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        Dietary carbohydrate and lipid sources affect differently the oxidative status of European sea bass (Dicentrarchus labrax) juveniles
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        Dietary carbohydrate and lipid sources affect differently the oxidative status of European sea bass (Dicentrarchus labrax) juveniles
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        Dietary carbohydrate and lipid sources affect differently the oxidative status of European sea bass (Dicentrarchus labrax) juveniles
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Abstract

This study aimed to evaluate the effects of dietary lipid source and carbohydrate content on the oxidative status of European sea bass (Dicentrarchus labrax) juveniles. For that purpose, four diets were formulated with fish oil (FO) and vegetable oils (VO) as the lipid source and with 20 or 0 % gelatinised starch as the carbohydrate source, in a 2×2 factorial design. Liver and intestine antioxidant enzyme activities (catalase (CAT), superoxide dismutase (SOD), glutathione peroxidase (GPX), glutathione reductase (GR), glucose-6-phosphate dehydrogenase (G6PD)), hepatic and intestinal lipid peroxidation (LPO), as well as hepatic oxidative stress index (OSI), were measured in fish fed the experimental diets for 73 d (n 9 fish/diet). Carbohydrate-rich diets promoted a decrease in hepatic LPO and OSI, whereas the lipid source induced no changes. Inversely, dietary lipid source, but not dietary carbohydrate concentration, affected LPO in the intestine. Lower intestinal LPO was observed in VO groups. Enzymes responsive to dietary treatments were GR, G6PD and CAT in the liver and GR and GPX in the intestine. Dietary carbohydrate induced GR and G6PD activities and depressed CAT activity in the liver. GPX and GR activities were increased in the intestine of fish fed VO diets. Overall, effects of diet composition on oxidative status were tissue-related: the liver and intestine were strongly responsive to dietary carbohydrates and lipid sources, respectively. Furthermore, different metabolic routes were more active to deal with the oxidative stress in the two organs studied.

Substitution of fishmeal (FM) and fish oil (FO) in aquafeeds by more readily available and sustainable alternative ingredients such as plant feedstuffs (PF) has become a priority issue in aquaculture( 1 ). However, dietary replacement of fishery by-products by PF may have negative health-related effects, especially in carnivorous species. For instance, vegetable oils (VO) were reported to induce alterations of the immune response in different fish species( 2 7 ), change the fatty acid profile of immune cells or modify eicosanoid production( 2 , 5 , 6 , 8 ). Inflammation and pathological morphological modifications in the intestine and steatosis in the liver were also reported in several species fed VO-rich diets( 9 13 ). Inadequate dietary nutrient balance, high fibre or carbohydrate content and the presence of anti-nutritional factors in PF are possible causes for such effects( 14 , 15 ).

Oxidative stress occurs when balance between the generation of reactive oxygen species (ROS) exceeds that of ROS removal, thus inducing oxidative damage. ROS are produced at a controlled rate during normal cell metabolism( 16 ). However, under stressful conditions such as nutritional alterations, ROS may dramatically increase and induce oxidative stress. Accordingly, variations in diet macronutrient sources( 7 , 17 25 ), dietary protein:non-protein ratios( 26 29 ) or non-protein energy sources( 30 ) have been linked to modifications of fish susceptibility to lipid peroxidation (LPO).

The knowledge about the effects of macronutrients on oxidative stress is still limited and the available data are contradictory. Regarding the effect of dietary lipid source, VO were reported to either decrease( 7 , 18 , 20 , 23 , 24 ), increase( 22 ) or promote no effects( 31 ) in fish hepatic and/or muscular LPO susceptibility. High dietary protein and/or lipid concentrations were observed to increase muscular LPO susceptibility( 26 , 28 ), whereas dietary carbohydrates decreased LPO in muscles and the liver( 27 , 28 , 30 ).

To prevent oxidative damage, organisms have evolved effective antioxidant systems, consisting of both non-enzymatic (GSH, vitamins and carotenoids, among others) and enzymatic mechanisms. Enzymes such as catalase (CAT), superoxide dismutase (SOD), glutathione peroxidase (GPX) and glutathione reductase (GR) play key roles in the antioxidant defence mechanism( 32 , 33 ). Available literature suggests that dietary lipid source has no effect on the liver and muscle activities of SOD, CAT and/or GPX( 22 , 31 , 34 ) or decreased SOD activity in VO-rich diets( 35 ) in a number of fish species.

Concerning macronutrient modifications, it was reported that in common dentex (Dentex dentex) high dietary lipid concentrations induced an increase in hepatic SOD activity at high dietary protein concentrations, and of GPX, independent of dietary protein concentrations( 28 ). The authors also observed an interaction between dietary protein and carbohydrate or lipid concentrations in muscular GR and hepatic SOD and CAT activities. In yellow catfish (Pelteobagrus fulvidraco), a decrease in CAT and GPX activities as dietary carbohydrate:lipid ratio increased was also observed( 30 ). In contrast, partial substitution of dietary proteins with carbohydrates induced no major alterations in antioxidant enzyme activities in Senegalese sole (Solea senegalensis) liver( 29 ).

Overall, current knowledge about the effects of alternative feed ingredients, which may be rich in carbohydrates or VO, on the oxidative status of animals is still very scarce. Thus, in this study, we evaluated the effect of the dietary lipid source and carbohydrate manipulation on oxidative defence mechanisms in the liver and intestine of European sea bass (Dicentrarchus labrax) juveniles.

Methods

Experimental diets

Four diets differing in carbohydrate content (0 and 20 % gelatinised starch, CH– and CH+, respectively) and lipid source (fish oil (FO) or a VO blend, FO and VO, respectively) were formulated (Table 1). The dietary carbohydrate content was increased at the expense of dietary protein. The VO blend was composed of 20 % rapeseed, 50 % linseed and 30 % palm oils and replaced approximately 70 % of dietary lipids, which were provided by cod liver oil and FM. All the ingredients were finely ground, well-mixed and dry-extruded in a laboratory pellet mill (California Pellet Mill), through a 3-mm die. The pellets were air-dried for 24 h and stored in a refrigerator (4°C) until used.

Table 1 Ingredients and chemical composition of the experimental diets

FO, fish oil; VO, blend of vegetable oils; carbohydrate content, 0 % (CH–) or 20 % (CH+) gelatinised maize starch.

* Steam-dried low-temperature fishmeal; Pesquera Diamante (CP: 71·1 % DM; CL: 8·8 % DM).

C-Gel Instant-12018; Cerestar.

Labchem, Laborspirit Lda.

§ 30 % Palm oil (Colmi), 50 % linseed oil (Sociedade Portuense de Drogas) and 20 % rapeseed oil (Huilerie Emile Noël SAS).

|| Vitamins (mg/kg diet): retinyl acetate, 18 000 IU (6·19 mg)/kg diet; cholecalciferol, 2000 IU (0·04 mg)/kg diet; α-tocopherol acetate, 35; sodium menadione bisulphate, 10; thiamin-HCl, 15; riboflavin, 25; calcium pantothenate, 50; nicotinic acid, 200; pyridoxine HCl, 5; folic acid 10; cyanocobalamin, 0·02; biotin, 1·5; ascorbic acid, 50; inositol, 400 (Premix).

Minerals (mg/kg diet): cobalt sulphate, 1·91; copper sulphate, 19·6; iron sulphate, 200; sodium fluoride, 2·21; potassium iodide, 0·78; magnesium oxide, 830; manganese oxide, 26; sodiumselenite, 0·66; zinc oxide, 37·5; dibasic calcium phosphate, 5·93 (g/kg diet); potassium chloride, 1·15 (g/kg diet); sodium chloride, 0·40 (g/kg diet) (Premix).

** Aquacube (Guar gum, polymethyl carbamide, Manioc starch blend, hydrate calcium sulphate) (Agil).

Animals, experimental conditions and sampling

This experiment was directed by accredited scientists (following FELASA category C recommendations) and conducted according to the European Union Directive (2010/63/EU) on the protection of animals for scientific purposes. The study was performed at the Marine Zoological Station, University of Porto, Portugal, in a thermo-regulated recirculation water system, equipped with twelve fibre-glass cylindrical tanks of 300 litre water capacity, and supplied with continuous flow of filtered seawater.

After 2 weeks of adaptation to the experimental conditions, twelve groups of twenty European sea bass juveniles (initial body weight 74·0 (sd 1·5) g) were established and randomly distributed into the tanks. The experimental diets were randomly assigned to triplicate groups of fish (n 3 tanks/diet). During the trial, salinity averaged 35 (sd 1·0) g/l, dissolved oxygen was kept near saturation and water temperature was regulated to 25·4 (sd 0·5)°C. The photoperiod was the natural one for June–August. The trial lasted 73 d, and during that period fish were hand-fed twice a day, 6 d/week, to apparent visual satiety. Food intake and mortality were recorded daily, and fish in each tank were bulk-weighted at the end of the trial, after 1 d of feed deprivation to empty gut contents. To eliminate handling stress, the fish were continued to be fed for one more week, and subsequently 18 h after the last meal (previous day afternoon meal) three fish per tank (n 9 fish/diet) were killed with a sharp blow to the head; the liver and intestines were removed, immediately frozen in liquid N2 and stored at –80°C until enzymatic analyses.

Diet analysis

Chemical analyses of the experimental diets were carried out according to Association of Official Analytical Chemists( 36 ) as follows: in brief, DM after drying at 105°C until constant weight; ash by incineration in a muffle furnace at 450°C for 16 h; protein content (N×6·25) by the Kjeldahl method after acid digestion using a Kjeltec digestion and distillation units (models 1015 and 1026, respectively; Tecator Systems); starch content according to Beutler( 37 ); gross energy by direct combustion in an adiabatic bomb calorimeter (Parr model 6200; Parr Instruments); and lipid content by petroleum diethyl ether extraction (Soxtec HT System; Höganäs). Fatty acid methyl esters were prepared by acid-catalysed transmethylation of total lipids using boron trifluoride in methanol (14 %) according to Santha & Ackman( 38 ) and were analysed by GC (Varian 3900; Varian) as described by Castro et al.( 39 ).

Enzymatic activity assays

Liver and whole intestine samples were homogenised (dilution 1:4) in ice-cold buffer (100 mm-Tris-HCl, 0·1 mm-EDTA and 0·1 % triton X-100 (v/v), pH 7·8). All the procedures were performed on ice. Homogenates were centrifuged at 30 000 g for 30 min at 4°C. After centrifugation, the resultant supernatant was collected, and the aliquots were stored at –80 °C until analysis.

All enzyme assays were carried out at 37°C. Changes in absorbance were monitored using a microplate reader (ELx808; BioTek Instruments) to determine enzyme activities. The optimal substrate and protein concentrations for maximal activity measurement of each enzyme were established by preliminary assays. The molar extinction coefficients used for H2O2 and NADPH were 0·039 and 6·22 mm −1×cm−1, respectively.

The specific assay conditions for each enzyme were as follows.

SOD (EC 1.15.1.1) activity was measured at 550 nm by the ferricytochrome C method using xanthine/xanthine oxidase as the source of superoxide radicals( 40 ). The reaction mixture consisted of 50 mm-potassium phosphate buffer (pH 7·8), 0·1 mm-EDTA, 0·1 mm-xanthine, 0·012 mm-cytochrome C and 0·025 IU/ml xanthine oxidase. Activity was reported as units per mg of protein. One unit of activity was defined as the amount of enzyme necessary to produce 50 % inhibition of ferricytochrome C reduction rate.

CAT (EC 1.11.1.6) activity was determined according to Aebi( 41 ) by measuring the decrease in H2O2 concentration at 240 nm. The reaction mixture contained 50 mm-potassium phosphate buffer (pH 7) and 10 mm-H2O2 freshly added.

GPX (EC 1.11.1.9) activity was assayed as described by Flohé & Günzler( 42 ). The GSSG generated by GPX was reduced by GR, and NADPH consumption rate was monitored at 340 nm. The reaction mixture consisted of 50 mm-potassium phosphate buffer (pH 7·1), 1 mm-EDTA, 3·9 mm-GSH, 3·9 mm-sodium azide, 1 IU/ml GR, 0·2 mm-NADPH and 0·05 mm-H2O2.

GR (EC 1.6.4.2) activity was determined at 340 nm by measuring the oxidation of NADPH as described by Morales et al.( 43 ). The reaction mixture consisted of 0·1 m-sodium phosphate buffer (pH 7·5), 1 mm-EDTA, 0·63 mm-NADPH and 0·16 mm-GSSG.

Glucose-6-phosphate dehydrogenase (G6PD; EC 1.1.1.49) activity was assayed by measuring the reduction of NADP+ at 340 nm as previously described by Morales et al.( 44 ), using a reaction mixture containing 50 mm-imidazole–HCl buffer (pH 7·4), 5 mm-MgCl2, 2 mm-NADP and 1 mm-glucose-6-phosphate.

Except for SOD, whose units of expression were described above, all other enzyme activities were expressed as units (CAT) or milliunits (G6PD, GPX and GR) per mg of soluble protein. One unit of enzyme activity was defined as the amount of enzyme required to transform 1 μmol of substrate/min under the above assay conditions. Protein concentration was determined according to Bradford( 45 ) using a protein assay kit (Sigma) and bovine serum albumin as the standard.

Total glutathione, GSH and oxidative stress index

A portion of liver was homogenised in nine volumes of ice-cold solution containing 1·3 % 5-sulphosalicylic acid (w/v) and 10 mm-HCl. The procedure was carefully performed always on ice in order to avoid the GSH. Homogenates were centrifuged at 14 000 g for 10 min at 4°C, and the resulting supernatants were immediately analysed.

Total glutathione (tGSH) and GSSG were measured following the method described by Griffith( 46 ) and Vandeputte et al.( 47 ) with modifications. Both tGSH and GSSG analyses were carried out at 37°C, and the changes in absorbance as a consequence of the reduction of 5,5′-dithiobis (2-nitrobenzoic acid) (DTNB) were monitored at 405 nm using a microplate reader (ELx808; BioTek Instruments). The optimal substrate and protein concentrations for the measurement of maximal activity for each enzyme were established by preliminary assays. The molar extinction coefficient used for DTNB was 13·6 mm −1×cm−1.

tGSH was determined using a reaction mixture containing 133 mm-phosphate buffer with 5·8 mm-EDTA at pH 7·4, 0·71 mm-DTNB, 0·24 mm-NADPH and 1·2 IU/ml GR.

GSSG was measured using an aliquot from the solution obtained after 60 min of incubation of 100 μl of the sample with 2 μl vinylpyridine and 6 μl 1·5 m-triethanolamine. The reaction mixture contained 122 mm-phosphate buffer with 5·4 mm-EDTA at pH 7·4, 0·71 mm-DTNB, 0·24 mm-NADPH and 1·2 IU/ml GR.

The results were calculated using the standard curves of GSH and GSSG for tGSH and GSSG measurements, respectively. GSH level was calculated by subtracting GSSG from tGSH values. The results are expressed as nmol per g of tissue.

Oxidative stress index (OSI) was calculated as follows:

OSI (%)=100×(2×GSSG/tGSH).

Lipid peroxidation

Concentration of malondialdehyde (MDA) was determined as a marker for LPO following the methodology described by Buege & Aust( 48 ). An aliquot of supernatant from the homogenate (100 μl) was mixed with 500 μl of a previously prepared solution containing 15 % (w/v) TCA, 0·375 % (w/v) thiobarbituric acid (TBA), 80 % (v/v) HCl 0·25 n and 0·01 % (w/v) butylated hydroxytoluene. The mixture was heated to 100°C for 15 min. After being cooled to room temperature and centrifuged at 1500 g for 10 min, the absorbance was measured at 535 nm in the supernatant. MDA concentration was expressed as nmol MDA per g of tissue, calculated from a calibration curve.

Statistical analysis

Data are presented as mean values and standard deviations. Data were checked for normality and homogeneity of variances and when appropriate were normalised (log, inverse and arcsin $\left( {\sqrt {\left( {x\,/\,{\rm 1}00} \right)} } \right)\ {\rm transformations})$ . Statistical evaluation of the data was carried out using a 2×2 factorial arrangement of treatments in a completely randomised experimental design (two-way ANOVA), with carbohydrate concentration and lipid source as fixed factors. The significance level of 0·05 was used for rejection of the null hypothesis. All the statistical analyses were carried out using SPSS 21.0 software package (IBM Corp.) for Windows.

Results

The diets presented small differences in the proportions of total SFA, which were slightly higher in the VO diets, and of MUFA, which were higher in the FO diets (Table 2). Within MUFA, higher concentrations of oleic acid (18 : 1n-9) were found in the VO diets, whereas the opposite occurred for palmitoleic acid (16 : 1n-7), eicosenoic acid (20 : 1n-9) and erucic acid (22 : 1n-9). Linoleic acid (LA, 18 : 2n-6), n-6 PUFA, was considerably higher in the VO diets. Regarding n-3 PUFA, VO diets were particularly rich in linolenic acid (LNA, 18 : 3n-3) and poor in EPA(20 : 5n-3) and DHA) (22 : 6n-3).

Table 2 Fatty acid composition (% of total fatty acids) of the experimental diets

FO, fish oil, VO, blend of vegetable oils; carbohydrate content, 0 % (CH–) or 20 % (CH+) gelatinised maize starch;·n-3 LC-PUFA, n-3 long-chain PUFA.

Results of the feeding trial were not the aim of this study and are presented elsewhere( 49 ).

In brief, growth performance and feed efficiency were not affected by dietary treatments. Final body mass averaged 221·7 (sd 5·0) g, feed efficiency (wet weight gain(g)/dry feed intake (g)) averaged 0·8 (sd 0·03) and feed intake (g/kg average body mass per d) averaged 16·6 (sd 0·7). Liver fatty acid composition is summarised in Table 3. Accordingly, liver samples of fish fed VO-based diets were characterised by higher concentrations of LA (18 : 2n-6), γ-LNA (GLA; 18 : 3n-6) and LNA (18 : 3n-3) and by lower concentration of arachidonic acid (ARA; 20 : 4n-6), EPA (20 : 5n-3) and DHA (22 : 6n-3) as well as n-3 PUFA:n-6 PUFA ratio. Except for ARA concentrations, lower in groups fed the CH+ diets, no other differences were found for hepatic n-6 PUFA concentrations. Regarding n-3 PUFA, groups fed the CH+ diets presented similar LNA concentrations but lower EPA and DHA concentrations compared with those fed CH– diets. Unsaturation index was lower in fish fed the CH+ diets. On the other hand, differences between lipid sources on unsaturation index were only evident in the CH– group. During the trial, no mortality occurred.

Table 3 Liver fatty acid (FA) profile (% of total FA) of European sea bass fed the experimental diets (Mean values and standard deviations; n 6 fish)

FO, fish oil; VO, blend of vegetable oils; CH content, 0 % (CH–) or 20 % (CH+) gelatinised maize starch.

* Significant differences at P<0·05 (two-way ANOVA).

Overall, both in the liver and in the intestines, there were no interactions between dietary carbohydrate and lipid source in the activities of oxidative stress enzymes or in LPO values. In the liver, dietary lipid source did not affect the activities of oxidative stress enzymes or LPO values, whereas dietary carbohydrates promoted a decrease in LPO values and in CAT activity and an increase in GR and G6PD activities (Table 4). Moreover, in the liver, SOD and GPX activities were not affected by dietary carbohydrates. In the intestine, the dietary carbohydrate level did not affect the activities of oxidative stress enzymes or LPO values, whereas dietary VO promoted a decrease in LPO and an increase in GPX and GR activities (Table 5). No effects on activities of CAT, SOD or G6PD in relation to diet composition were observed in the intestine.

Table 4 Specific activities of glucose-6-phosphate dehydrogenase (G6PD), glutathione peroxidase (GPX), glutathione reductase (GR) (mU/mg protein), catalase (CAT), superoxide dismutase (SOD) (U/mg protein) and lipid peroxidation (LPO) (nmol malondialdehyde per g tissue) in the liver of European sea bass fed the experimental diets(Mean values and standard deviations; n 9 fish)

FO, fish oil;, VO, blend of vegetable oils; CH content, 0 % (CH–) or 20 % (CH+) gelatinised maize starch.

* Significant differences at P<0·05 (two-way ANOVA).

Table 5 Specific activities of glucose-6-phosphate dehydrogenase (G6PD), glutathione peroxidase (GPX), glutathione reductase (GR) (mU/mg protein), superoxide dismutase (SOD), catalase (CAT) (U/mg protein) and lipid peroxidation (LPO) (nmol malondialdehyde per g tissue) in the intestine of European sea bass fed the experimental diets(Mean values and standard deviations; n 9 fish)

FO, fish oil; VO, blend of vegetable oils; CH content, 0 % (CH–) or 20 % (CH+) gelatinised maize starch.

* Significant differences at P<0·05 (two-way ANOVA).

In the liver, higher GSH content and lower GSSG levels and OSI were observed in the carbohydrate groups (Fig. 1), but no effects of dietary lipid source on glutathione (total, reduced and oxidised) or OSI were observed.

Fig. 1 Total glutathione (tGSH), GSH, GSSG and oxidative stress index (OSI) in the liver (L) of European sea bass fed experimental diets. Fish oil (FO), blend of vegetable oils (VO); carbohydrate content: 0 % (, CH–) or 20 % (, CH+) gelatinised maize starch. Values are means (n 9 fish), with standard deviations represented by vertical bars. Significant differences at P<0·05 (two-way ANOVA).

Discussion

In this study, low susceptibility to LPO was observed in the liver of fish fed carbohydrate-rich diets, which is in agreement with previous observations on muscles of this and other fish species( 27 , 28 ). The beneficial effect of dietary carbohydrates on lipid oxidative damage may be related to the nature of the glucose molecule, which can scavenge ROS by itself( 50 ), or through increased activity of the pentose phosphate pathway( 27 , 28 , 50 , 51 ). The pentose phosphate pathway, via G6PD, generates reducing power in the form of NADPH, which is required for the activity of two H2O2-scavenging pathways involving CAT and GPX( 43 , 52 ). In fact, along with lower hepatic LPO levels, higher G6PD activity was observed in European sea bass fed carbohydrate-rich diets. Furthermore, dietary carbohydrates also promoted an increase in hepatic GR activity, an enzyme that also requires NADPH to regenerate reduced glutathione (GSH) to its oxidised form (GSSG). Consistent with this observation, higher concentration of GSH and lower concentration of GSSG were observed in the liver of fish fed carbohydrate-rich diets. GSH is an essential substrate for GPX, but as it is also an effective ROS scavenger it plays an important role by preventing some of the deleterious effects of oxidative stress( 53 ). Accordingly, liver OSI, a biomarker of the cellular redox state, was also lower in groups fed carbohydrate-rich diets. Studies using OSI to assess the effects of oxidative stress in fish are scarce( 54 58 ) and mainly report a huge variation in OSI values (from 0·6 to 30 %) at the hepatic level( 55 58 ). As several variables (such as fish species, abiotic factors, fish size, time elapsed from the last meal or nutritional history) may regulate OSI values, comparisons among studies or establishment of reliable reference values for this parameter are difficult. Furthermore, a number of parameters or techniques have traditionally been used as measurements of oxidative stress. Among the most studied are antioxidant defence enzymes, TBA reactive substances (TBARS), concentrations of tGSH and GSSG and resulting OSI. In this study, the response of most parameters analysed (including OSI) was well correlated. However, other studies( 55 , 58 ) have demonstrated that OSI was not a suitable biomarker of oxidative stress under stressful conditions. As stated by Peres et al.( 59 ), reliable parameters are those that respond to stressful conditions but have low variation under normal conditions. This highlights the importance of taking into consideration the response of more than one parameter involved in the oxidative status( 55 , 58 ).

On the other hand, the carbohydrate-rich diets promoted a lower hepatic CAT activity. Previously, Lygren & Hemre( 50 ) also reported that hepatic CAT and SOD activities decreased in Atlantic salmon (Salmo salar) fed high-carbohydrate diets. These observations suggest that the lower LPO observed in fish fed the carbohydrate diets was mainly related with the higher concentrations of reducing equivalents such as NADPH and GSH and that activation of primary antioxidant enzymes, which are more energy-demanding, was not required.

Fatty acid composition of reserve lipids in tissues (liver, muscle, intestine, etc.) reflects dietary fatty acid profile( 60 ). In the present study, the reduced n-3:n-6 ratios (2·3 to 2·7 in VO groups v. 7·5 to 5·4 in FO groups) in the hepatic fatty acid profile of European sea bass fed VO-blend diets are consistent with previously reported values (1·9 to 2·5 in VO groups v. 4·3 in FO group) for this species also fed similar VO-blend based diets( 5 ). FO is rich in long-chain PUFA (LC-PUFA), whereas VO is rich in PUFA, and it is well established that due to their high degree of unsaturation LC-PUFA are more susceptible to peroxidation than PUFA or MUFA, thus promoting the formation of more free radicals( 16 ). Accordingly, in this study, higher hepatic LC-PUFA content was observed in European sea bass fed the FO-based diets than the VO-rich diets( 49 ), although liver oxidative status was not affected by dietary lipid source. In fact, although unsaturation index was lower in fish fed the carbohydrate-rich diets, differences related to lipid sources were only evident in the CH– group (lower in VO diet). Similarly, partial replacement of FO by VO did not induce changes in hepatic lipid oxidative damages and antioxidant enzyme activities of Atlantic cod (Gadus morhua) or rockfish (Sebastes schlegeli)( 31 , 34 ) or in LPO concentration in Japanese sea bass (Lateolabrax japonicas)( 23 ).

In contrast to what was observed in the liver, in the present study, the dietary lipid source affected the intestinal oxidative status in European sea bass juveniles. Overall, oxidative lipid stress measured as LPO concentration was higher in fish fed the FO-based diets than the VO-rich diets. Studies addressing the effect of dietary lipid source on fish oxidative status were mainly focused on liver and muscle tissues( 7 , 18 , 20 , 22 24 , 31 , 35 ). To the best of the authors’ knowledge, the impact of dietary lipid source on intestinal oxidative status of fish was only evaluated in Japanese sea bass, and it was limited to the assessment of LPO concentration( 23 ). In that study, dietary inclusion of palm oil led to decreased intestinal LPO concentration. Available studies dealing with the modulator role of nutrition on intestinal antioxidant enzymatic mechanisms were mainly dedicated to the role of amino acids( 61 , 62 ), anti-nutritional factors( 63 ) and NSP( 64 ).

GPX catalyses the reduction of H2O2 and lipid peroxides (ROOH) into the corresponding stable alcohols (ROH) at the expense of GSH( 32 ). GR catalyses the reduction of GSSG to GSH and its activity is tightly bound to that of GPX, in an attempt to keep the adequate GSSG:GSH ratio necessary for GPX function. In the present study, GPX and GR activities were responsive to dietary treatments and had increased activities in groups fed the VO diets. Although flavonoids and phenolic compounds of VO were not measured in the present study, their potential role in the intestinal GPX and GR activation cannot be discarded. There is evidence that some plant constituents (flavonoids, phenolic compounds) also present in VO have antioxidant functions in mammals( 65 67 ). This antioxidant effect may result from a direct free-radical scavenging action by these compounds or from their role in activation of important antioxidant enzymes( 65 67 ). Accordingly, Nagata et al ( 65 ) observed in vitro that in rat hepatocytes two flavonoids (quercetin and catechin) exert their antioxidant activity against H2O2 through activation of GPX. In human hepatocytes, it was also demonstrated that cocoa flavonoids exert an antioxidant protective effect against tert-butyl hydroperoxide via induction of GPX and GR activities. In fish, studies are not yet available. However, Sitjà-Bobadilla et al.( 68 ) suggested that replacement of FM by a mixture of plant ingredients in diets for gilthead sea bream (Sparus aurata) exerts an antioxidative effect through activation of glutathione metabolism, as plant-containing diets increased muscular GSH and hepatic glutathione redox status, as well as GR activity, in both hepatic and muscular tissues. According to the authors, this increased antioxidant potential related to plant ingredients was probably linked to the presence of phenolic compounds and/or flavonoids. Further, Saera-Vila et al.( 69 ) reported that FO replacement by VO in plant protein-based diets increased hepatic GSH:GSSG ratio, despite the reduction of absolute glutathione concentrations. According to the authors, this suggests that fish fed these diets may be less susceptible to LPO and oxidative damage. Overall, both the fatty acid profile of VO-based diets and the presence of plants constituents with antioxidant potential may explain the lower intestinal susceptibility to oxidative stress/damages in VO groups.

Transcriptional and proteomic studies have also evidenced that intestinal oxidative status may be affected by changes in dietary lipid source( 70 , 71 ). Accordingly, Morais et al. ( 71 ) reported that in Atlantic salmon the dietary FO substitution by VO led to an up-regulation of transcripts and proteins involved in detoxification and protection from oxidative stress, CAT and selenoprotein, as well as haemopexin-like protein and peroxiredoxin-1 proteins. On the contrary, a previous study on the same species showed that dietary FO replacement by VO decreased the transcription levels of antioxidant-related genes such as glutathione S-transferase and GR( 70 ).

Similarly to our previous studies on sturgeon (Acipenser naccarii) and rainbow trout (Oncorhynchus mykiss)( 72 ), in this study, European sea bass intestines also showed LPO damage values and GPX activity 2- to 5-fold higher compared with the liver. GPX and GR activities were also 2-fold higher in the intestine than in the liver of European sea bass. On the contrary, G6PD and CAT activities were, respectively, more than 10- and 5-fold lower in the intestine than in the liver. These differences between the two organs are a reflex of the distinct antioxidant mechanism strategies that are more relevant in each organ. The intestine being a tissue with high cell turnover is highly susceptible to oxidative status, and seems to require high activity of antioxidant enzymes.

In conclusion, results of the present study indicate that liver and intestine oxidative status balance were regulated through different metabolic pathways, and that dietary macronutrients affected metabolic pathways differently in each tissue. In addition, the observed LPO values suggest that susceptibility to oxidative stress was more important in the intestine than in the liver. Dietary carbohydrates contributed to reduced oxidative stress only in the liver, whereas the intestine was exclusively responsive to the dietary lipid source, with VO having a positive effect on the reduction of oxidative stress. Nowadays a wide range of PF are already used in commercial aquafeeds, including cereals and peas, which are rich sources of starch, and may therefore have a protective effect on liver oxidative stress. Given the importance of the intestine on whole-body homoeostasis and health, increased attention should be paid to this tissue in future studies evaluating the impact of dietary alternative ingredients on oxidative stress.

Acknowledgements

The authors express their thanks to P. Correia for technical assistance and L. Larroquet for the fatty acid analyses.

This work was partially supported by the FCT (Foundation for Science and Technology), Portugal (project PTDC/MAR-BIO/4107/2012), and co-financed by the European Regional Development Fund (ERDF) through the COMPETE – Operational Competitiveness Programme and national funds through FCT – under the project ‘PEst-C/MAR/LA0015/2011’. C. C., A. P.-J. and F. C. were supported by grants (SFRH/BD/76297/2011; SFRH/BPD/64684/2009; SFRH/BD/86799/2012, respectively) from FCT. P. D.-R. and C. R. S. were supported by grants (NORTE-07-0124-FEDER-000038-BPD-2013-07; NORTE-07-0124-FEDER-000038-BPD-2013-05, respectively).

C. C. carried out the main experimental work and wrote the draft of the manuscript under the direction of the project designer and leader A. O.-T., G. C., S. P. A. P.-J., F. C., P.D.-R., C. R. S. and H. P. assisted with the biochemical analyses and draft writing. All the authors contributed to and approved the manuscript.

The authors declare that there are no conflicts of interest.

References

1. National Research Council (2011) Replacement of marine resources: effects of alternative ingredients and strategies on nutritional quality. In Nutrient Requirements of Fish, pp. 304322. Washington, DC: National Academies Press.
2. Montero, D, Kalinowski, T, Obach, A, et al. (2003) Vegetable lipid sources for gilthead seabream (Sparus aurata): effects on fish health. Aquaculture 225, 353370.
3. Montero, D, Grasso, V, Izquierdo, MS, et al. (2008) Total substitution of fish oil by vegetable oils in gilthead seabream (Sparus aurata) diets: effects on hepatic Mx expression and some immune parameters. Fish Shellfish Immunol 24, 147155.
4. Mourente, G, Good, JE & Bell, JG (2005) Partial substitution of fish oil with rapeseed, linseed and olive oils in diets for European sea bass (Dicentrarchus labrax L.): effects on flesh fatty acid composition, plasma prostaglandins E2 and F2α, immune function and effectiveness of a fish oil finishing diet. Aquac Nutr 11, 2540.
5. Mourente, G, Good, JE, Thompson, KD, et al. (2007) Effects of partial substitution of dietary fish oil with blends of vegetable oils, on blood leucocyte fatty acid compositions, immune function and histology in European sea bass (Dicentrarchus labrax L.). Br J Nutr 98, 770779.
6. Bell, JG, Strachan, F, Good, JE, et al. (2006) Effect of dietary echium oil on growth,fatty acid composition and metabolism, gill prostaglandin production and macrophage activity in Atlantic cod (Gadus morhua L.). Aquac Res 37, 606617.
7. Lin, YH & Shiau, SY (2007) Effects of dietary blend of fish oil with corn oil on growth and non-specific immune responses of grouper, Epinephelus malabaricus. Aquac Nutr 13, 137144.
8. Ganga, R, Bell, JG, Montero, D, et al. (2005) Effect of feeding gilthead sea bream (Sparus aurata) with vegetable lipid sources on two potential immunomodulator products: prostanoids and leptins. Comp Biochem Physiol B 142, 410418.
9. Olsen, RE, Myklebust, R, Kaino, T, et al. (1999) Lipid digestibility and ultrastructural changes in the enterocytes of Arctic charr (Salvelinus alpinus L.) fed linseed oil and soybean lecithin. Fish Physiol Biochem 21, 3544.
10. Caballero, MJ, Obach, A, Rosenlund, G, et al. (2002) Impact of different dietary lipid sources on growth, lipid digestibility, tissue fatty acid composition and histology of rainbow trout, Oncorhynchus mykiss . Aquaculture 214, 253271.
11. Benedito-Palos, L, Navarro, JC, Sitjá-Bobadilla, A, et al. (2008) High levels of vegetable oils in plant protein-rich diets fed to gilthead seabream (Sparus aurata L.): growth performance, muscle fatty acid profiles and histological alterations of target tissues. Br J Nutr 100, 9921003.
12. Kjaer, MA, Vegusdal, A, Gjøen, T, et al. (2008) Effect of rapeseed oil and dietary n-3 fatty acids on triacylglycerol synthesis and secretion in Atlantic salmon hepatocytes. Biochim Biophys Acta 1781, 112122.
13. Kortner, TM, Gu, J, Krogdahl, A, et al. (2012) Transcriptional regulation of cholesterol and bile acid metabolism after dietary soyabean meal treatment in Atlantic salmon (Salmo salar L.). Br J Nutr 30, 112.
14. Gatlin, DM III, Barrows, FT, Brown, P, et al. (2007) Expanding the utilization of sustainable plant products in aquafeeds: a review. Aquac Res 38, 551579.
15. Montero, D & Izquierdo, M (2010) Welfare and health of fish fed vegetable oils as alternative lipid sources to fish oil. In Fish Oil Replacement and Alternative Lipid Sources in Aquaculture Feeds, pp. 439485 [GM Turchini, WK Ng and DR Tocher, editors]. Boca Raton, FL: CRC Press, Taylor & Francis.
16. Halliwell, B & Chirico, S (1993) Lipid peroxidation: its mechanism, measurement, and significance. Am J Clin Nutr 57, 715S725S.
17. López-Bote, CJ, Diez, A, Corraze, G, et al. (2001) Dietary protein source affects the susceptibility to lipid peroxidation of rainbow trout (Oncorhynchus mykiss) and sea bass (Dicentrarchus labrax) muscle. Anim Sci 73, 443449.
18. Menoyo, D, Izquierdo, MS, Robaina, L, et al. (2004) Adaptation of lipid metabolism, tissue composition and flesh quality in gilthead sea bream (Sparus aurata) to the replacement of dietary fish oil by linseed and soyabean oils. Br J Nutr 92, 4152.
19. Rueda-Jasso, R, Conceição, LEC, Dias, J, et al. (2004) Effect of dietary non-protein energy levels oncondition and oxidative status of Senegalese sole (Solea senegalensis) juveniles. Aquaculture 231, 417433.
20. Peng, SM, Chen, LQ, Qin, JG, et al. (2008) Effects of replacement of dietary fish oil by soybean oil on growth performance and liver biochemical composition in juvenile black seabream, Acanthopagrus schlegeli. Aquaculture 276, 154161.
21. Kiron, V, Thawonsuwan, J, Panigrahi, A, et al. (2011) Antioxidant and immune defences of rainbow trout (Oncorhynchus mykiss) offered plant oils differing in fatty acid profiles from early stages. Aquac Nutr 17, 130140.
22. Ostbye, TK, Kjaer, MA, Rora, AMB, et al. (2011) High n-3 HUFA levels in the diet of Atlantic salmon affect muscle and mitochondrial membrane lipids and their susceptibility to oxidative stress. Aquac Nutr 17, 177190.
23. Gao, J, Koshio, S, Ishikawa, M, et al. (2012) Effects of dietary palm oil supplements with oxidized and non-oxidized fish oil on growth performances and fatty acid compositions of juvenile Japanese sea bass, Lateolabrax japonicas . Aquaculture 324–325, 97103.
24. Ng, WK, Chong, CY, Wang, Y, et al. (2013) Effects of dietary fish and vegetable oils on the growth, tissue fatty acid composition, oxidative stability and vitamin E content of red hybrid tilapia and efficacy of using fish oil finishing diets. Aquaculture 372, 97110.
25. Bañuelos-Vargas, I, López, LM, Pérez-Jiménez, A, et al. (2014) Effect of fishmeal replacement by soy protein concentrate with taurine supplementation on hepatic intermediary metabolism and antioxidant status of totoaba juveniles (Totoaba macdonaldi). Comp Biochem Physiol B 170, 1825.
26. Alvarez, MJ, López-Bote, CJ, Díez, A, et al. (1998) Dietary fish oil and digestible protein modify susceptibility to lipid peroxidation in the muscle of rainbow trout (Oncorhynchus mykiss) and sea bass (Dicentrarchus labrax). Br J Nutr 80, 281289.
27. Alvarez, ML, López-Bote, CJ, Díez, A, et al. (1999) The partial substitution of digestible protein with gelatinized starch as an energy source reduces susceptibility to lipid oxidation in rainbow trout (Oncorhynchus mykiss) and sea bass (Dicentrarchus labrax) muscle. J Anim Sci 77, 33223329.
28. Pérez-Jiménez, A, Hidalgo, MC, Morales, AE, et al. (2009) Antioxidant enzymatic defenses and oxidative damage in Dentex dentex fed on different dietary macronutrient levels. Comp Biochem Physiol C 150, 537545.
29. Castro, C, Pérez-Jiménez, A, Guerreiro, I, et al. (2012) Effects of temperature and dietary protein level on hepatic oxidative status of Senegalese sole juveniles (Solea senegalensis). Comp Biochem Physiol A 163, 372378.
30. Wang, LN, Liu, WB, Lu, KL, et al. (2014) Effects of dietary carbohydrate/lipid ratios on non-specific immune responses, oxidative status and liver histology of juvenile yellow catfish Pelteobagrus fulvidraco . Aquaculture 426–427, 4148.
31. Kjaer, MA, Aursnes, IA, Berge, GM, et al. (2014) The influence of different dietary oil qualities on growth rate, feed utilization and oxidative stress in Atlantic cod. Aquac Nutr 20, 192204.
32. Mourente, G, Bell, JG & Tocher, DR (2007) Does dietary tocopherol level affect fatty acid metabolism in fish? Fish Physiol Biochem 33, 269280.
33. Lushchak, VI (2011) Environmentally induced oxidative stress in aquatic animals. Aquat Toxicol 101, 1330.
34. Aminikhoei, Z, Choi, J, Lee, SM, et al. (2013) Effects of different dietary lipid sources on gowth performance, fatty acid composition, and antioxidant enzyme activity of juvenile rockfish, Sebastes schlegeli . J World Aquacult 44, 716725.
35. Kjaer, MA, Todorcevic, M, Torstensen, BE, et al. (2008) Dietary n-3 HUFA affects mitochondrial fatty acid beta-oxidation capacity and susceptibility to oxidative stress in Atlantic salmon. Lipids 43, 813827.
36. Association of Official Analytical Chemists (2000) Official Methods of Analysis. Gaithersburg, MD: Association of Official Analytical Chemists.
37. Beutler, HO (1984) Starch. In Methods of Enzymatic Analysis 6, pp. 210 [HU Bergmeyer, editor]. Basel: Verlag, Chemie Weinheim.
38. Santha, NC & Ackman, RG (1990) Nervonic acid versus tricosanoic acid as internal standards in quantitative gas chromatographic analyses of fish oil longer-chain n-3 polyunsaturated fatty acid methyl esters. J. Chromatogr Biomed Appl 533, 110.
39. Castro, C, Corraze, G, Panserat, S, et al. (2014) Effects of fish oil replacement by a vegetable oil blend on digestibility, postprandial serum metabolite profile, lipid and glucose metabolism of European sea bass (Dicentrarchus labrax) juveniles. Aquac Nutr (epublication ahead of print version 29 October 2014).
40. McCord, JM & Fridovich, I (1969) Superoxide dismutase: an enzymic function for erythrocuprein. J Biol Chem 244, 60496055.
41. Aebi, H (1984) Catalase in vitro . Methods Enzymol 105, 121126.
42. Flohé, L & Günzler, WA (1984) Assay of glutathione peroxidase. Methods Enzymol 105, 115121.
43. Morales, AE, Pérez-Jiménez, A, Hidalgo, MC, et al. (2004) Oxidative stress and antioxidant defenses after prolonged starvation in Dentex dentex liver. Comp Biochem Physiol C 139, 153161.
44. Morales, AE, García Rejón, L & de la Higuera, M (1990) Influence of handling and/or anaesthesia on stress response in rainbow trout. Effects on liver primary metabolism. Comp Biochem Physiol A 95, 8793.
45. Bradford, M (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein using the principle of protein dye-binding. Anal Biochem 72, 248254.
46. Griffith, OW (1980) Determination of glutathione and glutathione disulfide using gluthatione reductase and 2-vinylpyridine. Anal Biochem 106, 207212.
47. Vandeputte, C, Guizon, I, Genestie-Denis, I, et al. (1994) A microtiter plate assay for total glutathione and glutathione disulfide contents in cultured/isolated cells: performance study of a new miniaturized protocol. Cell Biol Toxicol 10, 415421.
48. Buege, JA & Aust, SD (1978) Microsomal lipid peroxidation. Methods Enzymol 52, 302310.
49. Castro, C, Corraze, G, Pérez-Jiménez, A, et al. (2015) How dietary carbohydrates and lipid source affect lipid and glucose metabolism of European sea bass (Dicentrarchus labrax) juveniles? Br J Nutr (epublication ahead of print version 26 August 2015).
50. Lygren, B & Hemre, GI (2001) Influence of dietary carbohydrate on antioxidant enzyme activities in liver of Atlantic salmon (Salmo salar L.). Aquac Int 9, 421427.
51. Sagone, AL, Greenwald, J, Kraut, EH, et al. (1983) Glucose: a role as a free radical scavenger in biological systems. J Lab Clin Med 101, 97103.
52. Nelson, DL & Cox, MM (2004) Glycolysis, gluconeogenesis, and the pentose phosphate pathway. In Lehninger Principles of Biochemistry , 4th ed., pp. 521559 [DL Nelson and MM Cox, editors]. New York: W.H. Freeman.
53. Wu, G, Fang, YZ, Yang, S, et al. (2004) Glutathione metabolism and its implications for health. J Nutr 134, 489492.
54. Olsvik, PA, Kristensen, T, Waagbo, R, et al. (2005) mRNA expression of antioxidant enzymes (SOD, CAT and GSHPx) and lipid peroxidative stress in liver of Atlantic salmon (Salmo salar) exposed to hyperoxic water during smoltification. Comp Biochem Physiol C 141, 314323.
55. Olsvik, PA, Kristensen, T, Waagbo, R, et al. (2006) Effects of hypo- and hyperoxia on transcription levels of five stress genes and the glutathione system in liver of Atlantic cod Gadus morhua . J Exp Biol 209, 28932901.
56. Heier, LS, Lien, IB, Strømseng, AE, et al. (2009) Speciation of lead, copper, zinc and antimony in water draining a shooting range – time dependant metal accumulation and biomarker responses in brown trout (Salmo trutta L.). Sci Tot Environ 407, 40474055.
57. Karlsson, A, Heier, LS, Rosseland, BO, et al. (2011) Changes in arterial PO2, physiological blood parameters and intracellular antioxidants in free-swimming Atlantic cod (Gadus morhua) exposed to varying levels of hyperoxia. Fish Physiol Biochem 37, 249258.
58. Pérez-Jiménez, A, Peres, H, Cruz Rubio, V, et al. (2012) The effect of hypoxia on intermediary metabolism and oxidative status in gilthead sea bream (Sparus aurata) fed on diets supplemented with methionine and white tea. Comp Biochem Physiol C 155, 506516.
59. Peres, H, Santos, S & Oliva-Teles, A (2012) Selected plasma biochemistry parameters in gilthead seabream (Sparus aurata) juveniles. J Appl Ichthyol 29, 17.
60. Rosenlund, G, Corraze, G, Izquierdo, M, et al. (2010) The effects of fish oil replacement on nutritional and organoleptic qualities of farmed fish. In Fish Oil Replacement and Alternative Lipid Sources in Aquaculture Feeds CRC Press, pp. 487522 [GM Turchini, WK Ng and DR Tocher, editors]. Boca Raton, FL: CRC Press, Taylor & Francis.
61. Wen, H, Feng, L, Jiang, W, et al. (2014) Dietary tryptophan modulates intestinal immune response, barrier function, antioxidant status and gene expression of TOR and Nrf2 in young grass carp (Ctenopharyngodon idella). Fish Shellfish Immunol 40, 275287.
62. Zhao, Y, Hu, Y, Zhou, XQ, et al. (2015) Effects of dietary glutamate supplementation on growth performance, digestive enzyme activities and antioxidant capacity in intestine of grass carp (Ctenopharyngodon idella). Aquac Nutr (epublication ahead of print version 10 January 2015).
63. Zhang, JX, Guo, LY, Feng, L, et al. (2013) Soybean β-conglycinin induces inflammation and oxidation and causes dysfunction of intestinal digestion and absorption in fish. PLOS ONE 8, e58115.
64. Enes, P, Pérez-Jiménez, A, Peres, H, et al. (2012) Oxidative status and gut morphology of white sea bream, Diplodus sargus fed soluble non-starch polysaccharide supplemented diets. Aquaculture 358–359, 7984.
65. Nagata, H, Takekoshi, S, Takagi, T, et al. (1999) Antioxidativeaction offlavonoids, quercetin and catechin, mediated by the activation of glutathione peroxidise. Tokai J Exp Clin Med 24, 111.
66. Martín, , Serrano, ABG, Ramos, S, et al. (2010) Cocoa flavonoids up-regulate antioxidant enzyme activity via the ERK1/2 pathway to protect against oxidative stress-induced apoptosis in HepG2 cells. J Nutr Biochem 21, 196205.
67. Procházková, D, Boušová, I & Wilhelmová, N (2011) Review: antioxidant and prooxidant properties of flavonoids. Fitoterapia 82, 513523.
68. Sitjà-Bobadilla, A, Peña-Llopis, S, Gómez-Requeni, P, et al. (2005) Effect of fish meal replacement by plant protein sources on non-specific defence mechanisms and oxidative stress in gilthead sea bream (Sparus aurata). Aquaculture 249, 387400.
69. Saera-Vila, A, Benedito-Palos, L, Sitjà-Bobadilla, A, et al. (2009) Assessment of the health and antioxidant trade-off in gilthead sea bream (Sparus aurata L.) fed alternative diets with low levels of contaminants. Aquaculture 296, 8795.
70. Olsvik, PA, Torstensen, BE & Berntssen, MHG (2007) Effects of complete replacementof fish oil with plant oil on gastrointestinal cell death, proliferation and transcription of eight genes’ encoding proteins responding to cellular stress in Atlantic salmon Salmo salar L. J. Fish Biol 71, 550568.
71. Morais, S, Silva, T, Carneiro, O, et al. (2012) Effects of genotype and dietary fish oil replacement with vegetable oil on the intestinal transcriptome and proteome of Atlantic salmon (Salmo salar). BMC Genomics 13, 448.
72. Trenzado, C, Hidalgo, MC, García-Gallego, M, et al. (2006) Antioxidant enzymes and lipid peroxidation in sturgeon Acipenser naccarii and trout Oncorhynchus mykiss. A comparative study. Aquaculture 254, 758767.