![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
Institute of Aquaculture, University of Stirling, Stirling, FK9 4LA, Scotland, UK and * BioMar Limited, Grangemouth, FK3 8UL, Scotland, UK
2To whom correspondence should be addressed. E-mail: gjb1{at}stir.ac.uk.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
KEY WORDS: • Atlantic salmon • rapeseed • canola • polyunsaturated fatty acid
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
2
3
4
5)
.
However, global capture fisheries are a finite resource. As a result of
overfishing and subsequently tighter regulation, future demand for
wild-caught fisheries products will exceed supply (6)
.
Fish produced by aquaculture currently represent the fastest growing
food sector, which is projected to increase at 
10%/y for at least
the next 10 y (7)
. Aquaculture has traditionally
utilized products from industrial fisheries, namely, fish meal and oil,
to convert relatively cheap protein and oil into high value products, a
practice that is sound both scientifically and commercially. However,
of the global fish oil production in 1996 of 1.4 million metric tonnes
(mt), 576 kilo tonnes (kt) were used for salmon and trout production
with the predicted use of fish oil for aquaculture estimated to rise to
85% of the total available by 2010 (8
,9)
. In addition,
competition for fish oil for inclusion in human nutritional supplements
and agricultural feeds other than for aquaculture will soon make fish
oil a highly prized commodity. Clearly, if aquaculture is to continue
to expand, alternatives to its current dependence on fish oil must be
developed.
Many freshwater fish, including salmonids, can convert 18:3(n-3) to
20:5(n-3) and then to 22:6(n-3) (10
11
12)
; by the same
enzymatic pathways of desaturation and elongation, they can convert
18:2(n-6) to 20:4(n-6). In fresh water, Atlantic salmon parr consume
substantial quantities of invertebrates that contain an abundance of
18:2(n-6) and 18:3(n-3) with lower amounts of 20:5(n-3) and almost no
22:6(n-3) (13)
. It is not surprising, therefore, that
salmon parr have been grown successfully to smoltification (sea water
transfer) with diets containing rapeseed and linseed oil
(14
,15)
. After smoltification, salmon enter the marine
environment in which their natural zooplanktonic crustacean and piscine
prey are rich in 20:5(n-3) and 22:6(n-3). Therefore, it might be
expected that the genes encoding the desaturase and elongase enzymes
responsible for the conversion of 18:3(n-3) to 22:6(n-3) are
downregulated, possibly irreversibly, soon after salmon migrate to sea.
However, a number of studies have suggested that salmonids can utilize
vegetable oils in seawater provided the diets contain enough 18:3(n-3)
to satisfy essential fatty acid (EFA) requirements
(16
17
18)
. More recent evidence suggests that there is no
"switch off" of fatty acid–metabolizing enzymes in salmon
post-smolts and that fish fed vegetable oils showed increased
conversion of 18:3(n-3) to 22:6(n-3) [and 18:2(n-6) to 20:4(n-6)]
compared with those fed fish oil (FO) (19)
.
At the present time, the product of the salmon aquaculture industry is
of high nutritional quality with an abundance of (n-3) HUFA and a high
(n-3)/(n-6) polunsaturated fatty acid (PUFA) ratio (20)
.
Clearly, producers and consumers of salmon will want to minimize any
perceived reduction in quality arising from the inclusion of vegetable
oils, both in terms of growth and health of the fish and in the healthy
image of salmon as part of the human diet. The high growth rates of
cultured salmon are due to the use of high energy (oil) diets, which
are rich in the C-20 and C-22 monoenoic fatty acids characteristic of
northern hemisphere marine oils. Thus, any substitute for fish oil in
salmon feeds should meet the following criteria: 1) avoid
excessive deposition of 18:2(n-6); 2) enhance conversion of
18:3(n-3) to 20:5(n-3) and 22:6(n-3); and 3) provide
sufficient energy in the form of monoenoic fatty acids to maintain high
growth rates. In this regard, rapeseed oil (RO) is a potential
candidate in that it has moderate levels of 18:2(n-6) and 18:3(n-3), in
a ratio of 2:1, and an abundance of 18:1(n-9). The ratio of
18:2(n-6)/18:3(n-3) in rapeseed oil makes it of benefit to human
health, as well as fish health (21)
. In this study,
duplicate groups of Atlantic salmon post-smolts were fed diets
containing RO at 10, 25, 50 and 100% of the added oil, in comparison
to FO (0% RO), for 17 wk. Growth and feed utilization were measured
along with muscle and liver total lipid and fatty acid composition,
muscle proximate composition, muscle carotenoid content [astaxanthin
(Ax)] and hepatocyte fatty acid desaturation and elongation
activities.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Atlantic salmon S1 smolts (n = 350; initial mean
weight, 80 g) were obtained from the F.R.S. Marine Research Unit,
Aultbea, Wester Ross, Scotland. The smolts were randomly distributed
into 10 1-m diameter tanks of 500-L capacity supplied with
nonrecirculated sea water at 10 L/min at the F.R.S. Marine Research
Unit. The tanks were subjected to a photoperiod regime of 12 h
light:12 h dark and the temperature over the experimental period
(August–December 1999) ranged from 7.9 to 14.2°C with an average
temperature of 11.7 ± 1.5°C. The diets were supplied by
automatic feeders, which were activated for a few seconds every 15 min
during daylight hours and adjusted to provide 20 g/kg biomass each day.
Fish were weighed individually at the start and finish of the
experiment and bulk weighed every 28 d, with the ration adjusted
accordingly. An initial sample was taken at the start of the trial,
after 9 wk, and final sampling was performed after 17 wk of the feeding
experiment. The experiment was conducted in accordance with the British
Home Office guidelines regarding research on experimental animals.
Five practical-type commercial extruded diets were formulated
(BioMar, Brande, Denmark) to provide 47% crude protein and 25% lipid
and differed only in their content of rapeseed oil which was added at
0, 10, 25, 50 and 100% of the total added oil, the remainder of which
was FO (Table 1). The diets were designed to satisfy the nutritional requirements of
salmonid fish (22)
. All diets supplied sufficient (n-3)
PUFA to meet requirements for normal growth and development. In diets
with high levels of RO, the (n-3) PUFA requirement was met by PUFA
derived from dietary fish meal. The proximate composition of the
experimental diets including total carotenoid and Ax contents is shown
in Table 2
and the dietary fatty acid compositions are shown in Table 3
.
|
|
|
An initial sample of six fish was taken at the start of the experiment to determine baseline levels of lipid and fatty acid composition in liver and white muscle, and carotenoid content in muscle. A similar sample of 12 fish per dietary treatment was taken after 9 wk. After 17 wk, each analytical measurement was performed on at least 10 fish, selected at random, from each tank. Fish were killed by a blow to the head, and samples of liver were dissected and immediately placed in liquid nitrogen. A muscle sample, representative of the edible portion, was obtained by cutting a steak from the "Norwegian Quality Cut" region, between the dorsal and ventral fins. This section was then skinned, deboned, the red muscle trimmed off and the remaining white muscle homogenized, after removal of the dorsal fat body, in a Waring Blender (Fisher Scientific, Loughbourgh, UK). The homogenate was frozen in liquid nitrogen and all samples were stored at -40°C before analysis.
Proximate analysis.
The nutrient composition of the five experimental diets and muscle
samples was determined by proximate analysis. Moisture was determined
by thermal drying to constant weight in an oven at 110°C for 24 h. Sample weight was recorded before drying and after removal from the
oven, after cooling in a desiccator. Moisture was expressed as a
percentage of wet weight. Crude protein was determined by combustion
using the Dumas process (23)
. Crude lipid in diets was
determined by acid hydrolysis followed by Soxhlet extraction
(24)
. Ash content (g/100 g dry weight) was determined by
dry ashing in porcelain crucibles in a muffle furnace at 600°C
overnight (24)
.
Lipid extraction and fatty acid analysis.
Total lipid was extracted from liver and muscle by homogenizing in 20
volumes of chloroform/methanol (2:1, v/v) in an Ultra-Turrax tissue
disrupter (Fisher Scientific) and measured gravimetrically according to
the method of Folch et al. (25)
. Fatty acid methyl esters
(FAME) were prepared by acid-catalyzed transesterification of total
lipid according to the method of Christie (26)
. Extraction
and purification of FAME were performed as described by Ghioni et al.
(27)
. FAME were separated and quantified by gas-liquid
chromatography (Carlo Erba Vega 6000, Milan, Italy) using a 30 m
x 0.32 mm i.d. capillary column (CP Wax 52CB, Chrompak, London, UK).
Hydrogen was used as carrier gas and temperature programming was from
50 to 150°C at 40°C/min and then to 230°C at 2.0°C/min.
Individual methyl esters were identified by comparison with known
standards and by reference to published data (28)
. The gas
chromatography method used does not separate 22:1(n-13) from 22:1(n-11)
although in the text these combined fatty acids are referred to as
22:1.
Ax measurement.
Total carotenoid was extracted from salmon muscle largely by the method
of Barua et al. (29)
. Tissue samples were homogenized in 5
mL of absolute ethanol and 5 mL of ethyl acetate using an
Ultra-Turrax tissue disrupter (Fisher Scientific). The homogenate
was centrifuged (1000 x g, 5 min) and the
supernatant removed to a stoppered glass tube. The pellet was
rehomogenized in 5 mL of ethyl acetate and recentrifuged, and the
supernatant was combined with the first supernatant. Finally, the
pellet was rehomogenized in 10 mL of hexane and recentrifuged, and the
supernatant combined with the pooled supernatant. The pooled
supernatant was dried under a stream of nitrogen and
vacuum-desiccated for 2 h before redissolving the residue in 2
mL of hexane containing 0.2g/L BHT. Total carotenoid was measured
spectrophotometrically at 470 nm using the
E1% (wt/v) of 2100. Measurement of Ax was
carried out using a 5-µm Luna ODS2 column (4.6 x 150 mm, Phenomenex, Macclesfield, UK). The chromatographic system was
equipped with a Waters Model 501 pump and Ax was detected at 470 nm
using a Waters 490E multiwavelength UV/vis detector [Millipore (UK),
Watford, UK]. An isocratic solvent system was used containing ethyl
acetate/methanol/water (20:72:8, v/v/v) at a flow rate of 1 mL/min. Ax
was detected at 470 nm and quantified using an external standard of Ax
obtained from Roche (Heanor, UK). Ax in diets was extracted and
measured similarly after an enzymatic digestion of the ground extruded
pellets with Maxatase (International Biosynthetics, Rijswijk,
Netherlands). Portions of ground diet (1 g) were mixed with 10 mL water
and 10 mg of Maxatase in a 50-mL stoppered glass tube followed by
incubation in a water bath at 50°C for 30 min.
Preparation of isolated hepatocytes.
The gall bladder and main blood vessels were carefully dissected from the liver. The liver was perfused via the hepatic vein with solution A [calcium- and magnesium-free Hank’s balanced salt solution (HBSS) + 10 mmol/L HEPES + 1 mmol/L EDTA] to clear blood from the tissue. The liver was chopped finely with scissors and incubated with 20 mL of solution A containing 1 g/L collagenase in a 25 mL "Reacti-flask" in a shaking water bath at 20°C for 45 min. The digested liver was filtered through 100-µm nylon gauze and the cells collected by centrifugation at 1000 x g for 5 min. The cell pellet was washed with 20 mL of solution A containing fatty acid–free bovine serum albumin (FAF-BSA, 10 g/L) and recentrifuged. The hepatocytes were resuspended in 10 mL of Medium 199 containing 10 mmol/L HEPES, 2 mmol/L glutamine, 1 x 105 U/L penicillin and 0.1 g/L streptomycin. Cell suspensions (100 µL) were mixed with 400 µL of trypan blue; hepatocytes were counted and their viability assessed using a hemocytometer. Cell suspensions (100 µL) were also retained for protein determination as described below.
Assay of hepatocyte fatty acyl desaturation/elongation activities.
Fatty acyl desaturation and elongation activities were determined in
isolated hepatocytes as described by Buzzi (30)
.
Hepatocyte suspensions (5 mL) were dispensed into two
25-cm2 tissue culture flasks. Hepatocytes were incubated
with 9.25 KBq of [1-14C] 18:3(n-3) added as a complex
with FAF-BSA. Briefly, 925 KBq of fatty acid (0.5
µmol) in ethanol was placed in a reaction vial,
solvent evaporated under a stream of nitrogen and 100
µL of 0.1mol/L KOH added. The mixture was stirred for
10 min at room temperature before the addition of 5 mL of 50 g/L
FAF-BSA in HBSS containing 10 mmol/L HEPES buffer; the mixture was
stirred for 45 min at 20°C. After isotope addition, the flasks were
incubated at 20°C for 2 h. After incubation, the cell layer was
dislodged by gentle rocking, the cell suspension transferred to glass
conical test tubes and the flasks washed with 1 mL of ice-cold HBSS
containing 10 g/L FAF-BSA. The cell suspensions were centrifuged at
300 x g for 2 min, the supernatants discarded and the cell
pellets washed with 5 mL of ice-cold HBSS/FAF-BSA and
recentrifuged. The supernatants were again discarded and the tubes
placed upside down on paper towels to blot for 15 s before
extraction of total lipid using ice-cold chloroform/methanol (2:1,
v/v) containing 0.1 g/L BHT essentially according to Folch et al.
(25)
and as described in detail previously
(31)
.
Total lipid was transmethylated and FAME prepared as described by
Tocher et al. (31)
. The methyl esters were redissolved in
100 µL hexane containing 0.1 g/L BHT and applied as
2.5-cm streaks to TLC plates impregnated by spraying with 2 g
silver nitrate in 20 mL acetonitrile and preactivated at 110°C for 30
min. Plates were fully developed in toluene/acetonitrile (95:5, v/v)
(32)
. Autoradiography was performed with Kodak MR2 film
(Sigma, Poole, UK) for 4–7 d at room temperature. Silica corresponding
to individual PUFA was scraped into scintillation mini-vials
containing 2.5 mL of scintillation fluid (Ecoscint A. National
Diagnostics, Atlanta, GA) and radioactivity determined in a
TRI-CARB 2000CA scintillation counter (United Technologies Packard,
Pangbourne, UK). Results were corrected for counting efficiency and
quenching of 14C under exactly these conditions.
Protein determination.
Protein concentration in hepatocytes was determined according to the
method of Lowry et al. (33)
after incubation with 0.25 mL
of 2.5 g/L SDS, 1 mol/L NaOH for 45 min at 60°C.
Histology.
Samples of liver, heart, kidney and white muscle were fixed in 200 mL/L buffered formol saline and embedded in paraffin wax. Sections (5 µm) were cut and stained with hematoxylin and eosin for histological analysis. Pathological assessment was carried out on coded, randomized slides to eliminate bias in interpretation. Examination of slides was performed using a Reichert (Microstar IV) compound microscope at an objective magnification of X40 or X100. All organs were examined for general abnormalities such as tumors, infections, inflammation, loss or replacement of tissues.
Statistical analysis.
Significance of difference (P < 0.05) between dietary treatments was determined by one-way ANOVA. Differences between means were determined by Tukey’s test. Data identified as nonhomogeneous, using Bartlett’s test, were subjected to log or arcsin square-root transformation before applying the ANOVA. ANOVA was performed using a Graphpad Prism (version 2.0) statistical package (Graphpad Software, San Diego, CA).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The proximate composition of white muscle (Table 4) showed no significant differences in moisture content, but lipid
content was significantly greater, albeit only modestly, in fish fed
0% RO (FO) compared with those fed 50% RO. Conversely, the protein
content of muscle was significantly lower, again only modestly so, in
fish fed 0% RO compared with all other treatments. Ash content was
greatest in fish fed 10 and 50% RO followed by 0 and 25% RO and
lowest in fish fed 100% RO. Liver lipid content (Table 4)
was
significantly higher in fish fed 100% RO compared to fed fish fed 0,
10 and 25% RO. There were no significant differences in total
carotenoid pigment or Ax deposition in muscle in fish fed the different
experimental diets.
|
. Plots of concentrations of individual fatty acids in flesh total
lipids (g fatty acid/100 g total fatty acids) (Table 5)
against their
concentration in dietary lipid (Table 3)
yielded linear relationships;
typical examples are shown in Figure 1
. Correlation coefficients and slopes of the lines generated for
individual fatty acids are shown in Table 6
. Thus, the concentrations of individual fatty acids in dietary lipid
were linearly related to their concentrations in flesh total lipid.
However, the slopes and intercepts of the linear plots in Figure 1
and
Table 6
differed for different fatty acids, i.e., the relationship
between the concentration in the diet and the concentration in the
flesh differed for different fatty acids. This is seen more clearly in
Table 6
from the differences (
values) between the concentration of
individual fatty acids in dietary lipid and flesh total lipid for fish
fed the diets containing 0% RO (100% FO) and 100% RO. Thus,
22:6(n-3) and 18:1(n-9) were present in higher concentrations in flesh
lipid than in the 0% RO (100% FO) dietary lipid, whereas 20:5(n-3)
and 22:1 were present in lower concentrations (Table 6)
. Conversely,
18:1(n-9) and also 18:2(n-6) and 18:3(n-3), but not 22:1, were present
in lower concentrations in flesh lipid than in the 100% RO dietary
lipid, whereas 22:6(n-3) was higher in flesh lipid than in the dietary
lipid. Clearly, 22:6(n-3) was preferentially deposited in flesh lipid
relative to its level in dietary lipid with all dietary lipid blends
tested. However, 18:1(n-9), ,18:2(n-6) and 18:3(n-3) were
preferentially discriminated against in flesh relative to the diet at
high but not at low dietary lipid concentrations, and the same was true
for 22:1. This reflected the inverse concentrations of these fatty
acids in the 100% RO and 0% RO (100% FO) dietary lipids.
|
|
|
and correlation coefficients, slopes and values are shown in
Table 7
. The effects of the experimental diets on fatty acid compositions of
liver were qualitatively the same as described for muscle. However, a
number of notable quantitative differences occurred, particularly with
respect to monounsaturated fatty acids and HUFA. Thus, the
concentrations of monounsaturated fatty acids, especially 20:1(n-9) and
22:1 (results not shown), were much lower in liver than in flesh
lipids, whereas 20:5(n-3) and 22:6(n-3) were both notably greater in
liver lipids compared with flesh (Figs. 1
and 2)
. Moreover, both of
these PUFA and especially 22:6(n-3) were present in higher
concentrations in liver lipid than in dietary lipid.
|
|
and 4
). A linear relationship (r = 0.99) was found
between the recovery of total (summed) desaturation products of
[1-14C] 18:3(n-3) and the concentration of
18:3(n-3) in the diet (Fig. 3)
. The effect of diet on individual
desaturation and elongation products of [1-14C]
18:3(n-3) is shown in Figure 4
. The product of 6-desaturation, 18:4(n-3), increased significantly,
in stepwise fashion, in hepatocytes from fish fed diets containing 25,
50 and 100% RO, compared with fish fed 0% RO. The product of
6-desaturation and elongation, 20:4(n-3), was the most abundant
metabolite of radiolabeled 18:3(n-3) recovered and increased
significantly with increasing dietary RO inclusion in a manner similar
to 18:4(n-3). Although only very low quantities of 22:6(n-3) were
produced compared with other products, the highest quantities were
produced in fish fed 50 and 100% RO followed by 25% RO, with the
lowest levels in fish fed 10 and 0% RO.
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
,18
,34)
, and Atlantic salmon in
seawater (16)
showed no reduction in growth rate and FER
with dietary lipid contents of 14–19%. The recent study of Torstensen
et al. (35)
showed no differences in growth or feed
conversion when Atlantic salmon were fed diets containing 35% lipid
composed principally of capelin oil, palm oil and oleic acid–enriched
sunflower oil.
The percentage of lipid in the muscle was significantly greater in fish
fed 0% RO compared with fish fed 50% RO although the differences
between the dietary treatments were small. Conversely, the muscle
protein content was significantly lower in fish fed 0% RO compared
with all other treatments, and the second highest muscle lipid level,
found in fish fed 100% RO, was correlated with the second lowest
protein content. This relationship between muscle protein and lipid
content has been observed in previous studies with salmonids
(36
,37)
. Variation in the type and quantity of dietary
PUFA can influence uptake and deposition of carotenoid pigments in
salmon flesh (38)
. In the present study, there were no
significant differences in total muscle carotenoid and Ax
concentrations among the five dietary treatments. Flesh pigmentation is
an important factor in perception of flesh quality in salmonids
(20
,39)
. A previous study observed reduced pigment
deposition in salmon when fish oil was replaced with soybean oil or
beef tallow (40)
.
The fatty acid compositions of tissue lipids of Atlantic salmon are
readily influenced by the fatty acid composition of dietary lipid
(35
,41)
. This is amply confirmed in the present study,
which clearly establishes linear correlations, with different slopes
and intercepts, between the percentage of individual fatty acids in
dietary lipids and in muscle total lipid, consisting of 85%
triacylglycerols (TAG) and 10% phospholipids (PL), and in liver total
lipid, consisting of 30% TAG and 50% PL (20
,42)
.
Therefore, although the fatty acid composition of the total lipid is a
combination of the TAG and PL compositions, the former predominate in
salmon muscle and are thereby the primary influence on the composition
of total muscle lipid. In this study, we chose to concentrate on total
lipid rather than consider the different attributes of TAG and PL
because total lipid reflects the fatty acids available to the consumers
of fish. The correlations described above are of practical use for
predicting outcomes of feeding different blends of a given substituting
oil with fish oil. They also reveal how different fatty acids in
dietary lipid are selected for or against relative to tissue lipids.
Thus, the data in Tables 5
and 6
and Figure 1
establish that 22:6(n-3)
is selectively deposited in muscle lipids. Possible mechanisms
underlying this selective deposition include high specificity of fatty
acyl transferases for 22:6(n-3) and/or relative resistance of 22:6(n-3)
to ß-oxidation stemming from the complex catabolic pathway for this
fatty acid. It is of interest that complete replacement of fish oil in
the diet with rapeseed oil reduced the percentage of 22:6(n-3) in
dietary lipid by more than fourfold but in muscle lipid by only
twofold. This residual dietary 22:6(n-3) reflects the presence of fish
meal in the diet, and its associated 22:6(n-3)–rich PL and TAG.
In contrast to 22:6(n-3), the monoenes 22:1 and 18:1(n-9) are
discriminated against in muscle lipids relative to dietary lipids when
present at high concentrations, most clearly seen in the 0 and 100% RO
diets, respectively. This is readily accounted for by the ease of
ß-oxidation of these monoenes (43
,44)
. The high oil
diets used currently in salmon culture and in this study are of course
designed to generate energy through extensive fatty acid oxidation,
thereby sparing amino acid oxidation and enhancing the growth rate of
the fish. It is interesting that when 22:1 and 18:1(n-9) are present in
dietary lipid at similar percentages, as in the 0% RO diet, 22:1
appears to be catabolized in preference to 18:1(n-9) (Table 6)
.
Nevertheless, the data here are consistent with18:1(n-9) being
readily oxidized at high concentrations, as in the 100% RO diet, as is
the case also for 18:2(n-6) and 18:3(n-3).
The foregoing considerations for muscle total lipid hold also for liver
total lipid, except that the trends in liver lipid are quantitatively
greater than in muscle lipid. This reflects the predominance in liver
of PL, rich in 20:5(n-3) and especially 22:6(n-3), containing a modest
percentage of 18:2(n-6) and 18:3(n-3) and negligible 22:1, and whose
composition is not so readily influenced by dietary input as muscle
lipid, which is rich in TAG (20
,42)
.
Increases in hepatic desaturation and elongation activities have been
observed in previous studies in which salmon were fed vegetable oils or
vegetable oil blends (14
,15
,19)
. In the present
study, hepatic desaturation and elongation of 18:3(n-3) were clearly
enhanced progressively by replacing dietary FO with RO. However, only
very small amounts of end product 22:6(n-3) were formed in the short
term radiolabeling assay using hepatocytes; the major product was the
intermediate 20:4(n-3). This intermediate was not detected in large
amounts in fatty acid analyses of hepatic lipids, nor was there any
alteration in the ratio of 20:5(n-3) to 22:6(n-3) in hepatic lipids.
This suggests that although substituting FO in the diet with RO clearly
activates hepatic desaturation of 18:3(n-3) to 18:4(n-3) and its
further elongation to 20:4(n-3), the reactions may proceed too slowly
to influence significantly the dietary-induced changes in fatty
acid compositions of tissue lipids.
Salmon production worldwide in 1996 was 0.64 million mt and production
for 2000 is expected to be 0.8 million mt with a 1 million mt
harvest predicted 2002. With global fish oil supplies static, or
even in decline, the expansion of aquaculture production in
general, and salmon production in particular, will occur only if other
oil sources are utilized. A number of studies have investigated the
suitability of different vegetable oils as replacements for fish
oil in a range of fish species including, rainbow trout
(Oncorhynchus mykiss) (45
,46)
, brown trout
(Salmo trutta) (47)
, arctic charr
(Salvelinus alpinus) (48)
, brook charr
(Salvelinus fontinalis) (18)
, Atlantic salmon
(16
,17
,40)
and chinook salmon (Oncohynchus
tshawytscha) (34)
. A notable recent study is that of
Torstensen et al. (35)
investigating the effects of
capelin oil, palm oil and oleic acid–enriched sunflower as dietary
lipid sources for Atlantic salmon. Only the last-mentioned study
used an oil-rich diet formulation similar to the high energy feeds
currently favored for salmonid culture.
Given present concerns about the imbalance of (n-6) and (n-3) PUFA in
the diets of developed nations and the encouragement to consume oily
fish, such as mackerel, sardines, salmon and trout, it is important
that cultured salmon maintain a high level of essential (n-3) HUFA in
the edible flesh (3
4
5)
. Therefore, successful
substitution of fish oil with vegetable oils should minimize
depletion of 20:5(n-3) and 22:6(n-3), minimize deposition of 18:2(n-6)
and maximize desaturation and elongation of 18:3(n-3) to 20:5(n-3) and
22:6(n-3) while permitting growth rates equivalent to those presently
achieved with fish oils. The present study has investigated the
potential of rapeseed oil, fed at different inclusion levels, to
satisfy these criteria in Atlantic salmon post-smolts in sea water.
In our previous studies with Atlantic salmon fed vegetable oils,
including corn, sunflower, grape seed and safflower oils, there was a
significant increase in concentrations of arachidonic acid
[20:4(n-6)] in fish tissues, resulting in increased eicosanoid
production and the appearance of cardiac histopathologies (17
, 49
50
51)
. All of the oils mentioned above contain high levels of
18:2(n-6) and only trace levels of 18:3(n-3). Elevated percentages of
20:4(n-6) were not observed in tissue lipids in the present study, nor
were cardiac histopathologies observed, consistent with negligible
conversion of 18:2(n-6) to 20:4(n-6) resulting from replacement of
dietary FO with RO. The presence of substantial levels of 18:3(n-3) in
rapeseed oil, despite an excess of 18:2(n-6), can account for the lack
of a significant conversion of 18:2(n-6) to 20:4(n-6), due to
competitive interactions between (n-3) and (n-6) PUFA in hepatic
desaturation and elongation pathways (52
53
54)
. However,
fully understanding how simultaneously varying dietary levels of
precursor 18:2(n-6) and 18:3(n-3), and end product 20:4(n-6), 20:5(n-3)
and 22:6(n-3) affects hepatic elongation-desaturation pathways
requires further study. Nonetheless, the finding here that substituting
dietary FO with RO activates the pathways at least in part offers hope
that conditions can be identified whereby at least significant
conversion of 18:3(n-3) in the substituting oil to 22:6(n-3) occurs to
offset in part the decrease in dietary 22:6(n-3) caused by reduction of
dietary FO.
In summary, the present study suggests that rapeseed oil is an effective substitute for fish oil in Atlantic salmon in terms of permitting similar growth rates and feed efficiency, and having no apparent ill effects on fish health. However, inclusion of RO at levels in excess of 50% of supplementary lipid results in significant decreases in the (n-3)/(n-6) PUFA ratio and the EPA and DHA concentrations in fish flesh such that the nutritional benefits of the fish to the human consumer would be considerably reduced. This does not preclude the use of higher levels of rapeseed oil in dietary formulations for Atlantic salmon because such diets could be used for the majority of the growth cycle providing, at an appropriate time before marketing, the fish were returned to a fish oil-containing diet thereby restoring the 18:2(n-6), EPA and DHA concentrations to their "normal" values. These, and other aspects of fish oil substitution, are the subject of ongoing research activity in our laboratory.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
3 Present address; North Atlantic Fisheries College, Port Arthur, Scalloway, Shetland ZE1 0UN, Scotland, UK.
4 Abbreviations used: Ax, astaxanthin; BSA, bovine serum albumin; DHA, docosahexaenoic acid; EFA, essential fatty acid; EPA, eicosapentaenoic acid; FAF, fatty acid–free; FAME, fatty acid methyl esters; FER, feed efficiency ratio; FO, fish oil; HBSS, Hank’s balanced salt solution; HUFA, highly unsaturated fatty acids; kt, kilo tonnes; mt, metric tonnes; PL, phospholipid; PUFA, polyunsaturated fatty acids; RO, rapeseed oil; SGR, specific growth rate; TAG, triacylglycerol.
Manuscript received October 30, 2000. Initial review completed December 4, 2000. Revision accepted February 14, 2001.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
1. British Nutrition Foundation Task Force Unsaturated Fatty Acids. Nutritional and Physiological Significance. The Report of the British Nutritional Foundation Task Force 1992 Chapman & Hall London, UK.
2. British Nutrition Foundation Nutritional Aspects of Fish. British Nutrition Foundation Briefing Paper No. 10 1993 British Nutrition Foundation London, UK.
3. International Society for the Study of Fatty Acids and Lipids Health message statement. International Society for the Study of Fatty Acids and Lipids News Letter 1994;1:3-4
4. International Society for the Study of Fatty Acids and Lipids Recommendations for the essential fatty acid requirements for infant formulae. International Society for the Study of Fatty Acids and Lipids News Letter 1994;1:4-5
5.
Krauss R. M., Eckel R. H., Howard B., Appel L. J., Daniels S. R., Deckelbaum R.J ., Erdman J. W., Krisetherton P., Goldberg I. J., Kotchen T. A. AHA dietary guidelines—revision 2000: a statement for healthcare professionals from the nutrition committee of the American Heart Association. Circulation 2000;102:2284-2299
6. Sargent J. R., Tacon A.G.J. Development of farmed fish: a nutritionally necessary alternative to meat. Proc. Nutr. Soc. 1999;58:377-383[Medline]
7. Hempel E. Constraints and possibilities for developing aquaculture. Aquac. Int. 1993;1:2-19
8. Pike I. Future role of fish meal and oil in salmon diets. Salmon News 1996;4/5:3-5
9. Anonymous World fish meal production plunging to 15 year low. Oil World 1998;41:108
10. Castell J. D., Lee D. J., Sinnhuber R. O. Essential fatty acids in the diet of rainbow trout: lipid metabolism and fatty acid composition. J. Nutr. 1972;102:93-99
11. Sargent J. R., Bell J. G., Bell M. V., Henderson R. J., Tocher D. R. Requirement criteria for essential fatty acids. J. Appl. Ichthyol. 1995;11:183-198
12. Buzzi M., Henderson R. J., Sargent J. R. Biosynthesis of docosahexaenoic acid proceeds via 24-carbon intermediates. Comp. Biochem. Physiol. 1997;116B:263-267
13. Bell J. G., Ghioni C., Sargent J. R. Fatty acid compositions of ten freshwater invertebrates which are natural food organisms of Atlantic salmon parr (Salmo salar); a comparison with commercial diets. Aquaculture 1994;128:301-313
14. Bell J. G., Tocher D. R., Farndale B. M., Cox D. I., McKinney R. W., Sargent J. R. The effect of dietary lipid on polyunsaturated fatty acid metabolism in Atlantic salmon (Salmo salar) undergoing parr-smolt transformation. Lipids 1997;32:515-525[Medline]
15. Tocher D. R., Bell J. G., Dick J. R., Henderson R. J., McGhee F., Mitchell D. F., Morris P. C. Polyunsaturated fatty acid metabolism in Atlantic salmon (Salmo salar) undergoing parr-smolt transformation and the effects of dietary linseed and rapeseed oils. Fish Physiol. Biochem. 2000;23:59-73
16. Polvi S. M., Ackman R. G. Atlantic salmon (Salmo salar) muscle lipids and their response to alternative fatty acid sources. J. Agric. Food Chem. 1992;40:1001-1007
17. Bell J. G., Dick J. R., McVicar A. H., Sargent J. R., Thompson K. D. Dietary sunflower, linseed and fish oils affect phospholipid fatty acid composition, development of cardiac lesions, phospholipase activity and eicosanoid production in Atlantic salmon, Salmo salar. Prostaglandins Leukot. Essent. Fatty Acids 1993;49:665-673[Medline]
18. Guillou A., Soucy P., Khalil M., Adambounou L. Effects of dietary vegetable and marine lipid on growth, muscle fatty acid composition and organoleptic quality of flesh of brook charr (Salvelinus fontinalis). Aquaculture 1995;136:351-362
19.
Tocher D. R., Bell J. G., Dick J. R., Sargent J. R. Fatty acyl desaturation in isolated hepatocytes from Atlantic salmon (Salmo salar): stimulation by dietary borage oil containing 
-linolenic acid. Lipids 1997;32:1237-1247[Medline]
20. Bell J. G., McEvoy J., Webster J. L., McGhee F., Millar R. M., Sargent J. R. Flesh lipid and carotenoid composition of Scottish farmed Atlantic salmon (Salmo salar). J. Agric. Food Chem. 1998;46:119-127[Medline]
21. Ackman R. G. Canola fatty acids—an ideal mixture for health, nutrition and food use. Shahidi F. eds. Canola and Rapeseed 1990 Avi New York, NY.
22. National Research Council Nutrient Requirements of Fish 1993 National Academy Press Washington, DC.
23. Association of Official Analytical Chemists 15th ed. Official Methods of Analysis 1990;1:71-72 AOAC Washington, DC.
24. Woyewoda A. D., Shaw S. J., Ke P. J., Burns B. G. Recommended Laboratory Methods for Assessment of Fish Quality. Canadian Technical Report of Fisheries and Aquatic Sciences No. 1448 1986 NRC Canada. 156 pages
25.
Folch J., Lees M., Sloane-Stanley G. H. A simple method for the isolation and purification of total lipids from animal tissues. J. Biol. Chem. 1957;226:497-509
26. Christie W. W. Lipid Analyses 2nd ed. 1982:52-56 Pergamon Press Oxford, England.
27. Ghioni C., Bell J. G., Sargent J. R. Polyunsaturated fatty acids in neutral lipids and phospholipids of some freshwater insects. Comp. Biochem. Physiol. 1996;114B:161-170
28. Ackman R. G. Fish lipids, part 1. Connell J. J. eds. Advances in Fish Sciences and Technology 1980:86-103 Fishing News Books Farnham, UK.
29. Barua A. B., Kostic D., Olson J. A. New simplified procedures for the extraction and simultaneous high-performance liquid chromatographic analysis of retinol, tocopherols and carotenoids in human serum. J. Chromatogr. 1993;617:257-264[Medline]
30. Buzzi M. The Characterization of Docosahexaenoic Acid [22:6(n-3)] Biosynthesis in the Liver of Rainbow Trout (Oncorhynchus mykiss) 1996 University of Stirling Scotland, UK. Doctoral thesis,
31. Tocher D. R., Sargent J. R., Freriches G. N. The fatty acid compositions of established fish cell lines after long-term culture in mammalian sera. Fish Physiol. Biochem. 1988;5:219-227
32. Wilson R., Sargent J. R. High-resolution separation of polyunsaturated fatty acids by argentation thin-layer chromatography. J. Chromatogr. 1992;623:403-407
33.
Lowry O. H., Roseborough N. J., Farr A. L., Randall R. J. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 1951;193:265-275
34. Dosanjh B. S., Higgs D. A., Plotnikoff M. D., Markert J. R., Buckley J. T. Preliminary evaluation of canola oil, pork lard and marine lipid singly and in combination as supplemental dietary lipid sources for juvenile fall chinook salmon (Oncorhynchus tshawytscha). Aquaculture 1988;68:325-343
35. Torstensen B. E., Lie O., Froyland L. Lipid metabolism and tissue composition in Atlantic salmon (Salmo salar L.)—effects of capelin oil, palm oil and oleic acid-enriched sunflower oil as dietary lipid sources. Lipids 2000;35:653-664[Medline]
36. Reinitz G., Hitzel F. Formulation of practical diets for rainbow trout based on desired performance and body composition. Aquaculture 1980;19:243-252
37. Jokumsen A., Alsted N. Ørredvaekst I relation til forholdet mellem protein og fedt I foderet. Medd. Forsøgsdambruget 1990;80:1-15
38. Christiansen R., Waagbø R., Torissen O. J. Effects of polyunsaturated fatty acids and vitamin E on flesh pigmentation in Atlantic salmon, Salmo salar. Kaushik S. J. Luquet P. eds. Fish Nutrition in Practice 1993:339-343 INRA Editions Paris, France. Proceedings of the 4th International Symposium on Fish Nutrition and Feeding, Biarritz, France
39. Refsgaard H.H.F., Brockhoff P. B., Jensen B. Biological variation of lipid constituents and distribution of tocopherols and astaxanthin in farmed Atlantic salmon (Salmo salar). J. Agric. Food Chem. 1998;46:808-812
40. Hardy R. W., Scott T. M., Harrell L. W. Replacement of herring oil with menhaden oil, soybean oil, or tallow in the diets of Atlantic salmon raised in marine net-pens. Aquaculture 1987;65:267-277
41. Lie Ø., Waagbø R., Sandnes K. Growth and chemical composition of adult Atlantic salmon (Salmo salar) fed dry silage based diets. Aquaculture 1988;69:343-353
42. Henderson R. J., Tocher D. R. The lipid composition and biochemistry of freshwater fish. Prog. Lipid Res. 1987;26:281-347[Medline]
43. Henderson R. J., Sargent J. R. Chain length specificities of mitochondrial and peroxisomal ß-oxidation of fatty acids in rainbow trout (Salmo gairdneri). Comp. Biochem. Physiol. 1985;82B:79-85
44. Kiessling K.-H., Keissling A. Selective utilisation of fatty acids in rainbow trout (Oncorynchus mykiss Walbaum) red muscle mitochondria. Can. J. Zool. 1993;71:248-251
45. Cho C. Y., Bayley H. S., Slinger S. J. Partial replacement of herring meal with soybean meal and other changes in a diet for rainbow trout (Salmo gairdneri). J. Fish. Res. Board Can. 1974;31:1523-1528
46. Greene D.H.S., Selivonchick D. P. Effects of dietary vegetable, animal and marine lipids on muscle lipid and hematology of rainbow trout (Oncorhynchus mykiss). Aquaculture 1990;89:165-182
47. Arzel J., Martinez-Lopez F. X., Metallier R., Stephan G., Viau M., Gandemer G., Guillaume J. Effect of dietary lipid on growth performance and body composition of brown trout (Salmo trutta) raised in seawater. Aquaculture 1994;123:361-375
48. Olsen R. E., Henderson R. J. Muscle fatty acid composition and oxidative stress indices in Arctic charr, Salvelinus alpinus (L.), in relation to dietary polyunsaturated fatty acid levels and temperature. Aquac. Nutr. 1997;3:227-238
49. Bell J. G., Youngson A., Mitchell A. I., Cowey C. B. The effect of enhanced intake of linoleic acid on the fatty acid composition of tissue polar lipids of post-smolt Atlantic salmon (Salmo salar). Lipids 1989;24:240-242
50. Bell J. G., McVicar A. H., Park M. T., Sargent J. R. High dietary linoleic acid affects the fatty acid compositions of individual phospholipids from tissues of Atlantic salmon (Salmo salar): association with stress susceptibility and cardiac lesion. J. Nutr. 1991;121:1163-1172
51. Bell J. G., Dick J. R., Sargent J. R., McVicar A. H. Dietary linoleic acid affects phospholipid fatty acid composition in heart and eicosanoid production by cardiomyocytes from Atlantic salmon (Salmo salar). Comp. Biochem. Physiol. 1992;103A:337-342
52. Brenner R. R. Nutritional and hormonal factors influencing desaturation of essential fatty acids. Prog. Lipid Res. 1981;20:41-47[Medline]
53. Christiansen E. N., Lund J. S., Rørtveit T., Rustan A. C. Effect of dietary n-3 and n-6 fatty acids on fatty acid desaturation in rat liver. Biochim. Biophys. Acta 1991;1082:57-62[Medline]
54. Horrobin D. F. Interactions between n-3 and n-6 essential fatty acids (EFAs) in the regulation of cardiovascular disorders and inflammation. Prostaglandins Leukot. Essent. Fatty Acids 1991;44:127-131[Medline]
This article has been cited by other articles:
|
|
 |
|
  M. R. Miller, A. R. Bridle, P. D. Nichols, and C. G. Carter Increased Elongase and Desaturase Gene Expression with Stearidonic Acid Enriched Diet Does Not Enhance Long-Chain (n-3) Content of Seawater Atlantic Salmon (Salmo salar L.) J. Nutr., November 1, 2008; 138(11): 2179 - 2185. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. Menoyo, C. J. Lopez-Bote, A. Obach, and J. M. Bautista Effect of dietary fish oil substitution with linseed oil on the performance, tissue fatty acid profile, metabolism, and oxidative stability of Atlantic salmon J Anim Sci, December 1, 2005; 83(12): 2853 - 2862. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. G. Bell, D. R. Tocher, R. J. Henderson, J. R. Dick, and V. O. Crampton Altered Fatty Acid Compositions in Atlantic Salmon (Salmo salar) Fed Diets Containing Linseed and Rapeseed Oils Can Be Partially Restored by a Subsequent Fish Oil Finishing Diet J. Nutr., September 1, 2003; 133(9): 2793 - 2801. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. G. Bell, R. J. Henderson, D. R. Tocher, F. McGhee, J. R. Dick, A. Porter, R. P. Smullen, and J. R. Sargent Substituting Fish Oil with Crude Palm Oil in the Diet of Atlantic Salmon (Salmo salar) Affects Muscle Fatty Acid Composition and Hepatic Fatty Acid Metabolism J. Nutr., February 1, 2002; 132(2): 222 - 230. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
