QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) Online Supporting Material Purchase Article View Shopping Cart Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Diez, A. Articles by Bautista, J. M. Search for Related Content PubMed PubMed Citation Articles by Diez, A. Articles by Bautista, J. M.

---

Biochemical, Molecular, and Genetic Mechanisms

Conjugated Linoleic Acid Affects Lipid Composition, Metabolism, and Gene Expression in Gilthead Sea Bream (Sparus aurata L)^1–3,

⁴ Departamento de Bioquímica y Biología Molecular IV, Universidad Complutense de Madrid, Facultad de Veterinaria, Madrid, Spain; ⁵ Departamento de Producción Animal, Universidad Politécnica de Madrid, ETS de Ingenieros Agrónomos, Ciudad Universitaria, Madrid, Spain; ⁶ Instituto de Acuicultura de Torre de la Sal, (CSIC), Ribera de Cabanes, Castellón, Spain; ⁷ Nutreco Aquaculture Research Centre AS (ARC), Stavanger, Norway; ⁸ National Agricultural Research Foundation, Fisheries Research Institute, Nea Paramos, Kavala, Greece; and ⁹ Institute of Aquaculture, University of Stirling, Stirling, UK

^* To whom correspondence should be addressed. E-mail: jmbau{at}vet.ucm.es.

	ABSTRACT

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
To maximize growth, farmed fish are fed high-fat diets, which can lead to high tissue lipid concentrations that have an impact on quality. The intake of conjugated linoleic acid (CLA) reduces body fat in mammals and this study was undertaken to determine the effects of dietary CLA on growth, composition, and postprandial metabolic variables in sea bream. Fish were fed 3 diets containing 48 g/100 g protein and 24 g/100 g fat, including fish oil supplemented with 0 (control), 2, or 4% CLA for 12 wk. Feed intake, specific growth rate, total body fat, and circulating somatolactin concentration were lower in fish fed CLA than in controls. Feed efficiency was greater in fish fed 2% CLA than in controls. Liver triglyceride concentrations were higher in fish fed 4% CLA and muscle triglyceride concentrations were lower in fish fed both CLA diets than in controls. Hepatic fatty acyl desaturase and elongase mRNA levels in fish fed CLA were lower than in controls. Metabolic differences between controls and CLA-fed fish were observed at 6 h but not at 24 h after the last meal, including lower postprandial circulating triglyceride concentrations, higher hepatic acyl-CoA-oxidase, and lower L-3-hydroxyacyl-CoA dehydrogenase activities in CLA-fed fish than in controls. Dietary CLA did not affect enzymes involved in lipogenesis including hepatic fatty acid synthase and malic enzyme, but it decreased glucose 6-phosphate dehydrogenase activity at 24 h, but not at 6 h after feeding. The data suggest that CLA intake in sea bream has little effect on hepatic lipogenesis, channels dietary lipid from adipose tissue to the liver, and switches hepatic mitochondrial to peroxisomal ß-oxidation.

	Introduction

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

Conjugated linoleic acid (CLA) is a term used to describe positional and geometric isomers of linoleic acid [18:2(n-6)], the 2 main naturally occurring isomers being cis-9, trans-11 and trans-10, cis-12 (4). These compounds occur mainly in beef and dairy products but are widespread in lower levels in many foodstuffs (4). The dietary inclusion of CLA can cause significant alterations in energy and lipid metabolism in mammals leading to reductions in overall body fat mass and is thought to be beneficial in several farmed species and animal disease models and, by extension, in humans (5). CLA has also been shown to alter HUFA biosynthesis in cell models (6) and to increase the expression of genes involved in the HUFA biosynthetic pathway (7). The effects of dietary CLA have also been examined in several fish species (8–11) and, in general, effects on growth performance or on lipid metabolism have not been observed. As for land animals (12), the effects of CLA could be isomer-, dose-, time-, and/or species-dependent. The aim of the present study was to investigate the effects of dietary CLA, provided as equal amounts of the cis-9, trans-11 and trans-10, cis-12 isomers, on lipid metabolism, HUFA biosynthesis and lipid composition in the liver and flesh of sea bream.

	Materials and Methods

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
    Fish and experimental diets. The in vivo feeding trial was conducted over the period of July 25 to October 16 (2005) in an indoor system (Instituto de Acuicultura de Torre de la Sal) fed with running seawater. The oxygen content of the outlet water was always higher than 85% saturated. Day and night cycles and water temperatures (17°C to 25°C) were dictated by natural conditions. Juvenile sea bream, obtained from a commercial hatchery (Cupimar), were graded by size and checked for the absence of anatomical malformations. After 20 d of acclimatization to the experimental facilities, groups of 90 fish were placed in 12 circular glass fiber tanks (500 L), with 4 replicates set up for each dietary treatment. Fish were fed once a day (0900 h), 6 d/wk. The diets (Table 1) were formulated to contain 24 g/100 g fat (see Supplemental Table 1 for fatty acid composition), 48 g/100 g protein as proportions of the total mass, and prepared at the Skretting Aquaculture Research Centre (Stavanger, Norway).

    Sampling protocol. Whole body composition (proximate and fatty acid composition) was determined in a pooled sample of 10 fingerlings at the start of the trial and in pools of 5 fish per tank at the end of the trial (at 24 h). Specimens for whole body analysis were ground, and small aliquots dried to estimate water content. The remaining samples were freeze-dried and stored at –80°C until analysis. To assess dietary effects on postprandial and basal metabolism, 12 fish per dietary treatment (i.e., 3 fish per tank) were sampled at 6 h and 24 h after last feeding. Fish were anesthetized and killed with MS 222 (1 g/10 L), and blood samples were taken for plasma metabolite analysis. Specimens of liver, white muscle, intestine, and perivisceral adipose tissue for biochemical and gene expression analyses were rapidly excised, frozen in liquid nitrogen, and stored at –80°C until analysis. Tissues for gene expression analyses were obtained from individual fish (1–3 fish/tank, 3–6 fish/dietary treatment at the 6-h and 24-h time points).

All procedures were carried out according to national and institutional regulations (Consejo Superior de Investigaciones Científicas Instituto de Acuicultura de Torre de la Sal Review Board) and the current European Union legislation on handling experimental animals.

    Proximate analyses, lipid extraction, and fatty acid analysis. Moisture and crude protein, lipid, and ash contents of fish whole bodies were determined according to AOAC procedures (13). The liver and flesh (skinned and deboned) of 3 fish were mixed to generate pooled homogenates. Total lipid was extracted using chloroform/methanol (2:1, v:v) and prepared according to the method of Folch et al. (14). The weight of lipid was determined gravimetrically after evaporation of the solvent and overnight desiccation in vacuo. Lipids were separated into classes by high-performance thin-layer chromatography (HPTLC), visualized by charring for 15 min at 160°C after spraying with 3% (w:v) aqueous copper acetate containing 8% (v:v) phosphoric acid and quantified by densitometry using a Camag 3TLC scanner and winCATS software (15). Individual lipid classes were confirmed by comparison with Rf values for authentic standards run alongside samples on HPTLC plates developed in the above solvent systems.

Fatty acid methyl esters (FAME) were prepared from total tissue lipid and purified according to the methods of Christie (16) and Ghioni et al. (17). FAME were separated and quantified by GLC (17,18). Data were collected and processed using Chromcard software for Windows, version 1.19 (Thermoquest Italia S.p.A.).

    Key lipogenic and ß-oxidation enzyme activities. Activities of glucose-6-phosphate dehydrogenase (G6PD; EC 1.1.1.49), malic enzyme (EC 1.1.1.40), and fatty acid synthetase (EC 2.3.1.38) were assayed as previously described (19–21) in liver cytoplasm extracts. Acyl-CoA oxidase (ACO; EC 1.3.99.3) in peroxisome-enriched liver fractions was determined according to previously described procedures (22) with the modifications of Ruyter et al. (23). Mitochondrial extracts from livers were prepared according to the method of Harper and Saggerson (24), and L-3-hydroxyacyl-CoA dehydrogenase activity (L3HOAD; EC 1.1.135) was measured using the assay described by Bradshaw and Noyes (25). All enzyme assays were performed in duplicate or triplicate. Enzyme activity units (IU), defined as µmol of substrate converted to product per minute at the assay temperature, were expressed per mg of soluble protein (specific activity). Protein was determined by the Bio-Rad dye reagent method using bovine serum albumin as the standard. Total protein contents, determined in the mitochondrial extracts and peroxisomal enriched fractions, did not differ among experimental groups.

    Plasma metabolite and hormone concentrations. Plasma glucose, cholesterol (CHOL), and triglyceride (TAG) concentrations were measured spectrophotometrically using commercial kits (Sigma, Cat. 315–310, 401–25P, and 337-B, respectively). Plasma growth hormone (GH) and somatolactin (SL) concentrations were measured by homologous radioimmunoassays (RIA) as previously described (26,27). The midrange of the assay was 1.8 µg/L for GH and 2.1 µg/L for SL. Plasma insulin-like growth factor (IGF-I) was extracted by ethanol-cryoprecipitation and measured by fish RIA. The gilthead sea bream assay was based on the use of bream (Pagrus auratus) IGF-I (GroPep: 5PAF-AGU100) as tracer and standard. Anti-barramundi (Lates calcarifer) IGF-I serum (GroPep: 5PAF1-YU100) (1:8000) was used as primary antibody. A goat anti-rabbit IgG (1:20) (Biogenesis: 5196–2104) was used as a precipitating antibody. The sensitivity and midrange of the assay were 0.05 and 0.7–0.8 µg/L, respectively.

    RNA isolation and real-time quantitative RT-PCR. Total RNA was extracted from the fish tissue using an automated nucleic acid isolation system (ABI PRISM 6100 Nucleic Acid Prep Station, Applied Biosystems) according to the manufacturer's instructions. Total RNA was quantified with RIBOGreen TM (Molecular Probes, Europe) using a Perkin-Elmer LS-50B fluorimeter, and RNA integrity was checked by electrophoresis in 2% agarose gels. First strand cDNA was synthesized using the High-Capacity cDNA Archive kit (Applied Biosystems) according to the manufacturer's instructions.

Relative abundance of peroxisome proliferator acitvated receptor (PPAR)-mRNA was assessed using the 5' fluorogenic nuclease assay (TaqMan) in an ABI Prism7000 Sequence Detector System (Applied Biosystems) using primers and protocols described elsewhere (28). All samples were analyzed in triplicate and quantified by normalizing the PPAR signal to that of -tubulin by the 2^–Ct method (28).

    Riboprobes and ribonuclease protection assay. For the sea bream -6 fatty acyl desaturase (DES) riboprobe, oligonucleotide primers 5'-GAC CAT GCA GTT ACA AGC CAC C and 5'-TCC CCT GAG TTC TTC AGT GAC C were used in the PCR amplification of a 216 bp fragment (nucleotides 1225–1441) of cDNA [Genbank AY055749, (29)]. For the fatty acyl elongase (ELO) riboprobe, oligonucleotide primers 5'-TGC CAG GAC ACT CAC AGT GC and 5'-GGA CGA AGC TGT TTA GGG AGG were used in the PCR amplification of a 226 bp fragment (nucleotides 303–529) of cDNA [Genbank AY660879, (30)]. The ribonuclease protection assay was performed as previously described (28) and the relative expression of genes were compared among individual fish and treatments by normalizing to ß-actin expression (28).

    Statistical analysis. Results are expressed as means ± SD. Data were analyzed as a randomized design with type of diet as the main source of variation using the General Linear Model procedure of the SAS computer package (SAS Institute). For the plasma metabolite (glucose, TAG, and CHOL), tissue enzyme, and gene expression analyses, data were analyzed according to a factorial arrangement of treatments with type of diet, sampling time, and their interactions tested. The percentage of fatty acid data not showing homogeneity of variance were arcsine square-root transformed before analysis. Significant differences between treatments were assessed by the Newman-Keul's multiple comparison procedure. The level of significance was set at P < 0.05.

	Results

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
    Growth, feeding, and lipid contents. Differences among the groups in growth, feeding, and lipid contents were determined from samples taken at the start of the trial and at the end of the trial after 12 wk. Compared with control fish, either 2 or 4% CLA supplementation led to reduced feed intake, final body and liver weights, specific growth rates, and whole body lipid concentration and retention (Table 2). These effects were accompanied by greater feed efficiency in fish fed the 2% CLA diet.

    Plasma biochemistry. Plasma, enzyme, and endocrine variables were measured in samples taken at 6 h and 24 h after last feeding. The plasma glucose concentration was unaffected by dietary treatment but decreased from 6 h to 24 h postprandially, irrespective of diet (Supplemental Table 3). The plasma TAG concentration also decreased over the course of the postprandial period, with the CLA diets leading to reduced circulating TAG levels relative to controls 6 h after feeding. The diets did not affect the plasma CHOL concentration, although it was significantly lower after 24 h than after 6 h.

The activity of the lipogenic enzyme G6PD was higher in the 6 h postfeed liver samples than in 24 h samples in all groups, which did not differ from one another 6 h after the final feeding (Fig. 1A). However, G6PD activity was lower in fish fed both CLA diets 24 h after feeding compared with controls. Diet and time of sampling did not affect the specific activity of liver fatty acid synthase or malic enzyme (data not shown), other indicators of lipogenic activity. In contrast, dietary CLA affected lipolytic enzymes. In fish fed the control diet, the activity of hepatic ACO, the rate-limiting enzyme in peroxisomal fatty acid ß-oxidation, was the same at 6 h as at 24 h (Fig. 1B). However, in fish fed 4% CLA, ACO activity at 6 h after feeding was 2.8-fold that of controls (P < 0.05). At 24 h after feeding, ACO activity did not differ among the groups (P < 0.05 for the interaction). Liver L3HOAD activity, a marker of mitochondrial ß-oxidation, exhibited a large decrease from 6 h to 24 h postfeeding in fish fed the control diet (Fig. 1C). Similar to the results for ACO, CLA did not affect L3HOAD activity at 24 h after feeding, but in contrast to ACO, CLA-fed fish had lower activity 6 h after feeding compared with controls.

View larger version (23K): [in this window] [in a new window]   FIGURE 1  Hepatic glucose-6-phosphate dehydrogenase (A), acyl-CoA oxidase (B) and L-3-hydroxyacyl-CoA dehydrogenase (C) activities in gilthead sea bream after being fed diets containing 0, 2, or 4% CLA for 12 wk. Values are means ± SD, n = 6. Means in a panel for a time point without a common letter differ, P < 0.05. Interaction between type of diet and sampling (B, C), P < 0.05.

    DES and ELO gene expression. Levels of ELO and DES mRNA, as assessed by the ribonuclease protection assay, revealed that the gene coding for these enzymes were differentially expressed in the tissues examined (Fig. 2A). We detected ELO expression in both the liver and intestine but not in white muscle or adipose tissue, whereas DES seemed to be liver specific. We therefore limited our analysis of diet-dependent gene expression to these tissues using specimens obtained 24 h after the end of the trial.

View larger version (29K): [in this window] [in a new window]   FIGURE 2  mRNA expression of potential PPAR target genes in gilthead sea bream after being fed diets containing 0, 2, or 4% CLA for 12 wk. Fatty acid elongase (ELO) and desaturase (DES) in liver [L], intestine [I], white muscle [M], and adipose tissue [A] (A); mRNA expression in response to the 3 dietary treatments is shown in panels (B, C, and D). The mRNA abundance range in fish fed the control diet was set at 100. All values are means ± SD, n = 3. Means in a panel without a common letter differ, P < 0.05.

    PPAR mRNA expression in fish tissues. Messenger RNA expression for the PPAR isotypes was determined in the muscle, liver, and adipose tissue of fish specimens collected 6 h and 24 h after the last feeding (Fig. 3). In muscle, PPAR or PPARß expression did not differ at 6 h or 24 h after the final feeding in fish fed the control diet (Fig. 3A upper and middle). However, at 24 h after feeding, mRNA expression of both subtypes was higher in both groups of CLA-fed fish than in controls and was also greater at 6 h after feeding in fish fed the 4% CLA diet (Fig. 3A, upper and middle). Muscle PPAR expression did not differ between controls and fish fed 4% CLA at either time point but expression was greater in the 2% CLA-fed group (P < 0.05) after 6 h than in both the controls and the 4% CLA group (Fig. 3A, lower).

View larger version (32K): [in this window] [in a new window]   FIGURE 3  Effects on mRNA levels of PPAR, ß and isotypes in gilthead sea bream after being fed diets containing 0, 2, or 4% CLA for 12wk in muscle (A), liver (B) and adipose (C) (mesenteral) tissues. Values are means ± SD, n = 6. Means in a panel without a common letter differ, P < 0.05. Asterisks in (A, upper and middle) and (B, upper, middle and lower) indicate interaction between type of diet and sampling time, P < 0.05.

In adipose tissue, the 3 PPAR isotypes did not change at 6 h and 24 h after feeding in fish fed the control diet, and, at 6 h, the groups did not differ from one another in expression levels of PPARß and PPAR (Fig. 3C). However, PPAR expression was increased 6 h after feeding compared with 24 h after feeding in fish fed 2% CLA (Fig. 3C, upper), whereas PPAR mRNA levels were lower at 6 h compared with 24 h after feeding in fish fed 4% CLA (Fig. 3C, lower). In contrast, at 24 h after feeding, dietary CLA reduced PPARß levels and increased PPAR levels compared with controls (Fig. 3C, middle and lower).

	Discussion

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
The dietary intake of CLA in mammals affects body weight and fat deposition by reducing body fat, increasing lean body mass, and lowering serum lipids (5). The mechanisms proposed for such changes include the modification of gene expression and physiology of the tissues that play a crucial role in lipid homeostasis, such as adipose tissue and liver. In this study, sea bream juveniles were fed a commercial diet containing 24% fat, supplemented with up to 4% CLA (1:1 mixture of c9,t11 and t10,c12), for 12 wk to assess the effects of including CLA in aquaculture diets on growth, lipid deposition, and metabolism. Our results indicate that the inclusion of CLA in the feed promotes significant growth of the fish along with both metabolic and gene expression changes. These effects, however, were accompanied by a reduction in feed intake, and it could be argued that this also accounted for the reductions in growth and in whole body lipid contents and retention. However, there were no apparent CLA-linked changes in conditioning factors such as relative visceral or liver weights or total protein retention, and we would expect to see reductions in these factors in sea bream as a response to diminished ration size (31). Moreover, decreasing the feed ration in sea bream also has well-documented effects on circulating GH and IGF-I levels (32), neither of which were affected by CLA-feeding. In contrast, levels of SL, an emerging marker of adiposity regardless of feed intake (33), were reduced in response to CLA intake by the sea bream. Taken together, these results suggest that CLA had no effects on overall growth physiology and had small but significant effects on feed intake. Indeed, the few pair-feed trials involving dietary CLA performed on mammals have also revealed that a decreased feed intake cannot solely account for fat mass reductions or several other metabolic effects (34,35). It is clear, however, that the dietary CLA levels used here have no beneficial effects on the growth performance of sea bream. Previous studies in fish have demonstrated a similar lack of benefit (11), although, as also observed with sea bream, an increased feed efficiency has been reported (8).

The fish tissue buildup of dietary CLA varies according to species, dietary lipid source, CLA inclusion level, and fish size (8–11). In our study, juvenile gilthead sea bream incorporated both CLA isomers (c9, t11, and t10, c12) in liver and muscle. The concentration of a given fatty acid in the fish tissue, divided by its concentration in the diet, provides a deposition ratio (R_D) value that serves to assess fatty acid composition changes in response to diet (10). Accordingly, CLA was deposited at a lower rate than expected considering the dietary amount (R_D < 1). This deposition ratio was higher for muscle (R_D 0.70) than liver (R_D 0.50) with no effects observed from the dietary CLA level, as previously reported for Atlantic salmon (10). Notably, an accumulation of CLA in muscle was associated with increases in total phospholipids, particularly PE and PS, indicating that CLA may be incorporated into structural membrane lipid. In both muscle and liver, the incorporation of CLA led to decreased total MUFA content and to increased 18:0 concentrations in muscle. This phenomenon is seen both in mammals (12) and in several fish species (8–11) and seems to be among the most consistent effects of CLA. The main MUFA, 16:1(n-7) and 18:1(n-9), are synthesized through stearoyl-CoA desaturase-1 and, in mammals, the above effects have been attributed to the suppression or direct inhibition of SCD (6). The effects observed here in sea bream could thus involve similar mechanisms.

In addition to the changes recorded in MUFA and saturated fatty acid, we detected lowered total hepatic 20:5(n-3) levels, and raised levels of the precursor fatty acid 18:3(n-3) following CLA intake. These changes can be directly related to the reductions observed in hepatic DES and ELO expression, the genes responsible for HUFA biosynthesis. Diminished HUFA biosynthesis would not be desirable in aquaculture, particularly in diets in which dietary fish oils rich in HUFA have been replaced with HUFA-deficient plant oils.

It should be emphasized that some of the most significant CLA-induced changes observed 6 h after last feeding were not evident 24 h after feeding. Thus, the careful design of experiments and sampling procedures will provide greater insight into the varied effects of CLA. A clear effect in sea bream was an increased liver TAG concentration following the 4% CLA diet and a decreased postprandial plasma TAG for both CLA diets. Moreover, TAG levels in muscle and total body lipids were also lowered in response to the CLA diets. These effects were not associated with augmented liver lipogenesis and there was even some evidence of decreased lipogenesis in this tissue. Thus, it would seem that TAG derived from dietary sources are being diverted to the liver rather than to muscle for fuel or to adipose tissue for storage. Further, the increase in liver TAG was related to major changes in ß-oxidation pathways, as shown by the switch from postprandial mitochondrial metabolism to the peroxisomal pathway indicated by changes in L3HOAD and ACO activity, respectively. In other species, CLA seems to have different effects on liver TAG levels. For example, CLA increases liver TAG in mice but decreases CLA in hamsters (36,37). Nevertheless, as observed here, several studies in rodents have indicated enhanced peroxisomal metabolism (38) suggesting that this may be a common effect of CLA feeding across species. Peroxisomal ß-oxidation serves to metabolize atypical fatty acids, such as branched-chain or long-chain molecules that are structurally incapable of undergoing mitochondrial metabolism (39). Studies on CLA-fed rats have detected significant amounts of probable peroxisomal metabolites of CLA (40).

Although the mechanisms of the effects of CLA on lipid metabolism are unclear, there is increasing evidence for the involvement of PPAR-dependent gene regulation (41–43). PPAR are ligand-dependent transcription factors known to play critical roles in regulating lipid homeostasis in tissues such as liver, muscle, and adipose. These proteins act by regulating the activity of numerous genes involved in fatty acid storage, uptake, and metabolism (44). Studies in PPAR-null mice have shown that most of the effects of CLA on the genes involved in mitochondrial ß-oxidation are independent of PPAR, whereas CLA increases peroxisomal ACO via a PPAR-dependent mechanism (42). More recently, it has been suggested that the lipid-lowering effects of CLA are attributable to the diminished expression of PPAR and many of its downstream target genes (43). Sea bream PPAR isotype mRNAs are differentially expressed in a range of tissues (28). Thus, in muscle, PPAR and PPARß are the isotypes predominantly expressed, whereas PPAR expression is dominant in adipose tissue. All 3 isotypes are expressed in liver. CLA is a highly effective activator of sea bream PPAR and, to a lesser extent, of PPARß (28). In the present study, mRNA levels of PPAR were affected by dietary CLA inclusion, particularly in muscle in which basal levels of both PPAR and PPARß increased. The levels of PPAR in liver showed a postprandial decrease, confirming the results of previous studies (28), yet were not much affected by CLA feeding, indicating that PPAR-dependent gene regulation may not be affected by CLA. Interestingly, PPARß levels in liver were postprandially reduced by CLA, coinciding with the observed reduction in mitochondrial ß-oxidation capacity. In addition, CLA led to the downregulated postprandial expression in the liver and upregulated basal expression in adipose tissue of PPAR. Given the proposed functions of PPAR in regulating fat deposition (44), the observed decreases in whole body fat and increases in liver fat in sea bream do not correlate with the CLA-induced expression profile of PPAR. However, it should be noted that gene expression levels do not necessarily bear a relation to functional protein levels. Our data, therefore, do not provide sufficient evidence that the effects of CLA observed in sea bream are mediated through PPAR. In addition, the greater complexity of PPAR biology in fish compared with mammals must also be considered in view of the presence of multiple PPAR isoforms in fish species, which was previously inferred (28). Hence, it is possible that PPAR isoforms, in addition to the ones assayed here, may be functionally expressed in sea bream and more directly involved in mediating CLA effects.

In summary, the lipid-lowering effect of CLA observed here in sea bream juveniles may be the combined result of the following factors: a reduced feed intake, the endocrine status of the fish, the diversion of dietary-derived TAG from muscle and adipose tissue to liver, and increased hepatic peroxisomal ß oxidation. This induction of peroxisomal ß oxidation suggests a subtoxic response to CLA, which may have unknown consequences for fish health. Thus, despite promoting reduced adiposity, the slightly negative growth effects and reduction in HUFA biosynthesis indicate that the inclusion of CLA in aquaculture diets would be of little benefit.

Accordingly, dietary CLA supplementation would not be of obvious interest in intensive sea bream farming. However, its effects on fish physiology at the different developmental stages needs to be further evaluated, insofar as fish fed CLA may offer potential benefits for humans by combining the lipid-lowering properties of CLA with the beneficial health effects of high levels of HUFA that naturally occur in fish.

	ACKNOWLEDGMENTS

 
The authors thank Mr. Guillermo Borés, DVM, for his help and advice, and for organizing the in vivo growth trial.

	FOOTNOTES

 
¹ Supported by the European Commission (Q5RS-2000–30360) and Spanish MEC (AGL2004-06319-C02-01). Some of the analyses performed were supported by a grant from the Access to Research Infrastructure Action of the Improving Human Potential Program of the European Commission (contract HPRI-CT-2001-00180 to A.D. and J.M.B.).

² Author disclosures: A. Diez, D. Menoyo, S. Pérez-Benavente, J. A. Calduch-Giner, S. Vega-Rubin de Celis, A. Obach, L. Favre-Krey, E. Boukouvala, M. J. Leaver, D. R. Tocher, J. Pérez-Sanchez, G. Krey, and J. M. Bautista, no conflicts of interest.

³ Supplemental Tables 1–3 are available with the online posting of this paper at jn.nutrition.org.

¹⁰ Abbreviations used: ACO, acyl-CoA oxidase; CHOL, cholesterol; CLA, conjugated linoleic acid; DES, 6-fatty acyl desaturase; ELO, fatty acyl elongase; G6PD, glucose 6-phosphate dehydrogenase; GH, growth hormone; HUFA, (n-3) highly unsaturated fatty acids; IGF-I, insulin-like growth factor; L3HOAD, L-3-hydroxyacyl-CoA dehydrogenase; ME, malic enzyme; PE, phosphatidylethanolamine; PPAR, peroxisome proliferator acitvated receptors; SL, somatolactin; TAG, triglyceride.

Manuscript received 15 October 2006. Initial review completed 2 November 2006. Revision accepted 28 March 2007.

	LITERATURE CITED

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 

1. Tocher DR. Metabolism and functions of lipids and fatty acids in teleost fish. Rev Fish Sci. 2003;11:107–84.

2. Naylor RL, Goldburg RJ, Primavera JH, Kautsky N, Beveridge MCM, Clay J, Folke C, Lubchenco J, Mooney H, Troell M. Effect of aquaculture on world fish supplies. Nature. 2000;405:1017–24.[Medline]

3. Bell G, Torstensen B, Sargent J. Replacement of marine fish oils with vegetable oils in feeds for farmed salmon. Lipid Technol. 2005;17:7–11.

4. Pariza MW, Park Y, Cook M. The biologically active isomers of conjugated linoleic acid. Prog Lipid Res. 2001;40:283–98.[Medline]

5. Belury MA. Dietary conjugated linoleic acid in health: physiological effects and mechanisms of action. Annu Rev Nutr. 2002;22:505–31.[Medline]

6. Eder K, Slomma N, Becker K. Trans-10,cis-12 conjugated linoleic acid suppresses the desaturation of linoleic and -linolenic acids in HepG2 cells. J Nutr. 2002;132:1115–21.[Abstract/Free Full Text]

7. Takahashi Y, Kushiro M, Shinohara K, Ide T. Activity and mRNA levels of enzymes involved in hepatic fatty acid synthesis and oxidation in mice fed conjugated linoleic acid. Biochim Biophys Acta. 2003;1631:265–73.[Medline]

8. Twibell RG, Watkins BA, Rogers L, Brown PB. Effects of dietary conjugated linoleic acids on hepatic and muscle lipids in hybrid striped bass. Lipids. 2000;35:155–61.[Medline]

9. Twibell RG, Watkins BA, Brown PB. Dietary conjugated linoleic acids and lipid source alter fatty acid composition of juvenile yellow perch Perca flavescens. J Nutr. 2001;131:2322–8.[Abstract/Free Full Text]

10. Berge GM, Ruyter B, Asgard T. Conjugated linoleic acid in diets for juvenile Atlantic salmon (Salmo salar), effects on fish performance, proximate composition, fatty acid and mineral content. Aquaculture. 2004;237:365–80.

11. Kennedy SR, Campbell PJ, Porter A, Tocher DR. Influence of dietary conjugated linoleic acid (CLA) on lipid and fatty acid composition in liver and flesh of Atlantic salmon (Salmo salar). Comp Biochem Physiol B Biochem Mol Biol. 2005;141:168–78.[Medline]

12. Evans ME, Brown JM, McIntosh MK. Isomer-specific effects of conjugated linoleic acid (CLA) on adiposity and lipid metabolism. J Nutr Biochem. 2002;13:508–16.[Medline]

13. Association of Official Analytical Chemists. Official methods of analysis, vol. 1, 15th ed. Washington, DC: AOAC; 1990.

14. Folch J, Lees M, Sloane-Stanley GH. A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem. 1957;226:497–509.[Free Full Text]

15. Henderson RJ, Tocher DR. Thin-layer chromatography. In: Hamilton RJ, Hamilton S., editors. Lipid analysis: a practical approach. Oxford: Oxford University Press; 1992: 65–111.

16. Christie WW. Lipid analysis, 2nd ed. Oxford: Pergamon Press; 1982: 207.

17. Ghioni C, Bell JG, Sargent JR. Polyunsaturated fatty acids in neutral lipids and phospholipids of some freshwater insects. Comp Biochem Physiol. 1996;114B:161–70.

18. Ackman RG. Fish lipids, part 1. In: Connell JJ, editor. Advances in fish sciences and technology. Farnham, UK: Fishing News Books; 1980: 86–103.

19. Bautista JM, Garrido-Pertierra A, Soler G. Glucose-6-phosphate dehydrogenase from Dicentrarchus labrax liver: kinetic mechanism and kinetics of NADPH inhibition. Biochim Biophys Acta. 1988;967:354–63.[Medline]

20. Alvarez MJ, López-Bote CJ, Diez A, Corraze G, Arzel J, Dias J, Kaushik SJ, Bautista JM. 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. 1998;80:281–9.[Medline]

21. Alvarez MJ, Diez A, López-Bote CJ, Gallego M, Bautista JM. Short-term modulation of lipogenesis by macronutrients in rainbow trout (Oncorhynchus mykiss) hepatocytes. Br J Nutr. 2000;84:619–28.[Medline]

22. Small GM, Burdett K, Connock MJ. A sensitive spectrophotometric assay for peroxisomal Acyl-Coa oxidase. Biochem J. 1985;227:205–10.[Medline]

23. Ruyter B, Andersen Ø, Dehli A, Östlund-Farrants AK, Gjøen T, Thomassen MS. Peroxisome proliferator activated receptor in Atlantic salmon (Salmo salar): Effect on PPARs transcription and acyl-Coa Oxidase activity in hepatocytes by peroxisome proliferators and fatty acid. Biochim Biophys Acta. 1997;1348:331–8.[Medline]

24. Harper RD, Saggerson ED. Some aspects of fatty acid oxidation in isolated fat-cell mitochondria from rat. Biochem J. 1975;152:485–94.[Medline]

25. Bradshaw RA, Noyes BE. L-3-hydroxyacyl coenzyme A dehydrogenase from pig heart muscle. EC 1.1.1.35 L-3-hydroxyacyl-CoA: NAD oxidoreductase. Methods Enzymol. 1975;35:122–8.[Medline]

26. Martinez-Barbera JP, Pendon C, Marti-Palanca H, Calduch-Giner JA, Rodriguez RB, Valdivia MM, Perez-Sanchez J. The use of recombinant gilthead sea bream (Sparus-auratus) growth-hormone for radioiodination and standard preparation in radioimmunoassay. Comp Biochem Physiol A Physiol. 1995;110:335–40.[Medline]

27. Calduch-Giner JA, Pendon C, Valdivia MM, Pérez-Sánchez J. Recombinant somatolactin as a stable and bioactive protein in a cell culture bioassay: development and validation of a sensitive and reproducible radioimmunoassay. J Endocrinol. 1998;156:441–7.[Abstract]

28. Leaver MJ, Boukouvala E, Antonopoulou E, Diez A, Favre-Krey L, Tariq Ezaz M, Bautista JM, Tocher DR, Krey G. Three peroxisome proliferator-activated receptor isotypes from each of two species of marine fish. Endocrinology. 2005;146:3150–62.[Abstract/Free Full Text]

29. Seiliez I, Panserat S, Corraze G, Kaushik S, Bergot P. Cloning and nutritional regulation of a Delta6-desaturase-like enzyme in the marine teleost gilthead seabream (Sparus aurata). Comp Biochem Physiol B Biochem Mol Biol. 2003;135:449–60.[Medline]

30. Agaba MK, Tocher DR, Dickson CA, Zheng X, Dick JR, Teale AJ. Cloning and functional characterisation of polyunsaturated fatty acid elongases from marine and freshwater teleost fish. Comp Biochem Physiol Biochem Mol Biol. 2005;142:342–52.

31. Company R, Calduch-Giner JA, Kaushik S, Perez-Sanchez J. Growth performance and adiposity in gilthead sea bream (Sparus aurata): risks and benefits of high energy diets. Aquaculture. 1999;171:279–92.

32. Company R, Astola A, Pendon C, Valdivia MM, Pérez-Sánchez J. Somatotropic regulation of fish growth and adiposity: growth hormone (GH) and somatolactin (SL) relationship. Comp Biochem Physiol C Toxicol Pharmacol. 2001;130:435–45.[Medline]

33. Mingarro M, Vega-Rubin de Celis S, Astola A, Pendón C, Valdivia MM, Pérez-Sánchez J. Endocrine mediators of seasonal growth in gilthead sea bream (Sparus aurata): the growth hormone and somatolactin paradigm. Gen Comp Endocrinol. 2002;128:102–11.[Medline]

34. Hargrave KM, Li C, Meyer BJ, Kachman SD, Hartzell DL, Della-Fera MA, Miner JL, Baile CA. Adipose depletion and apoptosis induced by trans-10, cis-12 conjugated linoleic acid in mice. Obes Res. 2002;10:1284–90.[Medline]

35. Ntambi JM, Choi Y, Park Y, Peters JM, Pariza MW. Effects of conjugated linoleic acid (CLA) on immune responses, body composition and stearoyl-CoA desaturase. Can J Appl Physiol. 2002;27:617–27.[Medline]

36. Zabala A, Churruca I, Macarulla MT, Rodriguez VM, Fernandez-Quintela A, Martinez JA, Portillo MP. The trans-10,cis-12 isomer of conjugated linoleic acid reduces hepatic triacylglycerol content without affecting lipogenic enzymes in hamsters. Br J Nutr. 2004;92:383–9.[Medline]

37. Javadi M, Beynen AC, Hovenier R, Lankhorst AE, Lemmens AG, Terpstra AHM, Geelen MJH. Prolonged feeding of mice with conjugated linoleic acid increases hepatic fatty acid synthesis relative to oxidation. J Nutr Biochem. 2004;15:680–7.[Medline]

38. Degrace P, Demizieux L, Gresti J, Chardigny JM, Sebedio JL, Clouet P. Hepatic steatosis is not due to impaired fatty acid oxidation capacities in C57BL/6J mice fed the conjugated trans-10,cis-12-isomer of linoleic acid. J Nutr. 2004;134:861–7.[Abstract/Free Full Text]

39. Reddy JK, Mannaerts GP. Peroxisomal lipid metabolism. Annu Rev Nutr. 1994;14:343–70.[Medline]

40. Banni S, Petroni A, Blasevich M, Carta G, Cordeddu L, Murru E, Melis MP, Mahon A, Belury MA. Conjugated linleic acids (CLA) as precursors of a distinct family of PUFA. Lipids. 2004;39:1143–6.[Medline]

41. Moya-Camarena SY, Van den Heuvel JP, Belury MA. Conjugated linoleic acid activates peroxisome proliferator-activated receptor and ß subtypes but does not induce hepatic peroxisome proliferation in Sprague-Dawley rats. Biochim Biophys Acta. 1999;1436:331–42.[Medline]

42. Peters JM, Park Y, Gonzalez FJ, Pariza MW. Influence of conjugated linoleic acid on body composition and target gene expression in peroxisome proliferators-activated receptor -null mice. Biochim Biophys Acta. 2001;1533:233–42.[Medline]

43. Brown JM, Sandberg M, Boysen S, Chung O, Fabiyi RF, Morrison S, Mandrup M. McIntosh. Conjugated linoleic acid (CLA) induces human adipocyte delipidation: autocrine/paracrine regulation of MEK/ERK signaling by adipocytokines. J Biol Chem. 2004;279:26735–47.[Abstract/Free Full Text]

44. Desvergne B, Wahli W. Peroxisome proliferator-activated receptors: nuclear control of metabolism. Endocr Rev. 1999;20:649–88.[Abstract/Free Full Text]

This Article Abstract Full Text (PDF) Online Supporting Material Purchase Article View Shopping Cart Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Diez, A. Articles by Bautista, J. M. Search for Related Content PubMed PubMed Citation Articles by Diez, A. Articles by Bautista, J. M.