conventional 180 mg EPA and 120 mg DHA per 1000 mg capsule (group 2). In the group 2 products, a four-fold cost difference occurred (\$20–\$79 pa, Table 3), although product specifications were identical. Consumers can have difficulties in compliance (e.g., consumption of multiple large capsules), and may prefer the range of enriched products (group 1) now available. The most expensive product based on EPA + DHA content was the krill oil, which was six-fold more expensive than the next most costly product.

**Figure 4.** Cost per annum of fish oil products to supply 500 mg/day EPA + DHA. Sample codes refer to brands shown in Table 3. Enriched products—denotes group 1 brands containing concentrated EPA and/or DHA. 18/12 oils denotes—group 2 standard fish oils containing 180 mg/120 mg of EPA and DHA respectively. Other oils denotes—group 3 products containing varying proportions of EPA and DHA and may contain other components.

#### **5. Conclusions**

In summary, in comparison to 2002 samples of two major farmed Australian finfish species, Atlantic salmon and barramundi sampled in 2010–2013 have been shown to contain decreased relative levels and content of LC omega-3 oils (from 2014 mg/100 g in 2002 decreasing to 975 mg/100 g in spring 2013 for Atlantic salmon; 1970 mg/100 g in 2002 decreasing to 790 mg/100 g per serve for barramundi). These changes have resulted from the use of new, lower cost and sustainable ingredients

in farmed fish feed, necessitated by the inability of existing supplies together with the increasing cost of the wild harvest fish oil resource to meet the expanding needs of the aquaculture industry. Notwithstanding, these two widely available farmed fish species still remain an excellent source of the LC omega-3 oils, and in a broader context remain one of the best of all foods available for Australian consumers. All fish oil capsule supplements examined generally contained EPA + DHA at levels indicated on the product labels and, similar to the farmed seafood, such products can also represent a viable alternative source of LC omega-3 oils for consumers.

Subject to consumer demand, one of the cost-effective strategies to increase the omega-3 content of the lipid profile of farmed fish could be to include enhanced levels of omega-3 rich vegetable oils, and/or finishing diets with higher inclusions of marine oils [6]; such a strategy may result in grades of farmed seafood product being available for purchase that contain higher content of LC omega-3 than standard products grown using a predominately non-marine oil based diet. In the longer term, other strategies to consider include: selection for enhanced delta-5 and delta-6 desaturase activities (the key enzymes converting shorter chain omega-3 to EPA and DHA) and the use of alternate long-chain omega-3 oil sources, including new oil seeds containing EPA and DHA.

#### **Acknowledgments**

The authors thank the CSIRO Food Futures Flagship Omega-3 research team for their contributions. 1998 and 2002 seafood data was produced by the CSIRO Marine Oils groups, in particular Ben Mooney, Nick Elliott and Patti Virtue, with funding support from the Fisheries Research and Development Corporation. We also thank Marine Produce Australia Pty Ltd for supplying the 2010 farmed barramundi samples, Matt Miller of NZ Plant and Food for providing NZ King salmon data, and Nick Elliott, Peter Kube and Carol Mancuso Nichols of CSIRO and two anonymous journal reviewers for valuable comments on the manuscript.

#### **Conflicts of Interest**

The authors declare no conflict of interest.

#### **References**


## **Characterization of Oilseed Lipids from "DHA-Producing**  *Camelina sativa***": A New Transformed Land Plant Containing Long-Chain Omega-3 Oils**

### **Maged P. Mansour, Pushkar Shrestha, Srinivas Belide, James R. Petrie, Peter D. Nichols and Surinder P. Singh**

**Abstract:** New and sustainable sources of long-chain (LC, ≥C20) omega-3 oils containing DHA (docosahexaenoic acid, 22:6ω3) are required to meet increasing demands. The lipid content of the oilseed of a novel transgenic, DHA-producing land plant, *Camelina sativa*, containing microalgal genes able to produce LC omega-3 oils, contained 36% lipid by weight with triacylglycerols (TAG) as the major lipid class in hexane extracts (96% of total lipid). Subsequent chloroform-methanol (CM) extraction recovered further lipid (~50% polar lipid, comprising glycolipids and phospholipids) and residual TAG. The main phospholipid species were phosphatidyl choline and phosphatidyl ethanolamine. The % DHA was: 6.8% (of total fatty acids) in the TAG-rich hexane extract and 4.2% in the polar lipid-rich CM extract. The relative level of ALA (α-linolenic acid, 18:3ω3) in DHA-camelina seed was higher than the control. Major sterols in both DHA- and control camelina seeds were: sitosterol, campesterol, cholesterol, brassicasterol and isofucosterol. C16–C22 fatty alcohols, including iso-branched and odd-chain alcohols were present, including high levels of iso-17:0, 17:0 and 19:0. Other alcohols present were: 16:0, iso-18:0, 18:0 and 18:1 and the proportions varied between the hexane and CM extracts. These iso-branched odd-chain fatty alcohols, to our knowledge, have not been previously reported. These components may be derived from wax esters, or free fatty alcohols.

Reprinted from *Nutrients*. Cite as: Mansour, M.P.; Shrestha, P.; Belide, S.; Petrie, J.R.; Nichols, P.D.; Singh, S.P. Characterization of Oilseed Lipids from "DHA-Producing *Camelina sativa*": A New Transformed Land Plant Containing Long-Chain Omega-3 Oils. *Nutrients* **2014**, *6*, 776–789.

#### **1. Introduction**

There are many beneficial health effects in humans of omega-3 long-chain (≥C20) polyunsaturated fatty acids (omega-3 LC-PUFA, also termed LC omega-3 oils). The major bioactive LC omega-3 are EPA (eicosapentaenoic acid, 20:5ω3) and DHA docosahexaenoic acid, 22:6ω3). These are largely obtained through dietary consumption of seafood, primarily fish and fish oil nutraceuticals. The positive effects are across a range of degenerative and inflammatory disorders such as: heart disease, stroke, rheumatoid arthritis, asthma and some cancers, diabetes mellitus, multiple sclerosis, dementia and clinical depression [1–3]. LC omega-3 oils are also important in infant nutrition and are present in high concentrations in brain and retina and are important in the development, health and correct functioning of these organs [4,5]. They are also nutritionally important for the survival, growth and general health of aquaculture species particularly at the larval stage [6]. Highly purified or enriched LC omega-3 oils are also sought after for their bioactive function as potential pharmaceutical products [7].

Future supplies of fish oil derived LC omega-3 oils from fisheries is unlikely to be able to meet increasing demands for their inclusion in aquafeeds, foods, nutraceuticals, or for enrichment to highly purified EPA and DHA, as pharmaceutical bioactives and other products [8,9]. Hence, there is a need to develop new and sustainable sources to supplement oils extracted from wild fish harvests.

It has been recognized that marine fish do not have the capacity to synthesize these oils, rather it is microalgae, at the base of the marine food web, that have the ability to produce these oils [10,11]. Microalgae contain the genes for the various elongase and desaturase enzymes that are responsible for the synthesis of LC omega-3 oils. Members from the brown algal line, including for example the algal class dinoflagellates in particular, produce high proportions of DHA [12–15]. These health-benefitting oils are then accumulated up the food chain into various seafoods, including shellfish and finfish.

The search for new and sustainable sources of LC omega-3 oils has led researchers to attempt to transform land plants with microalgal LC omega-3 genes. As a result, a suite of microalgal genes have successfully been incorporated into a variety of land plants including tobacco leaf [16] and *Arabidopsis* seed [17,18] as a proof of concept, showing that transformed terrestrial plants can serve as an alternative supply of these oils. A little known oilseed plant, *Camelina sativa*, a member of the Cruciferae (Brassicaceae) family is an ancient plant native to Europe and Central Asian areas. It is cultivated as an oilseed crop and used in animal feed mainly in Europe and North America and is known as camelina, gold-of-pleasure or false flax. It has several unique and desirable features which give it a competitive advantage over other commercial oilseed crops such as canola, soybean, and sunflower [19]. It grows well in semiarid, marginal and saline soils and unlike commercial oilseed crops, does not have high nutrient requirements, can tolerate insects and weeds and survive winter sowing and frost and freeze-thaw cycles after emergence during late winter and spring. *Camelina sativa* seeds can produce up to 40% oil containing high proportions of α-linolenic acid (ALA, 28%) and linoleic acid (LA, 19%) [19], which makes this plant a good candidate for transformation into a LC omega-3 oil producing land plant. A new DHA-producing *Camelina sativa* oilseed plant transformed with a suite of microalgal LC omega-3 genes sourced from several target microalgal strains has been recently reported by Petrie *et al*. [20]. The synthesis of DHA occurs by the elongation and desaturation of C18 PUFA through EPA and not by retro-conversion of 24:6ω3. It was also determined by 13C NMR regiospecificity analysis of the TAG-containing oil that DHA is preferentially esterified at the *sn*-1,3 positions of the TAG molecule [20]. Here for the first time we present the lipid class, fatty acid, sterol and fatty alcohol composition of this new transformed DHA-producing *Camelina sativa* oilseed (hereafter termed DHA camelina) and carry out a descriptive analysis with reference to an unmodified *Camelina sativa* control seeds.

#### **2. Experimental Section**

#### *2*.*1*. *Lipid Extraction*

Details of the *Camelina sativa* seed, including: source, the full details of the gene construct inserted, which is composed of a series of elongase and desaturase genes, the metabolic pathway and methods used for the transformation to form the transgenic DHA-producing camelina oilseed event used in this study has been reported by Petrie *et al*. [20]. We used hexane as the extracting solvent since it is the industry standard and it preferentially extracts TAG-containing oil due to its solvating properties and very poor solubilization of polar lipids, particularly at room temperature. We did not use Soxhlet extraction so as to minimize any potential degradation of DHA due to heating for prolonged periods during reflux. No antioxidants were added during extraction or analysis. Subsequent extractions of the hexane-extracted crushed seed using chloroform-methanol were used to exhaustively recover residual TAG and un-extracted polar lipids to determine the effectiveness of extraction by hexane of the TAG-containing oil from the crushed seed. Transformed and control camelina seeds (130 g and 30 g, respectively) were wetted with hexane and crushed using an electric agate mortar and pestle (Retsch Muhle, Germany), transferred to a separatory funnel and extracted four times using a total of 800 mL hexane including an overnight third extraction. For each extraction, extracts were filtered to remove fines through a GFC glass fiber filter, and then rotary evaporated at 40 °C. The extracts were pooled and constituted the TAG-rich hexane extract.

Following extraction with hexane, the remaining meal was further extracted using chloroformmethanol (CM 1:1 v/v) as above, the meal was then removed by filtration and the combined extracts rotary evaporated. The pooled CM total crude extract was then dissolved using a modified Bligh and Dyer [21] one-phase methanol-chloroform-water mix (2:1:0.8 v/v/v). The phases were separated by the addition of chloroform-water (final solvent ratio, 1:1:0.9 v/v/v methanol-chloroform-water). The purified lipid was partitioned in the lower chloroform phase, concentrated using rotary evaporation and constituted the polar lipid-rich CM extract.

#### *2*.*2*. *Lipid Class Analysis*

Lipid classes of the hexane and CM extracts were analyzed by thin-layer chromatography with flame-ionization detection (TLC-FID; Iatroscan Mark V, Iatron Laboratories, Tokyo, Japan) [12] using hexane/diethyl ether/glacial acetic acid (70:10:0.1, v/v/v) as the developing solvent system in combination with Chromarod S-III silica on quartz rods and suitable calibration curves of representative standards obtained from Nu-Chek Prep, Inc. (Elysian, MN, USA). Data was processed using SIC-480II software (SISC Version: 7.0-E). Phospholipid species were separated by applying the purified phospholipid fraction (Section 2.3) obtained from silica column chromatography and developing the rods in chloroform/methanol/glacial acetic acid/water (85:17:5:2, v/v/v) prior to FID detection.

#### *2*.*3*. *Separation of TAG*, *Glycolipid and Phospholipid Fractions from the CM Extracts*

Silica gel 60 (100–200 mesh) (0.3–1 g) in a short glass column or Pasteur pipette plugged with glass wool was used to purify 10 mg of the purified CM lipid extract. The residual TAG fraction in the CM extract was eluted using 20 mL of 10% diethyl ether in hexane, the glycolipids eluted with 20 mL of acetone and the phospholipids eluted in two steps, first 10 mL of methanol then 10 mL of methanol-chloroform-water (5:3:2). This second elution was shown to increase the recovery of phospholipids [22]. The yield of each fraction was determined gravimetrically and the purity checked by TLC-FID. All extracts and fractions were stored in dichloromethane at −20 °C until further analysis by GC and GC-MS.

#### *2*.*4*. *Fatty Acid Methyl Ester Preparation*

Aliquots of the hexane and CM extracts were *trans*-methylated according to the method of Christie [23] to produce FA methyl esters (FAME) using methanol–chloroform–conc. hydrochloric acid (3 mL, 10:1:1, 80 °C, 2 h). FAME were extracted into hexane–chloroform (4:1, 3 × 1.8 mL). The meal (after hexane and CM extraction) was also *trans*-methylated and the value for total lipid was determined by adding the lipid contents of the hexane and CM extracts and the FAME content of the transmethylated meal after solvent extraction**.** 

#### *2*.*5*. *Sterol and Fatty Alcohol Derivatization*

Samples (approximately 10 mg) from the TAG-rich hexane extract and the polar lipid-rich CM extract were saponified separately using 4 mL 5% KOH in 80% MeOH and heated for 2 h at 80 °C in a Teflon-lined screw-capped glass test tube. After the reaction mixture was cooled, 2 mL of Milli-Q water was added and the sterols and alcohols were extracted into 2 mL of hexane: dichloromethane (4:1, v/v, 3×) by shaking and vortexing. The mixture was centrifuged and the extract in the organic phase was washed with 2 mL of Milli-Q water by shaking and centrifugation. After taking off the top sterol containing organic layer the solvent was evaporated using a stream of nitrogen gas and the sterols and alcohols silylated using 200 μL of Bis(trimethylsilyl)trifluoroacetamide (BSTFA, Sigma-Aldrich) and heating for 2 h at 80 °C in a sealed GC vial; free hydroxyl groups were converted to their trimethylsilyl ethers.

#### *2*.*6*. *GC and GC-MS Analysis*

The sterol- and alcohol-OTMSi derivatives were dried under a stream of nitrogen gas on a heating block (40 °C) and re-dissolved in dichloromethane (DCM) immediately prior to GC/GC-MS analysis. The FAME and alcohol/sterol-OTMSi derivatives were analyzed by gas chromatography (GC) using an Agilent Technologies 6890A GC (Palo Alto, CA, USA) fitted with a Supelco Equity™-1 (Bellefont, PA, USA) fused silica capillary column (15 m × 0.1 mm i.d., 0.1 μm film thickness), an FID, a split/splitless injector and an Agilent Technologies 7683B Series auto sampler and injector. Helium was the carrier gas. Samples were injected in splitless mode at an oven temperature of 120 °C. After injection, the oven temperature was raised to 270 °C at 10 °C min<sup>−</sup><sup>1</sup> and finally to 300 °C at 5 °C min<sup>−</sup><sup>1</sup> . Eluted compounds were quantified with Agilent Technologies ChemStation software (Palo Alto, CA, USA). GC results are subject to an error of ±5% of individual component area.

GC-mass spectrometric (GC-MS) analyses were performed on a Finnigan Trace ultra Quadrupole GC-MS (model: ThermoQuest Trace DSQ, Thermo Electron Corporation). Data was processed with ThermoQuest Xcalibur software (Austin, TX, USA). The GC was fitted with an on-column injector and a capillary HP-5 Ultra Agilient J & W column (50 m × 0.32 mm i.d., 0.17 μm film thickness, Agilent Technologies ( Santa Clara, CA, USA) of similar polarity to that described above. Individual components were identified using mass spectral data and by comparing retention time data with those obtained for authentic and laboratory standards. A full procedural blank analysis was performed concurrent to the sample batch.

#### **3. Results**

#### *3*.*1*. *Total Lipid Content*

The "DHA-producing camelina" seeds analyzed here contained slightly less total lipid (36% of seed wt.) than control wild-type *Camelina* (41% of seed wt.).

Of the total lipid, 31%–38% of lipid per seed weight was extracted by hexane for DHA- and control camelina, which accounted for 86% and 92% of total lipid, respectively (Table 1). The chloroform-methanol extraction post hexane extraction recovered a further 4.8% and 2.4% polar lipid-rich extract from the DHA- and control camelina, respectively and the residual lipid released by transmethylation of the remaining solvent extracted oilseed meal was 0.3% and 0.4% of seed weight, respectively.

**Table 1.** Lipid content (as % of seed weight) of DHA- and control *Camelina sativa* seeds after hexane extraction, post hexane chloroform-methanol (CM) extraction and subsequent transmethylation of the extracted meal.


 Polar lipid rich extract containing glycolipid and phospholipid with some residual TAG obtained by CM extraction after hexane extraction of the meal; <sup>2</sup> Residual lipid (FAME) from transmethylated meal after hexane and CM extractions.

#### *3*.*2*. *Lipid Class Analysis*

The TAG-rich hexane extract (Section 2.1) contained 96% TAG in both DHA- and control *Camelina*. The post hexane chloroform-methanol extraction recovered residual TAG amounting to 44% and 13% (of the CM extract), respectively. The chloroform-methanol extracts were rich in polar lipids (glycolipids and phospholipids) amounting to 50% and 76% (of the CM extract) for the DHA- and control camelina, respectively (Table 2). The main phospholipid was phosphatidyl choline (PC) and accounted for 70%–79% of the total phospholipids followed by phosphatidyl ethanolamine (PE, 7%–13%) with smaller relative levels of phosphatidic acid (PA, 2%–5%) and phosphatidyl serine (PS, <2%). There were several other unidentified components separated in the phospholipid fraction (Table 3).


**Table 2.** Lipid class composition (% of total lipid obtained for each extraction step) of Hexane and CM extracts from DHA- and control *Camelina sativa* seeds.

Abbreviations: sterol esters (SE), wax esters (WE), hydrocarbons (HC), triacylglycerols (TAG), free fatty acids (FFA), unknown (UN), sterols (ST), monoacylglycerols (MAG), polar lipids (PL) consisting of glycolipids and phospholipids; <sup>1</sup> SE, WE and HC coelute with this system; <sup>2</sup> May contain fatty alcohols and diacylglycerols (DAG).

**Table 3.** Phospholipid composition (% of total phospholipids) of CM extracts from DHA- and control camelina seeds.


Abbreviations: Unknown (UN), phosphatidic acid (PA), phosphatidyl ethanolamine (PE), phosphatidyl serine (PS), phosphatidyl choline (PC), phosphatidyl inositol (PI); <sup>1</sup> PC is the major component and PI coelutes with PC.



**101** 

### **102** *3*.*3*. *Fatty Acid Composition*

Generally we have found that total seed fatty acid composition obtained by direct transmethylation of whole seed is very similar to that of the TAG fraction, since the bulk of the lipids present in the seed occur in the form of TAG, hence total seed fatty acid composition is not reported here. In DHA camelina, the DHA was distributed in the major lipid fractions (TAG, phospholipids and glycolipids) and the proportion ranged between 1.6% and 6.8% with an inverse relationship between the proportions of DHA and ALA. The TAG-rich hexane extract of the DHA camelina contained 6.8% DHA and 41% ALA (Table 4). The polar lipid-rich chloroform-methanol extract contained 4.2% DHA and 50% ALA. Residual TAG from the polar lipid-rich chloroform-methanol extract contained 6% DHA and 40% ALA. The glycolipid fraction isolated from the chloroform-methanol extract contained 3% DHA and 39% ALA and the phospholipid fraction contained the lowest level of DHA (1.6%) and the highest levels of ALA (54%). The DHA camelina contained higher levels of ALA and lower levels of LA (linoleic acid, 18:2ω6) compared with the control camelina in the major lipid classes (TAG, glycolipids and phospholipids). The proportions of ALA and LA for DHA- and control camelina were: ALA 39%–54% and LA 4%–9% for DHA camelina and ALA 12%–32% and LA 20%–29% for control camelina. The relative level of eurucic acid (22:1ω9) was lower in all fractions in the DHA camelina than in the control (e.g., hexane extracts–1.3% *versus* 2.7%; Table 4).

#### *3*.*4*. *Sterol Composition*

The major sterols in both DHA- and control camelina were: 24-ethylcholesterol (sitosterol, 43%–54%), 24-methylcholesterol (campesterol, 20%–26%) with lower levels of cholesterol (5%–8%), brassicasterol (2%–7%), isofucosterol (∆5-avenasterol) (4%–6%), stigmasterol (0.5%–3%), cholest-7-en-3β-ol, (0.2%–0.5%), 24-methylcholestanol (campestanol, 0.4%–1%) and 24-dehydrocholesterol (0.5%–2%) (Table 5). These nine sterols accounted for 86%–95% of the total sterols, with the remaining components being unconfirmed sterols with partial identification obtained–including the number of carbons and double bonds.

The overall sterol profiles were similar between DHA- and control camelina for both the hexane and chloroform-methanol extracts, although there were slightly higher levels of unknown sterols in the chloroform-methanol extracts of both the DHA- and control camelina than occurred in the hexane extracts (10%–14% and 4%–7%, respectively).


**Table 5.** Sterol composition (% of total sterols) of DHA- and control camelina.

Abbreviations: UN denotes unknown, C is number of carbon atoms and db denotes number of double bonds; 1 Polar lipid-rich extract containing glycolipids, phospholipids and residual TAG recovered by chloroform-methanol (CM) extraction post hexane extraction of crushed seed.

#### *3*.*5*. *Fatty Alcohol Composition*

A series of fatty alcohols from C16–C22, with accompanying iso-branched fatty alcohols, were identified in both the hexane and chloroform-methanol extracts (Table 6). Similar profiles were observed for the DHA- and control camelina, with some variation in the proportions of individual components observed. The odd-chain alcohols were present at higher levels in the chloroform-methanol extract (37%–38%) than in the hexane extract (16%–23%). Iso-17:0 (16%–38%) predominated over 17:0 (0.3%–5.7%). Another odd-chain alcohol present was 19:0 (4.5%–6.5%). Phytol, derived from chlorophyll, was the major aliphatic alcohol and accounted for 47% and 37% of the total fatty alcohols in the hexane fractions in the DHA- and control camelina, respectively. There were lower levels in the chloroform-methanol extract (9% and 12%, for DHA- and control camelina, respectively). Other alcohols detected included iso-16:0, 16:0, iso-18:0, 18:1, 18:0, with minor levels of iso-20:0, 20:1, 20:0, iso-22:0, 22:1 and 22:0 also present.


**Table 6.** Fatty alcohol composition (% of total sterols) of DHA- and control camelina.

1 Polar lipid-rich extract containing glycolipids and phospholipids with residual TAG obtained by chloroform-methanol (CM) extraction post hexane extraction of crushed seed.

#### **4. Discussion**

The oil content from camelina seeds can range from 25% to 48% [19,24–26] and can be dependent on the location where the plant is grown and the environmental conditions during growth. Previous researchers have published the fatty acid composition of camelina oil which is similar to what we report here for the control camelina [27]. Since this was only a descriptive analysis with no replication performed, we cannot yet conclusively determine whether the insertion of the omega-3 LC-PUFA microalgal-derived genes affected the oil yield. Hence, further work including replication would need to be done to determine if there is any statistical change in the oil yield. The results for the oil extraction suggest that slow crushing using a motorized mortar and pestle with multiple extractions with hexane at room temperature is effective in recovering the majority of the TAG oil.

In addition to the oil containing moderate levels of DHA, the DHA-containing camelina also had markedly higher levels of ALA in the major lipid classes (triacylglycerols, glycolipids and phospholipids) compared with the control camelina. This finding shows that the activity of the ∆-15 desaturase gene is considerably enhanced in DHA camelina [18,20] hence there is scope to increase the DHA content further by optimizing the elongation and desaturation of ALA. Variations in the level of ALA and DHA in the transformed plants may also be influenced by the effects of cultivar variety and other factors such as the quality of soil and climatic and weather conditions. It has also been reported that, in oilseed crops, the level of PUFA in general is promoted by low temperatures (winter and spring season) during the seed filling period, while at higher temperatures (summer season) the concentration of saturated fatty acids is higher [26].

Interestingly, there were some slight differences in the fatty acid profile and proportion of DHA in the various extracts and fractions with the DHA levels being higher in the TAG-rich hexane extract and TAG from CM extraction (6%–6.8%) and lower in the polar lipid fractions (3% in glycolipids and 1.6% in phospholipids), and 16:0 being higher in the polar lipid fractions of glycolipids and phospholipids from CM extraction (19%–21%) compared with the TAG-rich hexane extract and TAG from CM extraction (6%–7%). It is not known if the low levels of residual lipid present in the meal after solvent extraction (using hexane then chloroform-methanol) and recovered by transmethylation is derived from free solvent extractable lipid or bound lipid which is liberated only by hydrolysis under the hot acid methanolic transmethylation conditions. Future research will investigate this aspect.

The sterol composition of the camelina DHA and control camelina samples analysed here were similar to that found in refined camelina oil [28] with the same major sterols present, indicating that the added genes did not affect sterol synthesis. Previous workers reported the major sterols as being: cholesterol (4%), brassicasterol (3%), campesterol (21%), stigmasterol (2%), sitosterols (45%), ∆5-avenosterol (9%), cycloartenol (12%) and 24-methylene cycloartenol (3%). They also noted ten minor components which were unidentified due to their very low levels. We also observed several unidentified components at low relative levels (Table 5). Schwartz *et al*. [29] reported campestanol (1.6 mg), sitostanol (2.5 mg), stigmasta-5,24-dienol (6.2 mg), gramisterol + α-amyrin (1.9 mg), ∆7-avenosterol (trace levels) and citrostadienol (1.30 mg) in camelina oil in addition to the sterols identified by Shukla *et al*. [28]. They reported the levels of the common sterols (mg/100 g oil) in camelina oil as cholesterol (35 mg), brassicasterol (27 mg), campesterol (117 mg), stigmasterol (5.6 mg), sitosterol (300 mg), ∆5-avenosterol (37 mg), cycloartenol (10 mg) and 24-methylene cycloartenol (1.0 mg). The level of cholesterol in camelina oil was higher than occurs in most vegetable oils and brassicasterol is a characteristic sterol found in the Brassicaceae family of which camelina is one.

Based on the combined analyses of a wide suite of lipid classes in DHA camelina and control camelina, it would seem then that the omega-3 LC-PUFA genes have, as expected, had little or no effect on the sterol composition. Further work will need to be carried out to determine the contributions of these sterols from the sterol ester and free sterol fractions, in each of the extracts, by separating the sterol ester and free sterol lipid classes and analysing them separately.

In relation to the presence of fatty alcohols, those with chain length C16–C24 have previously been observed in camelina after release from wax esters following saponification [30], but to our knowledge the presence of iso-odd-chain fatty alcohols such as found here, in both DHA camelina and control camelina, have not been reported previously. Further research needs to be performed to determine which fractions these alcohols are present in, e.g., in free or in an esterifiedform such as in a wax ester.

#### **5. Conclusions**

This is the first detailed report of the lipid composition of a new transformed terrestrial oilseed capable of producing DHA-containing (6.8% of total fatty acids) TAG oil with a simple fatty acid profile and a high preference of ω3 over ω6 LC-PUFA. Fish oils also have a high preference of ω3 over ω6 LC-PUFA. However, several key distinguishing features found in the transformed camelina fatty acid profile is the high level of α-linolenic acid (ALA, 18:3ω3) which is only a minor fatty acid in fish oils and the much lower levels of EPA (eicosapentaenoic acid, 20:5ω3) making the fatty acid profile of the oil unique. The chloroform-methanol extract is rich in polar lipids (glycolipids and phospholipid), since these are not extracted by hexane. Hence, the hexane extracted seed meal, from which most of the TAG was removed may be useful as an animal feed supplement (e.g., in aquaculture) or as a source of high value DHA-containing polar lipids (e.g., phosphatidyl choline containing DHA). The profiles of other lipid classes such as sterols and fatty alcohols were very similar to the control camelina. These results hold promise for the development and commercial production of new and sustainable terrestrial sources of LC omega-3 oils, which will supplement or in part replace LC omega-3 containing marine oils, hence alleviating pressure on wild harvest fisheries arising from increasing demand for these oils. Future research will be extended to canola and include similar detailed lipid class characterisation, examination of extraction efficiencies, oil stability and enrichment of DHA.

#### **Acknowledgments**

This research was funded by CSIRO Food Futures Flagship, Nuseed Pty Ltd. Laverton, Australia and the Australian Grains Research and Development Corporation, Canberra, Australia. The funders had no role in study design, data collection and analysis, or preparation of the manuscript. The authors were granted permission to publish the manuscript by the funders. We acknowledge Daniel Holdsworth who managed the CSIRO Marine and Atmospheric Research laboratories GC-MS facility and Diana Davies, Graeme Dunstan and two unknown reviewers for reviewing the manuscript and providing useful suggestions.

#### **Conflicts of Interest**

The authors declare no conflict of interest.

#### **References**


## **DHA-Containing Oilseed: A Timely Solution for the Sustainability Issues Surrounding Fish Oil Sources of the Health-Benefitting Long-Chain Omega-3 Oils**

#### **Soressa M. Kitessa, Mahinda Abeywardena, Chakra Wijesundera and Peter D. Nichols**

**Abstract**: Benefits of long-chain (≥C20) omega-3 oils (LC omega-3 oils) for reduction of the risk of a range of disorders are well documented. The benefits result from eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA); optimal intake levels of these bioactive fatty acids for maintenance of normal health and prevention of diseases have been developed and adopted by national and international health agencies and science bodies. These developments have led to increased consumer demand for LC omega-3 oils and, coupled with increasing global population, will impact on future sustainable supply of fish. Seafood supply from aquaculture has risen over the past decades and it relies on harvest of wild catch fisheries also for its fish oil needs. Alternate sources of LC omega-3 oils are being pursued, including genetically modified soybean rich in shorter-chain stearidonic acid (SDA, 18:4ω3). However, neither oils from traditional oilseeds such as linseed, nor the SDA soybean oil have shown efficient conversion to DHA. A recent breakthrough has seen the demonstration of a land plant-based oil enriched in DHA, and with omega-6 PUFA levels close to that occurring in marine sources of EPA and DHA. We review alternative sources of DHA supply with emphasis on the need for land plant oils containing EPA and DHA.

Reprinted from *Nutrients*. Cite as: Kitessa, S.M.; Abeywardena, M.; Wijesundera, C.; Nichols, P.D. DHA-Containing Oilseed: A Timely Solution for the Sustainability Issues Surrounding Fish Oil Sources of the Health-Benefitting Long-Chain Omega-3 Oils. *Nutrients* **2014**, *6*, 2035–2058.

#### **1. Introduction**

There is a vast array of reviews, systematic reviews, meta-analysis and industry reports on seafood consumption in general and the health benefits of omega*-*3 long-chain (≥C20) polyunsaturated fatty acids (omega LC-PUFA) (also termed LC omega-3 oils) in particular. Our review begins from the premise that, as far as human health claims are concerned, the health benefits of seafood and their omega-3 fatty acids are largely related to the supply of eicosapentaenoic acid (EPA, 20:5ω3) and docosahexaenoic acid (DHA, 22:6ω3) [1–9]. Thus, one aim of our review is to examine the occurrence of a marked rise in demand for oils high in EPA and DHA and against this demand, the sustainability of marine sources of these LC omega-3 oils. The review summarises recommended intake targets proposed and/or set by national and international bodies. It analyses the bio-conversion of shorter-chain (C18) omega-3 fatty acids to EPA and DHA in humans, monogastric animals, ruminants and aquaculture species. As the major user of global fish oil (FO) supply, the aquaculture sector is considered in a relatively greater detail with respect to both biosynthesis of EPA and DHA as well as FO supply issues. Research trends in the area of metabolic engineering of plants to produce novel omega-3 oil sources, and the biological efficacy of currently available oils are covered.

### **2. Consensus for, and Recommendations on, Human Nutrition Needs for Long-Chain Omega-3 Oils**

Increasing numbers of clinical and epidemiological studies provide evidence supporting the case that omega-3 LC-PUFA are responsible for a multitude of health benefits [10]. The dietary intake of preformed omega-3 LC-PUFA—EPA and DHA—has been recognised as important [1] since *in vivo* conversion of the shorter-chain (C18) fatty acids, namely ω-linolenic acid (ALA, 18:3ω3) to DHA in particular is relatively poor [11,12]. In 2009, the International Society for the Study of Fatty Acids and Lipids (ISSFAL) [13] provided the following position statement: "The majority of evidence from isotopic tracer studies shows that the conversion of ALA to DHA is on the order of 1% in infants, and considerably lower in adults." Hence, it is critical that there are various and sustainable sources of oils containing preformed DHA for the whole population to meet targets for adequate intake of EPA plus DHA. Table 1 summarises the dietary intake targets proposed by various national and international bodies [14–20].

Although the recommended daily intakes vary, they are a culmination of years of clinical and epidemiological research that have clearly established a strong body of evidence for the beneficial health effects of LC omega-3 oils for the improvement of cardiovascular health [21]. The recommendations provided in Table 1 are based on consumption of both EPA and DHA. As currently there are only limited and more niche supply alternatives to the marine supply of DHA, we also aim to highlight specific roles of DHA in the following section to caution against reliance on LC omega-3 sources that do not provide DHA.

**Table 1.** Selected suggested long-chain (LC) omega-3 (eicosapentaenoic acid + docosahexaenoic acid (EPA + DHA)) intakes (mg/day) for adults available from various agencies and bodies.


\* For secondary prevention of coronary heart disease.

#### **3. The Role of DHA**

LC omega-3 oils containing EPA and DHA are considered beneficial for certain aspects of cardiovascular health and pharmaceutical-grade omega-3 LC-PUFA therapies have expanded rapidly for treatment of cardiovascular-related diseases [22–24]. Whilst the majority of studies have reported both EPA and DHA as being protective, there is a growing body of evidence that suggests differential effects depending on the nature of cardiovascular risk factor itself and/or the disease endpoint. For example, DHA has been more effective than EPA for its actions on blood pressure, heart rate and vascular health [10,25–27]. On their effects on plasma lipids, a meta-analysis of randomized placebo controlled trials of EPA or DHA monotherapy has concluded that DHA is more effective in lowering triacylglycerols (TAG) and raising HDL-cholesterol (HDL-c) than EPA [28]. Although DHA has been found to raise LDL-cholesterol (LDL-c), this was also associated with increased LDL and HDL particle sizes [29]—an outcome not observed with EPA [26]. Furthermore, only DHA was effective in reducing the number of small, dense LDL particles [27] which are known to be more atherogenic. Increased LDL and HDL particle sizes are negatively correlated with cardiovascular risk. Furthermore, DHA, but not EPA, was inversely associated with intima-media thickness—an independent predictor of cardiovascular events—in the Japanese [30], suggesting more potent anti-atherogenic properties of DHA. Similarly, DHA but not EPA supplementation reduced the vulnerability to experimentally-induced atrial fibrillation and secondary structural changes (re-modeling) of the atria [31]; these findings are in agreement with observations made in several human clinical studies that reported that lower incidence of atrial fibrillation is correlated positively with plasma DHA, but not EPA [32,33].

The human nervous system contains a significant amount of DHA, which is required for brain development and function especially in infants [12]. With the poor conversion of ALA and EPA to DHA, together with the particularly important roles for DHA in humans, inclusion of DHA in infant formula is now widespread.

With regard to mental health conditions, a range of studies have examined the effect of the LC omega-3 oils on mild cognitive impairment (MCI) [34]. Depressive symptoms may increase the risk of progressing from MCI to dementia. Consumption of LC omega-3 may alleviate both cognitive decline and depression. A recent study investigated the benefits of DHA and EPA supplementation for depressive symptoms, quality of life (QOL) and cognition in elderly people with MCI [34]. In a 6-month, double-blind, randomised controlled trial, individuals aged 65 years with MCI were allocated to receive a supplement rich in EPA, DHA or the omega-6 PUFA linoleic acid (LA, 18:2ω6). Compared with the LA group, Geriatric Depression Scale (GDS) scores improved in the EPA and DHA groups and verbal fluency (Initial Letter Fluency) improved in the DHA group. Improved GDS scores were correlated with increased DHA plus EPA. Improved self-reported physical health was associated with increased DHA. There were no treatment effects on other cognitive or QOL parameters. Increased intakes of DHA and EPA benefited mental health in older people with MCI. Increasing omega-3 LC-PUFA intakes may reduce depressive symptoms and the risk of progressing to dementia. The authors concluded that this needs to be investigated in larger, depressed sample groups with MCI.

The same research team also examined the effects of LC omega-3 oils on literacy and behaviour in children with attention-deficit/hyperactivity disorder (ADHD) [35]. The effects of an EPA-rich oil and a DHA-rich oil *versus* an omega-6 PUFA-rich safflower oil (control)—as LA—were compared in a randomized, controlled trial. The effect of supplementation on cognition, literacy, and parent-rated actions was assessed by linear mixed modelling. There were no significant differences between the supplement groups in the primary outcomes after four months. However, the erythrocyte fatty acid profiles indicated that an increased proportion of DHA was associated with improved word reading and lower parent ratings of oppositional behaviour. These effects were more evident in a subgroup of children with learning difficulties: an increased erythrocyte DHA was associated with improved word reading, improved spelling, an improved ability to divide attention, and lower parent ratings of oppositional behaviour, hyperactivity, restlessness, and overall ADHD symptoms. The authors concluded that increases in erythrocyte omega-3 PUFA, specifically DHA, may improve literacy and conduct in children with ADHD. The greatest benefit may be observed in children who have co-morbid learning difficulties. In a recent randomized controlled intervention trial, DHA supplementation was observed to improve both memory and reaction time in healthy young adults whose habitual diets were low in DHA [36]; the response was found to be modulated by gender.

Another very recent application of DHA has been in the development of neuroprotective strategies for treatment of spinal cord and head injuries. These studies—albeit at early stages involving animal models—illustrate the significant potential of DHA, but not EPA, in the treatment of acute neurological injury [37].

Against the increasing scientific literature pointing to the importance of the LC omega-3 oils and in particular DHA, in human health, global supplies of fish oil (the current main source of EPA and DHA) obtained from wild-harvest low-value marine species, termed forage fish, will not meet future market demands [23,38]. The following sections in this review paper examine sustainability of fish oil and future sources of the LC omega-3 oils, recent findings for the use of SDA containing oils, current practices with aquafeeds, and further research needs.

## **4. Supply, Demand and Environmental Issues—A Need for Alternative Sources of LC Omega-3 Oils**

The harvest of low trophic species such as anchovy, sardines, mackerel, menhaden, capelin and sandeel for the production of fish meal and fish oil (FO) represents the current main source of the health-benefiting LC omega-3 oils used in aqua and animal feeds, health supplements, pharmaceuticals and other products including functional foods.

Fish oil processing involves a range of steps after the initial meal and oil production [39]. Many of these steps use the same processes that are used for vegetable oils. The final use for the oil determines the level of processing, with human nutrition and pharmaceutical applications generally requiring greater processing and accompanying quality assurance and quality control procedures than for fish and animal nutrition. Several changes in the usage pattern have occurred over the past decades. FO was initially widely used in livestock feed, then, as aquaculture expanded this industry became the major user. Therapeutic uses of LC omega-3 PUFA are increasingly being recognized and recently new pharmaceutical grade LC omega-3 products (containing 85%–95% EPA and DHA) have gained increasing market share, and this industry has expanded its share of use of the fish oil resource, with less oil available for the aquaculture sector [31,40]. This changing pattern of use of FO is predicted to continue [40]. One aspect of this recent change is that the processing of the FO to achieve the higher grade (or more pure) LC omega-3 products results in considerable losses, with product yields in the 5%–10% range. For instance, in a purification process reported by Belarbi *et al.* [41], production of 1 kg EPA ester required 15 kg of FO, or 56 kg (dry basis) of the marine alga *Phaeodactylum*  *tricornutum* or 70 kg of another alga *Monodus subterraneus*. Further research and development for more efficient production of the pharmaceutical grade products will assist all users to maximize and better utilize this important finite resource.

Global production of FO is around 1 million tonnes per annum, with fish meal in the range of 6–7 million tonnes per annum, except during the periodic El Niño years [42]. Production has generally remained at this level for the past decade [23], and requires an annual catch of 25–30 million tonnes of feed-grade fish and unwanted fish processing waste; 4–5 kg of wet fish yields 1 kg of FO and fishmeal. Although the current harvest of low trophic (also termed forage) species for fish meal and FO has been regarded as sustainable for several decades, a recent development has been the foreseen need to reduce the harvest of small oceanic forage fish like sardines and anchovies in some areas by 20%–50% in order to protect larger predators that rely on these species for food [43,44]. Should such recommendations be implemented, this would have significant flow-on ramifications for the range of industries currently utilizing the FO resources.

An emerging source of LC omega-3 oils over the past decade has been krill oil. The current krill harvest is around 200,000 MT [40], and is actually much less than in the 1980s prior to the break-up of the USSR. The total allowable catch is set by the Committee for the Conservation of Antarctic Marine Living Resources (CCAMLR) and is three times greater than the present harvest. Of the major krill oil producers, a further development has been that Aker BioMarine has been granted certification by the Marine Stewardship Council (MSC) in 2012. MSC has validated the harvesting and traceability for Aker's Antarctic fisheries. The total krill harvest is deemed sustainable at present levels in view of a number of, although not all, bodies including various Non-Government Organizations. If there is to be a major expansion of the krill fishery, the sustainability topic will clearly need to be revisited by these and other expert groups including in particular CCAMLR.

#### **5. Alternative Sources of LC Omega-3 Oils**

Against the background provided above on the current and future status of marine-derived oils—fish oil and more recently krill oil—considerable progress has occurred with the development of new, alternate and sustainable sources of the LC omega-3 oils. Single cell organisms (SCO), such as heterotrophic dinoflagellates and thraustochytrids (both grown and harvested for DHA containing oils) and other algal groups and recently a genetically modified (GM) yeast (containing EPA) are now in commercial production; strong uptake has occurred for the DHA oils in particular areas including infant formula, health supplements and some functional foods [45]. In addition to this excellent progress with SCO production of the LC omega-3 oils, a large number of groups are conducting research and development with a suite of microalgae towards co-production of biofuels and other by-products [46]. Whilst the cost of production of microalgae for biofuel production is presently greater than for fossil fuels, future breakthroughs in culturing, harvesting and other processing are anticipated [46], which can be expected to reduce costs. It is anticipated that these uses for SCO will largely remain for the high value applications including in nutraceutical and pharmaceuticals rather than in aquafeeds.

The past decade has also seen several groups using genetic engineering to allow oilseed crops to produce LC omega-3 oils [23]. As this research field has progressed, important breakthrough steps have included: the isolation and characterization of genes from the marine microalgae encoding front-end desaturases involved in DHA biosynthesis [47], the isolation of highly efficient desaturases and elongases [48–51], the use of genes with omega-3 substrate preference [49–51] and the development and use of a land plant (tobacco) leaf-based assay using interchangeable design principles to rapidly assemble multistep recombinant pathways [52]. Progress with research on insertion of microalgal-derived genes leading to DHA production into a range of omega-3 C18 PUFA accumulating land plants has been reviewed [2,53–55]. Transfer of genes from microorganisms to land plants has seen accumulation in oilseeds of SDA, EPA and DHA [2] (Table 2, Figure 1). Good progress has been made in engineering the EPA genes into crop plants, with several groups reporting the production of EPA at levels similar to that observed in bulk fish oil (approximately 18%) [56,57]. The conversion of

the C20 EPA to the particularly important C22 DHA, however, had been problematic with many attempts resulting in the accumulation of EPA and DPA, but until very recently little DHA [56,58–62]. For SDA and EPA, levels achieved in the engineered oilseed plants are comparable to levels from

other naturally occurring land plant (SDA) and/or marine (EPA) sources [23]. Elevated levels of DHA had not been achieved prior to 2012, except for one *in planta* observation where the isolated TAG fraction from the leaf of *Nicotiana benthemiana* contained high DHA, although high levels of the omega*-*6 PUFA 18:2ω6 were also present [53]. In 2012, a further key breakthrough occurred, with the reporting, for the first time, of fish oil-like profiles for a DHA-containing oilseed plant *Arabidopsis thaliana* [58]. Features of the new oil were: (i) a DHA level of 15% (of the total FA); (ii) a total of 25% new omega-3 PUFA and omega-3 LC-PUFA; (iii) 30% ALA; and (iv) an omega-3/omega-6 ratio that was similar to that observed for marine oils. The latter feature is a further important and distinguishing attribute for the land plant derived LC omega-3 oils, with this report being, to our knowledge, the first time this oil trait has been observed. More recently a similar profile has been observed in a commercial oilseed plant (*Camelina sativa*) [59].


**Table 2.** Levels of SDA, EPA and DHA (as % of fatty acids) in new land plant oil seeds.

Abbreviations: SDA, stearidonic acid EPA; EPA, eicosapentaenoic acid; DHA, docosahexaenoic acid.

These findings clearly indicate the feasibility of developing oilseed crops with high concentrations of omega-3 LC-PUFA. Further research and development is needed to develop commercial oilseed crops with LC omega-3 oil enriched in DHA as has been achieved in model plants (*Arabidopsis,* tobacco) and also in *Camelina*. Future novel land-based plants can provide an economically viable source of LC omega-3 oil for aquaculture and other higher value applications. A land plant source of LC omega-3, if achieved, and assuming cultivation is permitted, will be considerably cheaper than that from microorganisms, and could be used as an additional source of these essential ingredients in feed, food and pharmaceutical products to improve human health (Figure 1).

#### **6. ALA and SDA in Animal Feeds**

The availability of the SDA-containing *Echium* oil (around 14% SDA of total FA) has enabled this potential EPA/DHA precursor, which is one step more advanced than ALA to be trialed in animal, including farmed fish, and human nutrition research. The general hypothesis driving this line of research has been that animals and farmed fish fed SDA oils would produce tissue containing greater proportion of EPA and DHA than those fed ALA oils. This is based on the assumption that the inefficiency in the biosynthesis of EPA and DHA from ALA is related to the rate-limiting step of converting ALA to SDA (Figure 2).

Selected poultry data from feeding experiments [66–74] where different oils have been used to enrich thigh muscle are summarised in Figure 3. There were only modest changes in EPA and DHA content in broiler muscle samples when the oil supplement itself did not contain the omega-3 LC-PUFA. The evidence from such a large range of experiments did not indicate marked benefit from using C18 oils in enriching tissues particularly with DHA (Figure 3). Similar trends were noted when we reviewed EPA and DHA enrichment of egg from omega-3 oil-supplemented laying hens and breast muscle in broilers (data not shown). Similarly, data from lamb meat studies [75–84] are summarized in Figure 4. Levels of EPA and DHA in lamb meat were lowest in studies where vegetable oils were used. As the dietary fat supplement shifted towards marine sources, the levels of EPA and DHA in lamb meat increased across experiments. The evidence so far suggests that the best way to enhance the DHA content of livestock products is to include DHA containing fat supplements in the diet. There is as yet no convincing evidence that current fat supplements containing ALA or SDA are suitable alternatives for those containing preformed EPA and especially DHA.

**Figure 1.** The potential future sources of omega-3 LC-PUFA are shown, with current sources (left) being seafood and microalgae, with possible future sources through genetically engineered plants also indicated at the right.

**Figure 2**. Schematic showing synthesis of shorter-chain fatty acids in land plants (**black horizontal arrows**), followed by addition of genes from marine microalgae (**blue vertical arrows**) resulting in new LC omega-3 containing oilseeds. elo, elongase; des, desaturase.

**Figure 3.** Levels of omega-3 PUFA (ALA) and omega-3 LC-PUFA (EPA, DPA and DHA) in thigh muscle from broilers on a diet with: (1) no oil supplement; (2) vegetable oil; (3) fish oil; (4) marine algae; or (5) SDA-containing oil [66–74].

**Figure 4.** Levels of omega-3 PUFA (ALA) and omega-3 LC-PUFA (EPA, DPA and DHA) in trimmed lean muscle of lambs on a diet with: (1) no oil supplement; (2) linseed or linseed oil; (3) fish oil–vegetable oil mix; (4) fish oil; or (5) fish oil–marine algae mix [75–84].

#### **7. ALA and SDA in Aquafeeds**

Aquaculture continues to be the main end user of global sources of FO, yet available sources of FO generally remain static and will likely not increase significantly. Against this background of either static or decreasing resource availability, the demand for FO and its range of uses continues to increase as noted earlier. The increasing demand for FO is in line with the growing global population (due to rise by a further 34% by 2050) [85], the expanding aquaculture industry (around 10% growth per annum), the increasing recognition of the health benefits of the LC omega-3 oils, and more recently the resulting and therefore competing use of FO for production of pharmaceutical grade products containing ≥85% EPA and/or DHA.

A number of trials have been conducted for Atlantic salmon and barramundi using the SDA rich *Echium* oil *(Echium plantagineum)*. Initial work with Atlantic salmon parr (freshwater stage) showed that SDA was effective in producing EPA and DHA [86,87]. This freshwater phase is only a short period of the total life cycle for farmed Atlantic salmon. Trials for the same species during the seawater stage (bulk of the life cycle) showed SDA was not so effective [63,64,88]. Some EPA was produced and also DPA, but no DHA. The same observations occurred for barramundi feeding trials [89–91], that is limited or no omega-3 LC-PUFA, in particular DHA, was produced or accumulated in the flesh of this species. Conversion of the C18 SDA for barramundi to the omega-3 LC-PUFA was even lower than that observed for Atlantic salmon. Other researchers have generally observed similar findings where SDA inclusion has occurred in aquafeed trials with other fish species, including Atlantic cod, striped bass, rainbow trout and gilthead seabream [92–95].

A further feeding trial with the SDA-containing *Echium* oil for early juvenile barramundi used the plant bioactive sesamin as a potential modulator of lipid biosynthesis [96]. Relative to the control fish, growth of the SDA treatment group was hindered, although interestingly both EPA and DHA increased. It was proposed that sesamin is a potent modulator for LC-PUFA biosynthesis in barramundi, but probably will have more effective impact at advanced ages. By modulating certain lipid metabolic pathways, the use of sesamin as a feed ingredient probably disrupted the body growth and development of organs and tissues in the early juvenile barramundi.

Atlantic salmon and barramundi have, when fed an FO-containing diet, provided an excellent source of beneficial omega-3 LC-PUFA for human consumption, but reduced concentrations of these acids, together with a markedly decreased omega-3/omega-6 ratio, as occurs through the use of vegetable oil and/or animal fat diets, may compromise their nutritional benefit to consumers. Limited research has been performed to examine this issue. In one study [97], dietary intake of differently fed salmon (100% FO, 50/50 FO/rapeseed oil, 100% rapeseed oil) and the influence on markers of human atherosclerosis were compared. Significant differences between the consumer groups were observed in the serum fatty acid profiles, especially for the levels of total omega-3 PUFA and the omega-3/omega-6 ratio, which were markedly increased in the FO-fed fish consuming group in contrast to the two other groups. The authors concluded that Atlantic salmon fed the FO-containing diet and containing very high concentrations of omega-3 LC-PUFA seemed to impose favorable biochemical changes in patients with CHD when compared with ingestion of fillets with intermediate and low levels of the marine omega-3 LC-PUFA, where FO was replaced in part or in full by rapeseed oil [93]. There have been no consumer trials with fish fed diets containing ALA / SDA rich oils *versus* FO derived EPA + DHA, and looking at the effects on consumers.

#### **8. SDA Oils in Animal Models and Humans**

It is apparent from examination of the research performed to date on SDA diets, that the benefits from use of SDA (like for ALA) are due to its conversion to EPA and DHA. Whilst SDA is more efficiently converted to EPA than ALA [98], it is important to record that the elongation of dietary SDA to DHA has been found to be absent (or negligible at best) in humans [99–101]. Furthermore, with respect to enrichment with EPA, the conversion efficiency of SDA to EPA was only 17% even after four months of feeding soybean oil preparation containing 16% SDA and 11% ALA. Similarly, three months treatment with soybean oil with even higher SDA content (28% SDA alone and 40% total omega-3) failed to change erythrocyte DHA from baseline values. These studies clearly show the inability of SDA-rich oils to influence the endogenous DHA pool in humans. The elongation and desaturation of SDA appears to terminate at the DPA level as increased levels of this fatty acid have been observed [101]. However, evidence of direct physiological benefits of DPA in humans is yet to be elucidated. The accumulation of DPA in the EPA and DHA biosynthesis pathways of many species, and its relative abundance in red meat and some marine species also point to a need to determine whether or not DPA should be included in the omega-3 content claim of foods and ingredients. Inclusion of DPA will broaden the range of foods that can reach the "good source" and "very good source" bars in the omega-3 content claims.

Similar results have also been found following animal feeding studies where dietary *Echium* oil rich in SDA (and ALA; 15% SDA, 29% ALA) failed to increase plasma or tissue DHA levels compared to supplementation with fish oil. In particular, *Echium* oil diet did not lead to any increase in EPA or DHA in cardiac muscle membranes, but resulted in a dose-related increase in DPA [102]. Fish oil

feeding on the other hand displayed considerable accumulation of DHA but not DPA. Also it is of interest to note that anti-arrhythmic action (protection against ischemia induced cardiac arrhythmia and sudden cardiac death in rats) was significantly greater following feeding with FO compared to SDA-rich *Echium* oil [65].

Albert *et al.* [103] reported over a decade ago that plasma LC omega-3 levels are inversely associated with the risk of sudden cardiac death. More recently, the omega-3 index which is the combined total proportion of erythrocyte membrane EPA and DHA has emerged as a novel biomarker that predicts cardiovascular risk [104]. It has also been reported that the omega-3 index correlates well with the EPA + DHA levels of myocardial membranes [105], thus favouring its use when assessing the risk of sudden cardiac death. Since there was no increase in DHA following feeding oils rich in SDA [99–101], the reported beneficial rise in omega-3 index has been driven solely by an increase in EPA. In view of the overall cardiovascular benefits specific to DHA, including the anti-arrhythmic actions discussed above, it can be concluded that further research is needed to determine the important question whether or not an omega-3 index that increased solely due to EPA, is of less benefit as compared to rise in omega-3 index achieved via greater incorporation of DHA into erythrocytes, and ultimately whether an increased SDA consumption would translate into improved human health outcomes [101]. Taken collectively, the observations summarized in these two sections covering the potential use of SDA-rich oils presently imply that direct supply of the pre-formed omega-3 LC-PUFA (EPA and in particular DHA) is the preferred strategy to improve the omega-3 status in diverse applications ranging from aquaculture, livestock and in humans.

#### **9. Current Practices with Commercial Aquafeeds and Future Sources of LC Omega-3 Oils**

Aquaculture can be considered as a traditional industry with fish culture occurring for many centuries. Modern aquaculture expansion began in the 1980s and has continued to rise steadily since, with the high value salmonoid fish such as Atlantic salmon being the species of choice. Fish oil, produced as a by-product of the fish meal industry, had been the main oil incorporated into fish feed until recent years. As noted earlier, the past decade has seen fish oil availability decrease and also prices increase substantially. From the use of 100% FO (of the oil component), feed manufacturers are now using up to 75% or higher of vegetable or animal-derived oils [106]. The topic of FO replacement and alternative lipid sources has been recently examined in considerable detail, with a review book now available for researchers, industry and other end users [107]. Given the availability of such a substantial resource, it is not the purpose of this section to further review this topic. This FO substitution can include mixes such as FO/rapeseed oil, FO/chicken fat, and also other combinations. Whilst fish growth and performance is generally not affected, the concentration of omega-3 LC-PUFA and the omega-3/omega-6 ratio in fillet products is markedly changed. For farmed Atlantic salmon grown in Tasmania, Australia, the concentration of EPA and DHA has reduced by ≥30%–50% or more (Figure 5), and the omega-3/omega-6 ratio also has reduced markedly. Marine fish typically show an omega-3/omega-6 ratio of between 5 and 10, and for the first time in 2013 the ratio in farmed salmon in Australia has decreased to less than 1 [108].

**Figure 5**. Farmed Atlantic salmon from Tasmania, Australia from 2002 (fish oil diet) [23] and 2010 to 2013 (chicken fat/fish oil diet) [108]: Content of EPA (**white bars**) and DHA (**black bars**) (mg/100 g, wet weight).

A recent study tested whether Atlantic salmon smolt fed a diet with a higher DHA: EPA ratio and a lower content of LC omega-3 oils to that of conventional FO based diets would enhance deposition of LC omega-3 in the liver and muscle [109]. Comparisons were made between fish fed: (1) a FO diet; (2) a blend of 50% rapeseed and 50% tuna oil diet (termed model oil, MO1); (3) a blend of 50% rapeseed, 25% tuna and 25% FO diet (MO2); and (4) a blend of 50% FO and 50% chicken fat diet (FO/CF). The latter diet was representative of commercial diets in use at the time of the study, with the proportion of chicken fat increasing even further since the study was performed. The dietary DHA:EPA ratio was in the order MO1 > MO2 > FO/CF ~ FO. The LC omega-3 content was approximately 2-fold lower in the MO1, MO2 and FO/CF diets compared to the FO diet, with the relative levels (as % total FA) lowest in the MO1 diet. For the feeding trial, there were comparable contents of LC omega-3 in the muscle of the FO, MO1 and FO/CF fed fish. A major outcome was that a higher DHA:EPA ratio than that commonly occurring with FO-only diets used for Atlantic salmon was better suited for more efficient deposition of LC omega-3, in particular DHA, with evidence therefore apparent for LC omega-3 "sparing" in Atlantic salmon smolts when fed a diet with a high DHA:EPA ratio [109].

The use of a 50% FO and 50% CF blend in aquafeeds for Atlantic salmon, as was in the range commercially practiced in Australia, resulted in comparable LC omega-3 content in the muscle [108] and liver of juvenile Atlantic salmon to a FO fed fish. It is noteworthy that such an oil blend decreases the inefficient utilization of a 100% FO diet, due to the high loss of EPA in particular, and can be considered as an appropriate current strategy, in terms of LC omega-3 sparing, for present use in aquafeeds for Atlantic salmon [108]. It is important to note that in spite of changes that have occurred in feeding practices, farmed Tasmanian Atlantic salmon still remains one of the best sources of omega-3 LC-PUFA oils available to Australian consumers. However, the scope can exist with the potential future use of new oilseed-derived LC omega-3 to restore the content of these health-benefitting ingredients, and also the omega-3/omega-6 ratio to that previously seen.

Further research is needed to determine the optimum relative and absolute concentrations of dietary EPA and DHA to enhance their deposition in larger-sized commercially farmed Atlantic salmon. The rationale to pursue such studies will be reliant on research in plant genomics since oils with the desired FA profiles, in particular containing a high DHA:EPA ratio [58,59], whilst not presently available, will likely be a commercial reality by the end of this decade.

### **10. TAG Structure of Plant-Based LC Omega-3 Oils for Optimum Bioactivity and Food Processing**

The melting characteristic of a fat/oil is an important determinant of its suitability for the manufacture of food products. For example, a certain melting range is required before a fat can be used for the manufacture of margarine or fat spreads. The positional distribution of omega-3 LC-PUFA within the TAG molecules can significantly influence the melting characteristics of LC omega-3 oils. In general, the melting point is increased when the omega-3 LC-PUFA is located at the *sn*-2 position compared with the *sn*-1(3) positions [110]. This has practical implications enabling the conversion of liquid oils to semi-solid fats for margarine manufacture or use as *trans* fat substitutes in bakery products. Furthermore, omega-3 LC-PUFA such as DHA are more resistant to oxidative deterioration when located at the *sn*-2 position compared to the *sn*-1(3) positions [111].

The effects of fatty acid positional distribution on absorption and nutrition of oils and fats are less well understood. Evidence from animal and human infant studies suggests that TAG structure affects digestibility, atherogenicity and fasting lipid levels, with fats containing palmitic and stearic acid in the *sn*-2 position being better digested and considered more harmful for cardiovascular health [112–114]. However, a few studies in human adults have indicated that fatty acid positional distribution has no effect on digestibility or fasting plasma lipids [115,116]. There have been very limited studies on the physiological effects of TAG positional distribution of omega-3 LC-PUFA such as EPA and DHA on either animals or humans. These fatty acids are predominantly located at the *sn*-2 position in fish oil TAG with the notable exception of seal blubber oil. Though it has been hypothesised, there are presently not sufficient data with humans to conclude that location of omega-3 LC-PUFA at the *sn-*2 position confers greater physiological benefit when compared to location at the *sn-*1 *or sn*-3 positions. We consider this as an area for further fruitful research including with animal model and clinical trials. The available evidence does not yet support a preference as to how the LC omega-3 containing TAG in novel oilseeds should be best assembled to maximize the nutritional benefits to human consumers. As we acquire this knowledge, the prospect of tailoring the DHA positional distribution of novel plant-based DHA oils (both during metabolic engineering and post-harvest) can be used to meet optimum health and food processing properties

#### **11. Conclusions**

The need for further clinical trials to better refine our understanding of the mode of action of the health-benefitting LC omega-3 oils will continue, including with emphasis towards the mode of action of the individual components namely EPA and DHA. Similar requirements exist for farmed species, in particular cultured fish. The latter are being increasingly fed non-marine oils, although limited research has occurred on the possible deleterious effects to the farmed species of the lower dietary proportions of LC omega-3 oils, accompanied by a lower omega-3/omega-6 ratio, and ultimately to human consumers. The issue of the finite supply of the fish oil resource is very clearly upon us, and new sustainable sources of these oils are required. SCO derived oils are in use, although remain, and likely will remain, relatively expensive and therefore better suited to niche applications. The past five years have seen expanded interest in applications with krill oil, although considerable care with exploitation of this environmentally sensitive and important resource must occur. After several decades of research for production of LC omega-3 oils from oil seeds, the prospects for such a supply are now a reality, including most recently the difficult to achieve yet nutritionally important DHA. LC omega-3 oils derived from GM oil seed crops may in the future provide the most economically viable source of these key essential ingredients for aquaculture and a range of other applications. It is estimated that the cost and availability of oils from GM plants would be similar to that of currently available commercial oilseed crops such as rapeseed and soya. Further research and development in this area has the potential for significant commercial, health, social and environmental benefits. This exciting field of research will now move into the commercial development phase, with feeding and other trials and associated approvals and consumer acceptance to occur. Other areas of research for continuing effort will include: improved processing and yields for pharmaceutical grade products, improvement and/or further development of novel delivery modes for application of LC omega-3 oils in functional foods, examination of the effects of omega-3 LC-PUFA positional distribution on the bioactivity and processing properties of various LC omega-3 food products, and also for the omega-3 index to gain increasing acceptance and use. Collectively research and development in these and other areas will ensure that enhanced intake of the LC omega-3 oils can occur for a wider range of consumers, with resultant global heath, economic and social benefits resulting.

#### **Acknowledgements**

We thank the wider CSIRO Food Futures Flagship Omega-3 research team for their contributions and Allan Green who also assisted in the preparation of Figure 1. Surinder Singh, James Petrie, Rick Phleger, Manny Noakes and Welma Stonehouse and three anonymous journal reviewers provided helpful review comments on the manuscript.

#### **Author Contributions**

All authors contributed to the review of the literature and the preparation of the manuscript.

#### **Conflicts of Interest**

The authors declare no conflict of interest.

#### **References**



## **Detailed Distribution of Lipids in Greenshell™ Mussel (***Perna canaliculus***)**

#### **Matthew R. Miller, Luke Pearce and Bodhi I. Bettjeman**

**Abstract:** Greenshell™ mussels (GSM–*Perna canaliculus*) are a source of omega-3 (*n-*3) long-chain polyunsaturated fatty acids (LC-PUFA). Farmed GSM are considered to be a sustainable source of LC-PUFA as they require no dietary inputs, gaining all of their oil by filter-feeding microorganisms from sea water. GSM oil is a high-value product, with a value as much as 1000 times that of fish oils. GSM oil has important health benefits, for example, anti-inflammatory activity. It also contains several minor lipid components that are not present in most fish oil products, and that have their own beneficial effects on human health. We have shown the lipid content of the female GSM (1.9 g/100 g ww) was significantly greater than that of the male (1.4 g/100 g ww). Compared with male GSM, female GSM contained more *n-*3 LC-PUFA, and stored a greater proportion of total lipid in the gonad and mantle. The higher lipid content in the female than the male GSM is most likely related to gamete production. This information will be useful to optimize extraction of oils from GSM, a local and sustainable source of *n-*3 LC-PUFA.

Reprinted from *Nutrients*. Cite as: Miller, M.R.; Pearce, L.; Bettjeman, B.I. Detailed Distribution of Lipids in Greenshell™ Mussel (*Perna canaliculus*). *Nutrients* **2014**, *6*, 1454–1474.

#### **1. Introduction**

The New Zealand Greenshell™ mussel (GSM), *Perna canaliculus*, is indigenous to the coastlines of New Zealand. GSM are distinguishable by the green coloration of the shell near the lip, which gives the mussel its name. Since the establishment of GSM aquaculture in New Zealand in 1969 [1], the GSM industry has seen consistent and substantial growth. In 2011, the export value of GSM was 218 million New Zealand dollars (increased from approximately 40 million NZ dollars in 1989) from a product volume of more than 38,000 t (~90,000 t at harvest) [2]. GSM are sold as food, and are also used to produce high-value nutraceuticals including oil extracts and freeze-dried mussel powders, for example, Lyprinol® and Seatone®. Studies have shown that lipid extracted from GSM has numerous health benefits, including the ability to reduce inflammation [3–13]. GSM lipid contains a high proportion of omega-3 long-chain (C ≥ 20) polyunsaturated fatty acids (*n-*3 LC-PUFA), predominantly docosahexaenoic acid (DHA, 22:6*n-*3) and eicosapentaenoic acid (EPA, 20:5*n-*3), which are split between the triacyglycerol and polar lipid classes [14]. There are also several minor lipid components in GSM oil, including non-methylene-interrupted (NMI)-FA, plasmalogen, phytosterols, and furan fatty acids [14,15]. These minor lipids are not present in most fish oil products, and some have been shown to have beneficial effects on human health [15–18]. Because of the low concentration of oil in GSM [14,19] and the expensive extraction techniques (supercritical CO2 with ethanol as a co-solvent, or chemical extraction) required to extract it, GSM oil is very expensive. The estimated price is NZ \$3000/kg.

There has been extensive research confirming the health benefits of increased consumption of *n-*3 LC-PUFA, especially DHA and EPA [20–23]. Consequently, there is an increasing market for *n-*3 marine oil supplements. The proportion of *n-*3 LC-PUFA is higher in GSM oil [14] than in certain fish oils, such as those extracted from sardine, anchovy, and cod liver. GSM obtain *n-*3 LC-PUFA from their diet, which is rich in zooplankton and phytoplankton [24,25]. GSM are considered to be one of the most sustainable sources of *n-*3 LC-PUFA as they are farmed, rather than wild harvested, and do not require any dietary inputs for their nutrition. The lipid content and FA profile of GSM have been analyzed previously [14,19,26,27]. Those studies reported that polar lipids (PL) were the major lipid class (42.5–61.7 g/100 g) in GSM, followed by triacylglycerides (TAG) (17.8–49.2 g/100 g) with the remainder made up of sterols (5.5–6.8 g/100 g), free fatty acids (FFA) (2.9–14.9 g/100 g), and trace amounts of wax esters [14,19]. Previous studies reported that the concentration of PUFA in GSM lipid ranged from 19 to 49.1 g/100 g, with 6–12 g/100 g DHA, and 8–24 g/100 g EPA [14,28]. The lipid content of GSM, and the lipid classes and FA profile of GSM oil, are affected by many factors, including the season, location, and the types and amounts of algae consumed. To date, there have been no reports on the FA profiles of lipid extracted from different organs of the GSM.

In this study, we investigated the lipid classes and fatty acid profile of GSM lipids, and analyzed the lipid content of male and female GSM. To analyze lipid localization within the GSM, we dissected the GSM body into five components: the mantle, gonad, heart and foot, posterior adductor muscle, and the digestive gland. A knowledge of the differences in the amounts and composition of lipids between genders, and of how physiological function affects GSM lipids, will be useful to optimize the lipid extraction process, which would benefit the GSM marine oil industry.

#### **2. Experimental Section**

#### *2*.*1*. *Sampling*

Whole live GSM were collected 12 November 2012 from the Marlborough Sounds (South Island, New Zealand) and stored in a recirculating sea water system until experiments began on 16 November 2012. After transport and dry storage, mussels were rehydrated for 24 h in 100 L bins supplied with 10 L/min filtered seawater on a flow-through basis with auxiliary aeration. The mussels were weighed, shucked, and then the mussel meat was weighed. 10 Female and 10 male mussels were selected for lipid extraction based on gonad color; gonads of males are creamy white while those of females are various shades of orange (Figure 1). The data for one mussel was excluded from the analysis because that individual was an outlier in terms of size and oil content. Six female and six male mussels were used for analyses of the various components of the GSM body. The shell and meat were weighed and then the meat was dissected into the mantle, gonad, posterior adductor muscle, heart/foot, and the digestive gland (Figure 1). The gonad was not able to be completely separated, and some gonad tissue remained attached to the digestive gland. Other un-dissectible smaller organs including the gill and labial palp were omitted from this analysis. All samples were stored at −80 °C prior to freeze drying before lipid extraction.

**Figure 1.** (**A**) Female Greenshell™ mussel (*Perna canaliculus*) with orange gonad; (**B**) Male Greenshell™ mussel with creamy white gonad. Dissected organs of a male Greenshell™ mussel: (**C**) mantle; (**D**) gonad; (**E**) digestive gland and digestive gland; (**F**) posterior adductor muscle; (**G**) heart and foot.

The mussels were assigned a condition index as follows:

condition index (CI) = dry meat weight/(whole weight-shell weight) × 100 [29] (1)

#### *2*.*2*. *Lipid Extraction*, *Fractionation*, *and Fatty Acid Analysis*

GSM oils were extracted using a modified Bligh and Dyer protocol [30]. A single phase extraction with CHCl3:MeOH:H2O (1:1:0.9, v/v/v) yielded the total lipid extract (TLE). Lipid classes were analyzed with an Iatroscan MK V thin*-*layer chromatography-flame ionization detector (TLC-FID) (Iatron Laboratories, Tokyo, Japan). Samples were spotted onto silica gel SIII Chromarods (5-μm particle size) and developed in a glass tank lined with pre-soaked filter paper. The solvent system used for lipid separation was hexane: diethyl ether: acetic acid (60:17:0.1, v/v/v). After development for 25 min, the Chromarods were oven*-*dried and analyzed immediately to minimize adsorption of atmospheric contaminants. Lipid classes were quantified using Azur v5.0 software (DATALYS, St Martin D'Heres, France). The FID was calibrated for each compound class using the following compounds: phosphatidylcholine; cholesterol; cholesteryl ester; oleic acid; hydrocarbon (squalene); wax ester (derived from fish oil); and triacylglycerol (TAG, derived from fish oil).

An aliquot of the TLE from each sample type was trans-methylated in methanol: chloroform: hydrochloric acid (10:1:1, v/v/v) for 1 h at 100 °C. After addition of water the mixture was extracted three times with hexane: chloroform (4:1, v/v) to obtain fatty acid methyl esters (FAME). Samples were completed to 1 mL with an internal injection standard (23:0 or 19:0 FAME) and analyzed by gas chromatography mass spectrometry (GC-MS). The analytical system consisted of a Shimadzu 2010 QP GC-MS equipped a Restek GTx silica capillary column (30 m × 0.25 mm i.d., 0.25 μm film thickness). Samples (1 μL) were injected via a splitless injector at 220 °C. The column temperature program was as follows: 60 °C at 0 min; 40 °C min<sup>−</sup><sup>1</sup> to 100 °C; then 10 °C min<sup>−</sup><sup>1</sup> to 170 °C; then 5 °C min<sup>−</sup><sup>1</sup> to 185 °C; 2 min hold; then 3 °C min<sup>−</sup><sup>1</sup> to 197 °C; then 0.5 °C min<sup>−</sup><sup>1</sup> to 199 °C; 1 min hold; then 5 °C min to 230 °C; 3 min hold; then 5 °C min<sup>−</sup><sup>1</sup> to 250 °C; 5 min hold. Helium was the carrier gas. GC results were typically repeatable to within ±5% of the area of each individual component in replicate analyses.

Different classes of lipids were separated using a solid phase extraction (SPE) cartridge containing 500 mg silica (Grace Pure, Grace Davison Discovery Sciences, North Shore City, New Zealand). The SPE column was pre-treated with chloroform, and then a TLE sample from whole mussel, mantle, gonad, or digestive gland samples was loaded onto the column. Lipids were eluted with 10 mL chloroform followed by 10 mL methanol [31]. The oil fractions were concentrated by rotary evaporation and added to vials for analysis. Success of elution was confirmed via thin*-*layer chromatography (TLC).

#### *2*.*3*. *Sterol Analysis*

Sterols were isolated by saponifying 200 μL TLE in 5% w/v KOH in methanol/water (8:2 v/v) at 60 °C for 3 h. The mixture was cooled to room temperature and then 1 mL water was added. The sterols were then extracted twice using 1 mL hexane/chloroform (4:1). The extracted total non*-*saponifiable neutral (TSN) lipid fraction was transferred to labeled vials for further analysis. The TSN lipid fraction was blown down under a nitrogen stream and then heated at 60 °C with 50 μL *N*,*O*-bis(trimethylsilyl)trifluoroacetamide (BSTFA) for 40 min. BSFTA was evaporated under nitrogen and the sample was made up to 2 mL with chloroform before GC-MS analysis. Samples were injected into the GC at an oven temperature of 100 °C, which was increased to 300 °C at a rate of 10 °C/min and then held for 30 min. The mobile phase carrier gas was He. The silylated sterols were identified by their retention times and characteristic peaks in the mass spectrum [32].

#### *2*.*4*. *Statistical Analysis*

Mean values and standard error are reported. Normality and homogeneity of variance were confirmed and mean values were compared by one-way analysis of variance (ANOVA). Multiple comparisons were conducted using the Tukey-Kramer HSD (honestly significant difference) test. Differences were considered significant at *p* ≤ 0.05. Statistical analyses were performed using GenStat 14th Edition software.

#### **3. Results**

#### *3.1. Lipid Content of Whole GSM*

The lipid content of whole female GSM (1.9 g/100 g ww; 9.3 g/100 g dw) was significantly (*p <* 0.01) higher than that of male GSM (1.4 g/100 g ww; 7.4 g/100 g dw). The average total meat mass per individual was 18.3 g ± 3.3 g for females and 19.87 g ± 2.2 g for males. The Condition Index (CI) was 11.25 ± 2.0 for females and 9.57 ± 1.6 for males. There was no statistically significant difference between the two genders in the CI, size, or weight.

The major lipid class in both genders of GSM was PL. There was a significantly (*p <* 0.03) higher concentration of PL in males (82.3 g/100 g TLE) than in females (77.8 g/100 g TLE) (Table 1). The concentration of TAG in females (19.4 g/100 g TLE) was significantly higher (*p <* 0.03) than that in males (13.2 g/100 g TLE). The minor lipid classes in males and females were sterols (2.8–2.9 g/100 g TLE) and trace amounts (0.1 g/100 g TLE) of free fatty acids (FFA).


**Table 1.** Absolute FA (wet weight) and proportion of lipid classes (g/100 g TLE) in female and male Greenshell™ mussel (GSM) oil.


Values are mean ± standard error (*n* = 9). ww; wet weight; Other SFA: Sum of 15:0, 17:0, 20:0, 22:0 and 24:0; Other MUFA: Sum of 16:1*n-*5, 18:1*n-*7trans, 18:1*n-*5, 22:*n-*11 and 22:1*n-*9; Other PUFA: Sum of 16:2*n-*6, 18:3*n-*6, 20:4*n-*6, 20:2*n-*6, 22:5*n-*6, 22:4*n-*6 and 22:2*n-*6; Other includes fatty aldehydes and 4,8,12 trimethyl tetradecanoic acid (4,8,12-TMTD); NMI, non-methylene interrupted; TAG; triacylglycerols; PL: polar lipid; ST: sterols, FFA; free fatty acids; tr: trace.

#### *3.3. FA Profiles of GSM*

The major FA class (in % FA) was PUFA (53.8–55.9 g/100 g TLE), the main component of which was *n-*3 LC-PUFA (42.4–45.6 g/100 g TLE). The *n-*3/*n-*6 ratio was 6.6 in male GSM and 8.3 in female GSM. The major fatty acids (>10 g/100 g TLE) in both male and female GSM were EPA, DHA, and palmitic acid (16:0 PA). There were small amounts of NMI FA, including 20:2 NMI FA (2.8–3.4 g/100 g TLE) and 22:2 NMI FA (1.0–1.9 g/100 g TLE). The absolute FA content of the male and female (in mg/g ww) are shown in Table 1. Compared with male GSM, female GSM showed significantly (*p <* 0.01) higher concentrations of alpha-linolenic acid (ALA, 18:3*n-*3), stearidonic acid (SDA 18:4*n-*3), EPA, docosapentaenoic acid (DPA(*n-*3), 22:5*n-*3), total *n-*3s, and total PUFA.

#### *3.4. Lipid Content and Lipid Classes in Different Organs*

We used SPE to separate the non*-*polar and polar lipid fraction of TLE from the whole male and female GSM and from the various organs (Figure 2). The success of chromatographic separation was confirmed by TLC-FID. The most lipid-rich organs were the gonad (male, 3.1 g lipid/100 g ww; female, 4.0 g lipid/100 g ww), digestive gland (male, 3.1 g lipid/100 g ww; female, 3.7 g lipid/100 g ww) and mantle (1.5 g lipid/100 g ww; female, 2.0 g lipid/100 g ww). There were lower concentrations of lipid in the heart/foot (male, 1.3 g lipid/100 g ww; female, 1.0 g lipid/100 g ww) and adductor muscle (male, 1.2 g lipid/100 g ww; female, 0.8 g lipid/100 g ww) (Table 2, Supplementary Table S1). In all organs, PL was the major lipid class in the TLE. There were higher concentrations of TAG in the

**Table 1.** *Cont.*

digestive gland (male, 28.8 g/100 g TLE; female, 33.5 g/100 g TLE) and gonad (male, 38.5 g/100 g TLE; female, 33.7 g/100 g TLE) than in the other organs. The lowest concentrations of TAG were in the heart/ foot and adductor muscle samples (<8 g/100 g TLE). The main lipid class in the TLE from those samples was PL (>80 g/100 g TLE).

**Figure 2.** Lipid distribution in female and male Greenshell™ mussels (GSM) organs. \* Gonad also contained gill and labial palp. Values are mean ± standard error (*n* = 9).

#### *3.5. FA Profiles of Lipids from Different Organs*

In both male and female GSM, there were no differences in the proportions of the major FA (those detected at more than 10 g/100 g TLE; *i*.*e*., EPA, DHA, and PA) among the five different organs. For the minor FA, the proportions of SDA and ALA in the TLE from the digestive gland were significantly (*p <* 0.01) greater in the female GSM than the male GSM.

#### *3*.*6*. *Sterols*

Saponification of the TLE produced a total non*-*saponifiable neutral (TSN) fraction, which contained mainly sterols (Table 3). GC-MS analysis was performed after silylation of the TSN fraction. In total, 18 sterols were identified in GSM, with cholesterol being the predominant sterol. There was a significantly (*p <* 0.01) higher proportion cholesterol (30.2% of total sterols) in male GSM than in female GSM (28.5% of total sterols). The second most abundant sterol was brassicasterol; the proportion of brassicasterol was significantly (*p <* 0.01) higher in female (23.4% of total sterols) than in male GSM (21.05% of total sterols). Other major sterols were 24-nordehydrocholesterol, occelasterol, trans-22-dehydrocholesterol, and 24-methlenecholesterol.

#### **4. Discussion**

#### *4*.*1*. *Male vs*. *Female GSM*

In this study the lipid content of female GSM was significantly (*p <* 0.01) higher than that of male GSM even though it was previously reported that female mussels contain lower levels of total lipids [33]. Also, there was a significantly (*p <* 0.01) higher concentration of the TAG lipid class in the female than in the male. Because female GSM contained larger amounts of lipid than did males, the total amount of FA was higher in females than in males. In particular, the amounts of FA in the *n-*3 LC-PUFA class (ALA, SDA, EPA, and DPA) were all significantly (*p <* 0.01) higher in female than in male GSM. Differences in the lipid profile between male and female GSM have not been published previously, although a recent report showed similar results for the mussel species *Mytilus galloprovincialis* [34]. In another study, the FA composition of male and female GSM from two locations (Marlborough Sounds and Stewart Island) was profiled over three seasons (winter, spring, summer) but the types of lipids, that is, the lipid classes, were not reported [26]. Further, it was reported that GSM contained higher levels of lipids in summer and autumn than in winter and spring, but that the highest *n-*3 content was in winter [28]. In our study, we analyzed GSM collected in late spring (November) of 2012. Our results showed differences in lipid content, lipid classes, and total FA content between male and female GSM. In future research, it would be interesting to analyze changes in lipid content, lipid class composition, and the FA profile of male and female GSM on a finer time scale.


**Table 2.** Concentrations of fatty acids in oil extracted from mantle, gonad, and digestive gland of Greenshell™ mussels.

**141** 




**Table 3.** Sterol content of female and male Greenshell™ mussel (GSM) oil extract.

Values are mean ± standard error (*n* = 9); Unknown sterols could not be identified from MS data.

There is significant biological investment by the GSM into the production/storage of lipids during gonad development during winter, before spawning [28]. In general, the amount of lipid increases in GSM before spawning, and decreases to its lowest level immediately after spawning as a result of lipid loss during reproduction. GSM can also spawn in autumn (March–April) or shed eggs throughout their life cycle [28]. These reproductive events could explain differences in FA and lipid accumulation throughout the seasons. The higher lipid content in the female GSM is likely associated with oogenesis, during which lipid globules and small quantities of glycogen accumulate in the eggs [35]. There is a large variation in the timing and duration of gonad development in GSM and other mollusks, and the timing of gametogenesis varies according to the season and location [35]. A better understanding of the reproductive cycle, and how it affects the amount of lipid in GSM, may allow GSM oil producers to optimize the harvest time to obtain higher lipid yields.

Little is known about FA biosynthesis in GSM. In general, most bivalve species gain the majority of FA from their diet, although biochemical modifications of some FA occur in some species. The idea of "you are what you eat" has led to the study of signature lipids that can help to shed light on the preypredator relationships in ecosystems. When analyzed using high-powered statistical models, signature FA profiles can reveal prey-predator relationships in a food web. GSM are filter feeders that consume a variety of phytoplankton and zooplankton [24,25]. Recently, studies on feeding blue mussels (*Mytilus edulis*) organic waste from an aquaculture facility showed that when they were fed on a nontraditional (fish waste) diet, they did not feed or grow, but instead showed significant decreases in growth and total lipid content [36,37]. Phytoplankton and zooplankton are high in *n-*3 LC-PUFA, which results in the FA profile of GSM oil being rich in EPA and DHA. The types of FA present in the food sources of GSM will depend on factors such as season, location, and temperature. The similarity in the relative proportions of FA (g/100 g TLE) in males and females (and even among organs) suggested that there was little adaption of FA from their shared diet by either gender. However, the larger amount of lipid in the female GSM suggests that it has a greater capacity than the male GSM to store, and possibly biosynthesize, FA, especially *n-*3 LC-PUFA.

The biosynthetic pathway for *n-*3 LC-PUFA is unknown in GSM. Vertebrates have a poor capacity to produce DHA from precursors via the Sprecher pathway [38]. However, there is some evidence that invertebrates, such as the nematode *Caenorhabditis elegans*, have different pathways to produce *n-*3 LC-PUFA [39]. The FA results confirmed that the major biosynthetic precursors in the *n-*3 LC-PUFA pathway are present in the FA profile of GSM. These biosynthetic precursors include ALA, SDA, EPA, and DPA (*n-*3), with DHA being the final product. All of these FA except for DHA were present in significantly (*p <* 0.01) higher proportions of total FA in the female than in the male GSM. The higher concentrations of ALA, SDA, EPA, and DPA (*n-*3) may indicate that the female GSM has a greater *n-*3 LC-PUFA biosynthetic capacity during gametogenesis. Furthermore, in the digestive gland, the proportions of ALA and SDA in the FA profile were significantly (*p <* 0.01) higher in female than in male GSM. This may indicate a higher rate of FA biosynthesis in the digestive gland before lipids are transported for storage in the gonad. However, these FA results could also indicate that female GSM have an enhanced capacity to filter, process, and store *n-*3 LC-PUFA. To date, there have been no published reports of feeding trials to clarify aspects of lipid metabolism and/or storage in GSM. Therefore, it remains a matter of speculation whether the higher concentrations of FA in female than in male GSM are a result of enhanced biosynthesis or greater storage capacity.

Minor bioactive FA present in the GSM profiles included NMI PUFA, 4,8,12 trimethyl tetradecanoic acid (4,8,12-TMTD), and fatty aldehydes, all of which have been reported previously [14,19]. NMI FA are present in various mollusks at concentrations of up to 20% of wet weight [16]. Mollusks can synthesize NMI, which are believed to have structural and functional roles in biological membranes [16], via biosynthetic pathways that are still not fully understood. Our results showed that there were differences in the amount of NMI between male and female GSM, and that there was significantly less NMI in the gonad than in the other organs. Fatty aldehydes and 4,8,12-TMTD were present in GSM, but at very low concentrations (<1 g/100 g TLE), and were not included in the FA profiles. The amounts of both 4,8,12-TMTD and fatty aldehydes were substantially lower than those detected in our previous analyses of GSM that were collected from a similar location but at a slightly earlier time of the year (August and September 2009) [14]. Both NMI and 4,8,12-TMTD are possible indicators of red algae and/or some zooplanktonic pteropods, which could have been directly or indirectly consumed [19]. However, in other mollusks such as the green abalone, *Haliotis fulgens*, there is some evidence that 20:2 and 22:2 NMI may be metabolic products of desaturation of LC-MUFA [40]. Therefore, their presence may indicate an endogenous biosynthetic capacity of GSM.

#### *4*.*2*. *Differences among Organs*

Although the gonad contained the highest concentrations of lipid, the digestive gland also contained high lipid concentrations, which may be indicative of lipid-rich stomach contents. The lipid extracted from the digestive gland was darker and stickier than that extracted from the other organs. The dark color may represent non*-*enzymatic browning during extraction; this reaction occurs between oxidized lipids and primary amine groups, and has been observed in PL emulsions [41]. The digestive gland will likely contain the highest concentrations of endogenous lipases and low pH which may lead to the oxidization of lipid because of the more extreme conditions. Further, the dark color could be due to pigment residues derived from algae from the GSM diet

Generally, different lipid classes perform different functions in biological systems. Neutral lipids (*i*.*e*., TAG) are used as energy storage, while PL are mainly components of structural and functional parts of the cell. The lipid content of bivalves is directly linked to the gametogenic cycle, as TAGs and PLs play important roles as structural components and energy reserves in the gonads and gametes and during embryonic development [35]. Previous studies have shown the accumulation of neutral lipid reserves in eggs for a number of bivalve species, although studies looking at lipid production in GSM are not as detailed with regards to the effects of the gametogenic cycle [26,28]. It has been suggested that the digestive gland plays an important role in storing metabolic reserves, such as lipids, for use in gametogenesis and during periods of stress [35,42]. As previously mentioned, the higher proportion of *n-*3 LC-PUFA biosynthetic precursors (ALA and SDA) in the GSM digestive gland indicates that this organ is the most likely site of FA biosynthesis, and that these FA may have structural and functional roles in gamete development. The GSM analyzed in this study were collected in single sampling during a period of gametic growth; therefore, we cannot comment on changes in lipid composition during the year.

The mantle was the largest of the organs analyzed in this study, and accounts for approximately 20% of the wet weight of the mussel (Figure 2). The mantle is an important store of lipids in GSM; our results showed that this organ contained 21.5% (male mussels) and 28.0% (female mussels) of the total lipid. The main lipid class in the mantle was PL (73.5 g/100 g TLE in females; 77.3 g/100 g TLE in males) because of the large amount of structural tissue in this organ. The mantle consists of vascular connective tissue and plays a role in directing particles to the gills and deflecting other materials [35]. Our results suggested that the mantle of GSM also functions in lipid storage. The proportion of neutral lipids, which are generally used for storage, was higher in the mantle (22.9 g/100 g TLE in females; 18.4 g/100 g TLE in males) than in organs such as the adductor muscle (<3.0 g/100 g of TLE), which have different functions.

The three major organs of the GSM that store and use lipids are the gonad, digestive gland, and mantle. Our results showed that the adductor muscle and the heart and foot contained lower concentrations of lipids (3.9 g/100 g of TLE for adductor muscle and 5.4 g/100 g of TLE for heat and foot). The lipid extracted from the adductor muscle and the heart and foot showed similar FA profiles to that of the mantle. In the TLE from both the adductor muscle and the heart and foot samples, the major lipid class was PL (83.1–91.78 g/100 g of TLE) with minor amounts of TAG (1.9–7.5 g/100 g of TLE) and sterols (4.7–8.1 g/100 g of TLE). Profiles are included in the Supplementary Files. In general, the differences in lipid classes among different organs and between genders were minimal, suggesting that the diet is the main source of these FA and has the greatest influence on FA composition.

#### *4*.*3*. *Sterols*

The sterols in marine invertebrates are generally derived from sterols in the diet. However, some unconventional sterols are more common in primitive invertebrates, while cholesterol is found in greater proportions in more complex organisms [43]. The sterol profile of GSM reported here (Table 3) is similar to that of the only other published GSM sterol profile [19]. In a previous study, GSM were sampled from a similar region (Marlborough Sounds) at a similar time of year (October, compared with our sampling time in early November). In that study, there were only minor differences in the sterol profile among samples collected from four different sites [19]. This finding suggested that there was little difference in the diets of GSM among sites.

Phytosterols have wide bioactivity in humans. In particular, they are considered to be effective in lowering cholesterol, and consequently may have a preventive role against vascular disease [18,44]. Further beneficial properties of phytosterols include cancer prevention [18,45]. Phytosterols and cholesterol are likely metabolized by internal and external microorganisms into other bioactive substances. Phytosterols cannot be synthesized by humans and, therefore, can only be obtained through the diet. However, GSM may have some capacity to biosynthesize cholesterol into different phytosterols depending on their stage of the life cycle, sexual maturity, and gender [46]. As they are lipophilic substances with positive health benefits, phytosterols have been added to margarines and spreads to create "functional" foods. The amount of phytosterols derived from land sources is increasing in the marine environment as a result of land base ingredients increased used in aquafeeds. Changes from marine to terrestrial sources of protein and oil for use in aquafeeds has resulted in increased amounts and different types of phytosterols in farmed Atlantic salmon [47]. Here, we report that GSM oil contained approximately 3% phytosterols, of which about one-third was cholesterol. Larger amounts of sterols (5.5%–6.9%) have been reported previously for GSM [19]. Phytosterols such as isofucosterol and occelasterol are not common in our diets as they are not typical in terrestrial food sources, but may have novel beneficial functionalities. Isofucosterol and occelasterol are most likely derived from marine algae. In future research, it would be interesting to investigate whether novel phytosterols, along with the high content of ω3 LC-PUFA in GSM, can provide increased protection against coronary heart disease in humans.

#### *4*.*4*. *Implications of Results for the GSM Industry*

The major implications of this work are for the GSM oil industry, which is growing in both value and volume. A detailed understanding of the location of *n-*3 LC-PUFA in GSM may allow novel processing techniques to be established. The firmer parts of the mussel (posterior adductor muscle, heart and foot) contained very low concentrations of oil and *n-*3 LC-PUFA, and removal of these organs would reduce the amount of material processed using expensive freeze-drying and super-critical oil extraction procedures. This could potentially improve yields and benefit producers. Further, development of all-female lines of mussels would give mussel oil producers a greater source of oil and *n-*3 LC-PUFA. We estimate that oil yields could increase by as much as 35%, *n-*3 yields by 27%, and EPA yields by 43% if all-female mussels were extracted at the appropriate time of year. Highest yields could be obtained if female GSM were harvested in the peak reproductive state, since the gonad is the major storage centre for lipid. Higher investment in reproduction by the female GSM gonad may be to supply ample nutrition to offspring/eggs. The FA data suggested that female GSM have a higher biosynthetic capacity than that of males, and that they also store more lipid than do males. Compared with male GSM, female GSM contained larger amounts of the *n-*3 LC-PUFA biosynthetic precursors SDA and ALA. If both genders obtain the same FA from the shared diet, the increase in these biosynthetic precursors provides further evidence of an enhanced biosynthetic capacity in female GSM.

#### **5. Conclusions**

The lipid profiles of GSM were evaluated on the basis of gender and anatomy. The lipid content of the female GSM (1.9 g/100 g ww) was significantly greater than that of the male (1.4 g/100 g ww). The major lipid class in both genders was PL. Compared with male GSM, female GSM contained more *n-*3 LC-PUFA, and stored a greater proportion of total lipid in the gonad and mantle. The higher lipid content in the female than the male GSM is most likely related to gamete production. The mantle and digestive gland were other important sites for lipid storage and/or function/production. Novel bioactives, such as NMI-FA, plasmalogens, and phytosterols were identified in GSM oil.

#### **Acknowledgments**

We thank Helen Cleaver for technical assistance. This research was supported by MBIE (NZ Ministry of Business, Innovation and Employment) under project C02X0806, "Engineered Marine Molecules" with additional funding from MBIE programme "Export Marine Products".

#### **Author Contributions**

Matthew R. Miller made substantial contributions to conception and design, interpretation of data and development of manuscript. Luke Pearce and Bodhi Bettjeman made substantial contributions to acquisition of data, analysis and interpretation of data and development of manuscript.

#### **Abbreviations**

ANOVA, 1-way analysis of variance ALA, alpha-linolenic acid BSTFA, *N*,*O*-bis(trimethylsilyl)trifluoroacetamide CI, Condition index DHA, docosahexaenoic acid DPA, docosapentaenoic acid EPA, eicosapentaenoic acid FA, fatty acid(s) FFA, free fatty acid(s) FALD, dimethylacetals of aliphatic aldehydes FAME, fatty acid(s) methyl ester GC-MS, gas chromatography mass spectrometry GSM, Greenshell™ mussel HSD, honestly significant difference LC, long chain (C ≥ 20) MUFA, monounsaturated fatty acid(s) NMI, non-methylene interrupted TSN, non*-*saponifiable neutral *n*-3, Omega 3 *n*-6, Omega 6

PUFA, polyunsaturated fatty acid(s) SDA, stearidonic acid SFA, saturated fatty acid(s) TAG, triacylglycerol TLE, total lipid extract TMTD, trimethyl tetradecanoic acid tr, trace amounts

#### **Supplementary Information**

**Table S1.** Fatty acid concentration (mg/g) of Greenshell™ mussels. Polar and neutral lipid fractions were separated by solid phase extraction (SPE).


Values are mean ± standard error (*n*=9). ww; wet weight; Other SFA: Sum of 15:0, 17:0, 20:0, 22:0 and 24:0; Other MUFA: Sum of 16:1*n-*5, 18:1*n-*7trans, 18:1*n-*5, 22:*n-*11, and 22:1*n-*9; Other PUFA: Sum of 16:2*n-*6, 18:3*n-*6, 20:4*n-*6, 20:2*n-*6, 22:5*n-*6, 22:4*n-*6, and 22:2*n-*6; Other includes fatty aldehydes and 4,8,12 trimethyl tetradecanoic acid (4,8,12-TMTD).

**148** 


**Table S2.** Fatty acid concentrations in oil extracted from Greenshell™ mussel (GSM) adductor muscle and heart and foot.

TLE: Total lipid extract; Values are mean ± standard error (*n* = 6); Other SFA: Sum of 15:0, 17:0, 20:0, 22:0, and 24:0; Other MUFA: Sum of 16:1*n-*5, 18:1*n-*7trans, 18:1*n-*5, 22:*n-*11, and 22:1*n-*9; Other PUFA: Sum of 16:2*n-*6, 18:3*n-*6, 20:4*n-*6, 20:2*n-*6, 22:5*n-*6, 22:4*n-*6, and 22:2*n-*6; Other includes fatty aldehydes and 4,8,12 trimethyl tetradecanoic acid (4,8,12-TMTD); TAG: triacylglycerols, PL: polar lipid; ST: sterols.

#### **Conflicts of Interest**

The authors declare no conflict of interest.

#### **References**


## **Evaluation of Bread Crumbs as a Potential Carbon Source for the Growth of Thraustochytrid Species for Oil and Omega-3 Production**

#### **Tamilselvi Thyagarajan, Munish Puri, Jitraporn Vongsvivut and Colin J. Barrow**

**Abstract:** The utilization of food waste by microorganisms to produce omega-3 fatty acids or biofuel is a potentially low cost method with positive environmental benefits. In the present study, the marine microorganisms *Thraustochytrium* sp. AH-2 and *Schizochytrium* sp. SR21 were used to evaluate the potential of breadcrumbs as an alternate carbon source for the production of lipids under static fermentation conditions. For the *Thraustochytrium* sp. AH-2, submerged liquid fermentation with 3% glucose produced 4.3 g/L of biomass and 44.16 mg/g of saturated fatty acids after seven days. Static fermentation with 0.5% and 1% breadcrumbs resulted in 2.5 and 4.7 g/L of biomass, and 42.4 and 33.6 mg/g of saturated fatty acids, respectively. Scanning electron microscopic (SEM) studies confirmed the growth of both strains on breadcrumbs. Attenuated total reflection Fourier transform infrared (ATR-FTIR) spectroscopy for both strains were consistent with the utilization of breadcrumbs for the production of unsaturated lipids, albeit at relatively low levels. The total lipid yield for static fermentation with bread crumbs was marginally lower than that of fermentation with glucose media, while the yield of unsaturated fatty acids was considerably lower, indicating that static fermentation may be more appropriate for the production of biodiesel than for the production of omega-3 rich oils in these strains.

Reprinted from *Nutrients*. Cite as: Thyagarajan, T.; Puri, M.; Vongsvivut, J.; Barrow, C.J. Evaluation of Bread Crumbs as a Potential Carbon Source for the Growth of Thraustochytrid Species for Oil and Omega-3 Production. *Nutrients* **2014**, *6*, 2104–2114.

#### **1. Introduction**

Thraustochytrids are marine protists that belong to Labyrinthulomycetes and were first reported by Sparrow in 1936. These microorganisms are epibiotic in nature and represent a diverse group of organisms living in marine and estuarine habitats throughout the world and exhibiting a saprotrophic mode of nutrition [1]. Due to their ability to produce a large amount of oil, including polyunsaturated fatty acids (PUFAs), research has focused on lowering their cost of production using low cost carbon and nitrogen sources, particularly for large-scale industrial fermentation [2]. Biomass produced through heterotrophic fermentation of thraustochytrids is a potentially sustainable approach to the production of PUFAs for food, feed and supplement applications and oil for biofuel applications [3].

Thraustochytrids can act as microbial cell factories for the production of omega-3 PUFAs, squalene, and other secondary metabolites such as carotenoids and sterols, along with enzymes and extracellular polysaccharides [2]. For commercial utility in the production of these materials, thraustochytrid growth should be low cost, particularly to compete in the food market as a replacement for fish oils. Because heterotrophic organisms require a carbon source for bioconversion into oil, they are more expensive to grow in terms of consumables than autotrophic organisms. Heterotrophic fermentation has some advantages over autotrophic fermentation, particularly the ability to use standard industrial fermentation equipment at scale and to get much higher cell density than can be achieved with autotrophic organisms, which often need complex and expensive equipment to enable light to reach the cells while preventing contamination [4,5]. The cost of glucose as a carbon source in the growth medium to produce biomass can account for up to 30% of the overall production cost and so commercial biofuel production using heterotrophic fermentation requires the use of low cost carbon sources such as glycerol or food waste [6,7]. Food wastes generated worldwide are about 1.3 billion tons [8]. Management of food wastes through landfill dumping is common and is environmentally problematic. Food in landfills can rot and release methane gas, which is a major contributor to carbon emissions worldwide [9].

Research on using microbial fermentation to convert food waste into value added products is limited [10]. *Schizochytrium mangrovei* and *Chlorella pyrenoidosa* have been grown using food waste obtained by fungal hydrolysis and were reported to produce biomass potentially useful as a feed supplement or for biodiesel production [6]. *Schizochytrium mangrovei* KF6 was also reported to utilize processed bread crust to produce docosahexaenoic acid (DHA) from shake flask fermentation at 200 rpm under fluorescent light for eight days [11]. Polyunsaturated fatty acids were produced by *Mortierella alpina* utilizing rice bran as the carbon source in a solid-state column reactor under static conditions. Static conditions, where there is no agitation during the growth phase, are often used in solid substrate fermentation, as opposed to submerged liquid fermentation which requires higher energy inputs for continuous shaking of the flask at constant speed [12]. Other inexpensive carbon sources derived from food wastes that were studied includes okara powder [13], residues from beer and potato processing [14], sweet sorghum juice [15], coconut water [16], marine aquaculture waste water [17] and crude glycerol [18].

The objective of the present study was to investigate the ability of some thraustochytrid strains to utilize bakery waste, specifically breadcrumbs (BC), as an alternate carbon source in the fermentation media to produce lipids under static conditions. Scanning electron microscopy (SEM) was used to investigate the growth pattern of these microorganisms during static fermentation, and attenuated total reflection Fourier transform infrared (ATR-FTIR) spectroscopic analysis was performed to observe the production of unsaturated fatty acids during fermentation. ATR-FTIR spectroscopy and SEM were shown to be useful for monitoring static fermentation.

#### **2. Experimental Section**

All chemicals including fatty acid methyl ester standards used in this study were procured from Sigma-Aldrich (Sydney, Australia) and Merck Chemicals (Victoria, Australia) and were of analytical grade.

#### *2.1. Preparation of Seed Culture*

Thraustochytrid strains used in the study were *Thraustochytrium* sp. AH-2 (ATCC® PRA-296™, Manassas, VA, USA) and *Schizochytrium* sp. SR21 (ATCC® MYA-1381™, Manassas, VA, USA) and these strains were procured from the American Type Culture Collection (ATCC, Manassas, VA, USA), and grown in liquid media containing 1 g yeast extract, 15 g peptone, 20 g glucose (1 g yeast extract, 1 g peptone, 5 g glucose for *Schizochytrium* sp. SR21) in 1 L of artificial sea water (ASW) at 70% strength, according to ATCC product information sheets. In brief, the cultures were grown at 20 °C with shaking speed of 120 rpm for 96 h. Seed culture of 5% (v/v) culture was used in subsequent submerged liquid glucose fermentation and static fermentation with breadcrumbs as an alternate carbon source.

#### *2.2. Submerged Liquid Fermentation with Glucose Medium*

Cultures were grown in 250 mL flasks with 50 mL of medium altered and adopted from Li *et al.* [19]. Flasks were kept at 20 °C with shaking speed of 120 rpm for 7 days with medium at a starting pH of 6.0 (prior to autoclaving) and the pH was not controlled over the fermentation period. Flasks were collected for the dry weight determination and fatty acid estimation, each for 24 h.

#### *2.3. Preparation of Bakery Waste Bread Crumbs (BC) for Static Fermentation*

Breadcrumbs (crude powder; non-uniform size) were purchased from a local bakery (Geelong, Australia) for evaluating their utilization under static fermentation. BC powder was used for carrying fermentation experiments. Static fermentation was carried with the same media as submerged fermentation but with BC (at 0.5% and 1%) substituted for glucose in the media. Flasks were kept under static conditions in an incubator at 20 °C for 7 days, with pH of medium adjusted at 6.0 prior to autoclaving but not adjusted during the fermentation phase. Flasks were inoculated with 5% (v/v) seed culture (Section 2.1) under aseptic conditions. Samples were collected for the cell dry weight and fatty acid estimation after 24 h. Freeze-dried BC were analyzed using an EuroEA elemental analyzer (Euro Vector, Milan, Italy) to determine the percentages of carbon and nitrogen content.

#### *2.4. Scanning Electron Microscopy (SEM)*

A small flake of freeze-dried cells was mounted onto carbon tape on an aluminum stub and air dried, after which 60 nm of gold was deposited on its surface using a sputter coater. The cells were examined under a scanning electron microscope (SEM Supra 55 VP, Zeiss, Berlin, Germany) at accelerating voltage 3–5 KV using secondary electron detector.

#### *2.5. Fatty Acid Extraction and Gas Chromatography (GC) Analysis*

Fatty acid extraction was performed as previously described with some modifications [20]. In brief, 10 mg of freeze-dried cells were used for lipid extraction. Fatty acids were extracted with a mixture containing a 2:1 ratio of chloroform to methanol and repeated 3 times. For trans-esterification, 1 mL toluene was added followed by addition of 200 μL of internal standard, methyl nonadecanoate (C19:0) and 200 μL of butylated hydroxytoluene (BHT). Acidic methanol (2 mL) was also added to the tube and kept for overnight incubation at 50 °C. Fatty acid methyl esters (FAMEs) were extracted into hexane. The hexane layer was removed and dried over sodium sulphate. FAMEs were concentrated using nitrogen gas prior to GC analysis [21]. The samples were analyzed using a GC-FID system (Agilent Technologies, 6890N, Santa Clara, CA, USA). The GC instrument was equipped with a capillary column (Suplecowax 10, 30 × 0.25 mm, 0.25 μm thickness). Helium was used as the carrier gas at a flow rate of 1.5 mL·min−1. The injector was maintained at 250 °C and a sample volume of 1 μL was injected. Fatty acids were identified by comparison to external standards (Sigma-Aldrich, Sydney, Australia). Peaks were quantified with Chemstation chromatography software (Agilent Technologies, Santa Clara, CA, USA) and corrected using theoretical relative FID response factors [22]. Samples are analyzed in duplicate and compared to external standards.

#### *2.6. Attenuated Total Reflection Fourier Transform Infrared (ATR-FTIR) Spectroscopic Analysis*

ATR-FTIR measurements of the freeze-dried samples were conducted using an Alpha FTIR spectrometer (Bruker Optik GmbH, Ettlingen, Germany) equipped with a deuterated triglycine sulfate (DTGS) detector and a single-reflection diamond ATR sampling module (Platinum ATR QuickSnap™, Ettlingen, Germany). The ATR-FTIR spectra were acquired at 4-cm−1 spectral resolution with 64 co-added scans within the 4000–400 cm−1 spectral region. Blackman-Harris 3-Term apodization, power-spectrum phase correction, and zero-filling factor of 2 were set as default acquisition parameters using OPUS 6.0 software suite (Bruker Optik GmbH, Ettlingen, Germany). Background spectra of a clean ATR surface were acquired prior to each sample measurement using the same acquisition parameters.

#### *2.7. Statistical Analysis*

Data were statistically compared using one-way analysis of variance (Anova), and the significant difference was identified using Tukey's and Scheffe tests. The analysis was carried out using SPSS software (IBM® SPSS® Statistics 20, Sydney, Australia).

#### **3. Results and Discussion**

*Thraustochytrium* sp. AH-2 was previously isolated from coastal and mangrove habitats of Goa and further studied for its extracellular alkaline lipase production [23]. In the present study, control fermentation profiles were obtained using submerged liquid fermentation with 3% glucose as the carbon source. Under glucose conditions, biomass of 4.3 g/L and total lipid yield of 941 mg/L were achieved (Table 1). The biomass and lipid yield for our species is similar to that reported for *T. aureum* ATCC 34304 [24], although optimization of controlled fermentation conditions would probably enable higher biomass and lipid production for strain AH-2. Oleic acid (C18:1n9) was the major fatty acid at 63.19 mg/g, followed by palmitic acid (C16:0) 32.33 mg/g, DHA (C22:6n3) 23.74 mg/g and stearic acid (C18:0) 11.82 mg/g. C14:0, C15:0. C16:1n7, C17:0, C17:1n7 were present in lower amounts. DHA (C22:6n3) was the major PUFA, followed by docosapentaenoic acid (DPA) (C22:5n6) at 4.32 mg/g and eicosapentaenoic acid (EPA) (C20:5n3) at 3.03 mg/g.


**Table 1.** Fermentation profiles for *Thraustochytrium* sp. AH-2.

1 Palmitic acid (C16:0); stearic acid (C18:0); <sup>2</sup> Oleic acid (C18:1n9).

#### *3.1. Fermentation Growth Using Bread Crumbs as the Carbon Source*

Elemental analysis of the freeze-dried BC revealed approximately 40.95% carbon and 3.23% nitrogen. The fatty acid analysis of unfermented BC is presented in Table 2 and shows oleic acid as the major fatty acid in the profile, followed by palmitic acid (C16:0), stearic acid (C18:0), linoleic acid (18:2n6) and other fatty acids were also present. The polyunsaturated fatty acids EPA, DPA and DHA were not detected in the fatty acid profile of BC.

*Thraustochytrium* sp. AH-2 was grown using BC as the carbon source and the results compared with those obtained using glucose. BC is known to contain primarily complex carbohydrate in the form of starch [25]. BC was used at levels of 0.5% and 1%. Higher levels of 3% and 5% BC gave no improvement in cell growth and made lipid extraction difficult and were not pursued further. Fermentation with 0.5% BC gave 2.5 g/L of biomass and 260 mg/L of total lipid yield. Fermentation with 1% BC gave 4.7 g/L of biomass and 390 mg/L of total lipid yield (Figure 1). Compared to liquid fermentation with glucose, the total lipid yield was relatively low for static fermentation.


**Table 2.** Fatty acid profile <sup>1</sup> (mg/g) of unfermented bread crumbs, submerged liquid fermentation and static fermentation with bread crumbs, for *Thraustochytrium* sp. AH-2.

1 Palmitic acid (C16:0); stearic acid (C18:0); oleic acid (C18:1n9); linoleic acid (C18:2n6); linolenic (C18:3n3); EPA (C20:5n3); DPA (C22:5n3); DHA (C22:6n3); <sup>2</sup> Submerged liquid fermentation media with 3% glucose incubated for 7 days at 20 °C with shaking speed of 120 rpm; <sup>3</sup> Static fermentation with same medium composition as submerged liquid fermentation, with breadcrumbs as 5 and 10 g substituted for glucose in 1litre of 70% ASW at pH 6; <sup>4</sup> Static fermentation with 0.5% BC, incubated at 20 °C for 7 days; <sup>5</sup> Static fermentation with 1% BC, incubated at 20 °C for 5 days; a,b indicates statistically significant difference for submerged liquid fermentation (<sup>a</sup> ) and static fermentation (<sup>b</sup> ) (*p* < 0.05).

Under static fermentation, unsaturated fatty acids were poorly synthesized, although the presence of C18:3n3 (linolenic acid), EPA and DHA were clearly observed. Previous study on *Schizochytrium mangrovei* KF6 under heterotrophic conditions with processed bread crust also reported low levels of unsaturated fatty acids [11]. In general, monosaccharide sugars in the media result in the production of higher levels of PUFAs compared to di- and poly-saccharides [26]. As BC contains mainly starch, the observed poor synthesis of unsaturated fatty acids is not unexpected. In this study, the maximum fatty acid profile with 1% BC was observed on day 5, after which fatty acid content decreased, probably due to lipid consumption by the organism. Lipids and fatty acids accumulated in oleaginous microorganisms can act as energy sources for growth that are utilized when there is a lack of available carbon in the media [27].

The total amount of saturated fatty acids, which was primarily C16:0 and C18:0, were 42.4 mg/g with 0.5% BC and 33.6 mg/g with 1% BC, whilst submerged liquid fermentation with glucose as the carbon source gave an only slightly higher level of saturated fatty acids at 44.1 mg/g. Since PUFA production is low and unsaturated production is relatively high, fermentation with BC provides a fatty acid profile more consistent with that of biofuel, than did submerged liquid fermentation. Static fermentation may be a useful method for converting BC to oil, since parameters are readily standardized at industrial scale.

#### *3.2. Scanning Electron Microscopy (SEM) Observation of Cell Growth*

The fermentation growth for *Thraustochytrium* sp. AH-2 and *Schizochytrium* sp. SR21 were compared for BC and glucose as the carbon source using SEM. The morphology of freeze-dried unfermented BC was observed using SEM as a control material. The SEM images of cells grown using submerged liquid fermentation with glucose show spherical cells that are clumped together (Figure 2a). When grown in the presence of BC, cell clusters are attached to the BCs, confirming that cells do grow on this complex carbon source (Figure 2b,c).

**Figure 2.** SEM images of freeze-dried cells for (**a**) freeze-dried cells of *Schizochytrium* sp. SR21 grown under submerged liquid fermentation; (**b**) *Schizochytrium* sp. SR21 cells grown with 1% BC as alternate carbon source; (**c**) *Thraustochytrium* sp. AH-2 cells grown with 1% BC as alternate carbon source; (**d**) *Thraustochytrium* sp. AH-2 grown under submerged liquid fermentation.

*3.3. ATR-FTIR Spectroscopy Analysis of Cell Lipid Content* 

ATR-FTIR spectroscopic measurement of *Schizochytrium* sp. SR21 was performed to confirm the production of unsaturated fatty acids when BC was used as an alternative carbon source, in comparison to the submerged liquid fermentation with glucose of the same strain. Figure 3 shows the comparison of the ATR-FTIR spectral features of the raw unfermented BC, the static fermented 1% BC and glucose fermented cells. In particular, the olefinic C=CH stretching vibration found at ~3014 cm<sup>−</sup><sup>1</sup> is commonly known as a representative band for unsaturated fatty acids [28,29]. This band is clearly observed in the freeze-dried cells that were grown in 1% concentration of BC under static fermentation and with glucose, suggesting that observable amounts of unsaturated fatty acids were produced in the cells grown by both fermentation methods. The triplet bands found in the range of 3000–2800 cm<sup>−</sup><sup>1</sup> , on the other hand, are attributed to C-H stretches of lipids and proteins [29]. At the low wavenumber region, the strong bands centered at 1650 and 1545 cm−1, known as amide I and II bands, respectively, occurred due to the protein moieties in the BC and the cells. The sharp band at 1725 cm<sup>−</sup><sup>1</sup> , on the other hand, represents ν(C=O) stretches of ester functional groups from lipids and fatty acids, and is therefore indicative of total lipids produced by the cells [28–30]. According to the intensities of this band, fermentation with glucose led to a substantially higher amount of total lipids

produced in the microorganisms. However, the ratios of unsaturated fatty acids per total lipids (*i.e.*, *I*3014/*I*1725) were found to be comparable between both fermentation approaches, suggesting that similar yields of unsaturated fatty acid can be achieved using BC as a carbon source under static fermentation of *Schizochytrium* sp. SR21. Therefore, growth of these strains on BC is potentially useful both for the utilization of food waste and the production of lipid.

**Figure 3.** ATR-FTIR spectra of: (**a**) *Thraustochytrium* sp. AH-2; and (**b**) *Schizochytrium* sp. SR21. Black line—submerged fermentation; Blue line—unfermented breadcrumbs; Red line—static fermentation with breadcrumbs.

#### **4. Conclusions**

*Thraustochytrium* sp. AH-2 and *Schizochytrium* sp. SR21 were tested for their ability to utilize BC during static fermentation as a low-cost and environmental friendly carbon source for producing oil for either biofuel (saturated fatty acid rich) or food (PUFA rich). The fatty acid profiles from *Thraustochytrium* sp. AH-2 indicated low levels of PUFA and higher levels of saturated oil. ATR-FTIR spectroscopy of *Schizochytrium* sp. SR21 was also consistent with higher levels of saturated fatty acids. Fermentation on BC containing complex carbohydrate appears to be more appropriate to the production of biofuel from these organisms than for the production of high levels of PUFA for food applications, due to the suppression of PUFA when BC is used as the carbon source.

#### **Acknowledgments**

We thank the Centre for Chemistry and Biotechnology for funding towards this project, including some scholarship costs for T. Thyagarajan.

#### **Author Contributions**

T. Thyagarajan performed the research and contributed to writing the manuscript. J. Vongsvivut performed and interpreted the ATR-FTIR. M. Puri contributed to research and supervision. C. Barrow supervised the project and contributed to writing the manuscript.

#### **Conflicts of Interest**

The authors declare no conflict of interest.

#### **References**

