Caffeic acid and hydroxytyrosol have anti-obesogenic properties in zebrafish and rainbow trout models

Some natural products, known sources of bioactive compounds with a wide range of properties, may have therapeutic values in human health and diseases, as well as agronomic applications. The effect of three compounds of plant origin with well-known dietary antioxidant properties, astaxanthin (ATX), caffeic acid (CA) and hydroxytyrosol (HT), on zebrafish (Danio rerio) larval adiposity and rainbow trout (Onchorynchus mykiss) adipocytes was assessed. The zebrafish obesogenic test (ZOT) demonstrated the anti-obesogenic activity of CA and HT. These compounds were able to counteract the obesogenic effect produced by the peroxisome proliferator-activated receptor gamma (PPARγ) agonist, rosiglitazone (RGZ). CA and HT suppressed RGZ-increased PPARγ protein expression and lipid accumulation in primary-cultured rainbow trout adipocytes. HT also significantly reduced plasma triacylglycerol concentrations, as well as mRNA levels of the fasn adipogenic gene in the adipose tissue of HT-injected rainbow trout. In conclusion, in vitro and in vivo approaches demonstrated the anti-obesogenic potential of CA and HT on teleost fish models that may be relevant for studying their molecular mode of action. Further studies are required to evaluate the effect of these bioactive components as food supplements for modulating adiposity in farmed fish.


Introduction
Obesity has become a worldwide epidemic and is considered one of the most serious public health problems of our time [1,2]. Overweight and obesity occur when energy intake exceeds energy expenditure, leading to increased storage of triacylglycerols (TAG), mainly in white adipose tissue (WAT). In addition to TAG storage in adipocyte lipid droplets, WAT has been recognized as a multi-functional endocrine organ that plays a critical role in modulating several physiological processes, such as appetite, whole-body energy metabolism and homeostasis, as well as tissue inflammation responses [3]. Consequently Forêt), under permit number A33-522-6. All experiments were conducted in conformity with the European Communities Council Directive (2010/63/EU) on the protection of animals used for scientific purposes and local French legislation on the care and use of laboratory animals. Larvae were obtained by natural mating and raised in embryo water (90 μg/ml Instant Ocean [Aquarium Systems, Sarrebourg, France], 0.58 mM CaSO 4 , 2H 2 O, dissolved in reverseosmosis purified water) at 28.5˚C with an 11L:13D photoperiod. From 5 days postfertilization until day 15, larvae were fed ad libitum on ZF Biolabs formulated diet flakes (Tres Cantos, Spain). They were then nourished with standard diet (SD) for late larvae (TetraMin Baby, Tetra GmbH, Melle, Germany). Animal stages were recorded according to standard length, i.e. the distance from the rostral tip of the larva to the base of the caudal fin. Juvenile rainbow trout, body weight approximately 80 g for in vivo studies and 250 g for extracting WAT to be used in adipocyte primary cultures, were obtained from the "Viveros de los Pirineos" fish farm (El Grado, Huesca, Spain). Animals were maintained according to the

Zebrafish obesogenic test (ZOT)
The short-term ZOT assay, using Nile red staining, is a non-invasive in vivo method for visualizing the effects of the molecules tested on the adiposity dynamics of zebrafish larvae by fluorescence microscopy. The 3-day, in vivo, animal treatment protocol was performed as previously described [30]. In replicated experiments, ten larvae ranging from 7 to 9 mm standard length were used per group. Larvae were exposed to the selected compounds or to vehicle alone for one day in a fasting state. All larvae were kept in a fasting state for 24 h prior to exposure and until the end of the trial, to avoid food auto-fluorescence. Treatments were as follows: vehicle control (CT) (0.1% DMSO), CA (0.1% DMSO plus 50 μM CA), HT (0.1% DMSO plus 100 μM HT), and ATX (0.1% DMSO plus 100 μM ATX). Concentrations of each molecule tested were the highest that did not induce any mortality. Quantitative analysis was performed, as previously described [30], by recording the image area of Nile red green fluorescence as a percentage of initial adiposity, using ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Rainbow trout adipocyte cell cultures
All cell-culture reagents were purchased from Sigma-Aldrich (Tres Cantos, Spain) and Life Technologies (Alcobendas, Spain). All plastic items and glass cover slips were obtained from Nunc (LabClinics, Barcelona, Spain). Cells were cultured according to the previously established procedure [31], with perivisceral WAT from four to six fish per culture. After counting, cells were seeded at a final density of 2-2.5Á10 4 cells/cm 2 in 1% gelatin on pretreated six-well plates (9.6 cm 2 /well) for real-time quantitative PCR (qRT-PCR) analyses or twelve-well plates (2.55 cm 2 /well), with or without coverslips, for immunofluorescence or Oil red O (ORO) staining, respectively. Plates were kept at 18˚C in Leibovitz's L-15 growth medium, supplemented with 10% fetal bovine serum and 1% antibiotic-antimycotic solution (growth medium, GM). When necessary, the standard procedure used for cell differentiation was the following: once confluence was reached (day 7), cells were induced to differentiate by incubating them with a differentiation medium (DM) based on GM and containing 10 μg/mL insulin, 0.5 mM 3-isobutyl-1-methylxanthine and 0.25 μM dexamethasone. The medium was changed every 2 days throughout the procedure.

Immunofluorescence assay
Post-confluent pre-adipocyte cells from day 7 of culture were incubated with vehicle CT (DM ± 0.1% DMSO) or vehicle plus CA (50 μM), HT (100 μM) or RGZ (1 μM), or a combination of CA or HT with RGZ, for 24 h. RGZ was used as a potential rainbow trout peroxisome proliferator-activated receptor gamma (PPARγ) agonist. PPARγ was detected by immunofluorescence, using the protocol described by [32]. The polyclonal rabbit anti-PPARγ (H-100) was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Secondary Alexa Fluor 1 conjugated antibody (A21069, goat anti-rabbit 568) was purchased from Life Technologies (Alcobendas, Spain). Nuclei were counterstained with Hoechst (H1399, Life Technologies, Alcobendas, Spain). Images were obtained at 36x magnification on a Leica TCS-SP5 confocal microscope. Nucleus fluorescence was quantified and normalized to the total number of nuclei in the same field, using ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Western blot analysis
Post-confluent pre-adipocyte cells from day 7 of culture were treated as described in the previous immunofluorescence section. Protein extraction and Western blot analysis were performed using the protocol described by [32]. Briefly, the amount of protein from each sample was measured [33] and 20 μg were subjected to electrophoresis (SDS-PAGE) on 10% polyacrylamide gels (125 V for 1 h 30 min). After overnight transfer to a PVDF membrane, a staining with Ponceau S solution (Sigma-Aldrich, Tres Cantos, Spain) was performed, showing similar amounts of transferred proteins on each lane and membranes were scanned for posterior band quantification. Subsequently, membranes were washed and then blocked in non-fat milk 5% and incubated with polyclonal rabbit anti-PPARγ (H-100), an antibody that has been previously shown to successfully cross-react with rainbow trout [31,34]. After washing, membranes were incubated with a peroxidase-conjugated secondary goat anti-rabbit antibody (Cat. No. 31460. Thermo Scientific, Alcobendas, Spain). The immunoreactive band was visualized using an enhanced chemiluminescence kit (Pierce ECL Western blotting Substrate; Thermo Scientific, Alcobendas, Spain) and quantified by densitometric scanning using ImageJ software (National Institutes of Health, Bethesda, MD, USA). Results from the densitometry analysis of each specific band were normalized by the densitometry values of the most abundant band of Ponceau S staining as previously reported [35]. In support of this methodology, it has been shown that reversible Ponceau S staining can be used advantageously over specific proteins detection for quality or control of equal loading in Western blotting [36,37].

Oil red O staining and lipid quantification
After confluence (day 7), cells were incubated with vehicle CT (DM ± 0.1% DMSO) or vehicle plus CA (50 μM), HT (100 μM), LIP (10 μl/ml) or the indicated combination of CA or HT with LIP, for 72 h. Cell differentiation and lipid accumulation were analyzed by ORO staining, as described elsewhere [38], with minor modifications. Briefly, cells were fixed with 10% formalin for 1 h. Fixed cells were rinsed with PBS, stained with 0.3% ORO prepared in 36% tri-ethyl phosphate for 2 h, and then rinsed three times with distilled water. A 100% 2-propanol solution was used to elute the ORO dye and absorbance was measured at 490 nm. The cells were then stained with Coomassie blue for 1 h and proteins were extracted using 85% propylene glycol at 60˚C for 1 h. Lipid quantification was calculated as the absorbance measured at 490 nm divided by the measurement corresponding to protein at 630 nm.

Cell viability assay
Pre-confluent pre-adipocyte cells (day 5 of culture) were incubated with vehicle CT (GM) or vehicle plus CA (50 μM), HT (100 μM), RGZ (1 μM), or a combination of CA or HT with RGZ, for 24 h. The methylthiazolyldiphenyl-tetrazolium bromide (MTT) assay was performed as previously described elsewhere [38]. Briefly, after 24 h incubation with a final concentration of 0.5 mg/mL MTT, cells were washed with PBS and the blue formazan crystals that formed were resuspended in 250 μL DMSO per well for 2 h. Cell viability values were obtained from the absorbance measured at 570 nm, with 680 nm as the reference wavelength, using a microplate reader (Infinite 200, Tecan).

Measuring cell proliferation
Pre-confluent pre-adipocyte cells (day 5 of culture) were incubated as described in the previous cell viability section. Cell proliferation was evaluated by immunocytochemical detection of proliferating cell nuclear antigen (PCNA), using a commercial staining kit (Cat. No. 93-1143, Life Technologies, Alcobendas, Spain). In brief, after 24 h incubation (see above), cells were washed and fixed in 4% paraformaldehyde (PFA, Sigma-Aldrich, Spain) at room temperature for 15 min. Subsequently, coverslips were post-fixed in 50% and 70% ethanol for 5 min and incubated in PCNA staining reagents, following the manufacturer's suggested protocol. The amount of PCNA-labeled nuclei (positive cells) was evaluated and normalized to the number of nuclei in that field, using ImageJ software (National Institutes of Health, Bethesda, MD, USA). Five to ten images were taken per coverslip with a CC2 camera coupled to a microscope at 40x using analySIS (Soft Imaging System) software.

In vivo experimental treatment in rainbow trout
After 15 days' acclimation in our facilities at 15˚C, juvenile rainbow trout fasted for 24 h were then anesthetized with MS-222 (0.1 g/L) prior to receiving an intraperitoneal injection of 4.64 μL volume per g body weight. Treatments were as follows: vehicle CT containing DMSO diluted in sesame oil (1:3, v/v), vehicle plus CA at 10 μg/g body weight, and vehicle plus HT at 20 μg/g body weight. After 24 h exposure to the compounds in a fasting state, trout were anesthetized, sacrificed by a blow to the head, and blood samples were taken from the caudal aorta. Liver and perivisceral WAT samples were then harvested and preserved at −80˚C pending analysis. None of the molecules tested induced mortality at the concentrations used.

Statistical analyses
Data were analyzed using IBM SPSS Statistics v.22 (IBM, Armonk, USA) and GraphPad prism 6 (La Jolla, USA, www.graphpad.com) and presented as mean ± SEM, unless stated otherwise. Normal distribution was first analyzed using the Shapiro-Wilk test, followed by Levene's to test homogeneity of variances. Statistical significance was assessed by two-tailed unpaired Student's t-test or one-way analysis of variance (ANOVA), followed by the Tukey post-hoc test. When data did not follow a normal distribution, a non-parametric Kruskal-Wallis test was applied, followed by a paired U-Mann Whitney test. Statistical differences were considered significant for all analyses when p-value 0.05, indicated by asterisks: Ã p 0.05, ÃÃ p 0.01, ÃÃÃ p 0.0001.
The mRNA level of each gene analyzed was calculated relative to the corresponding reference genes, i.e. geometric means of ef1a and ubiquitin for the in vitro cell culture assays and ef1a and actb for in vivo assays, using the Pfaffl method [40] implemented in the Biorad CFX manager 3.1 (Bio-Rad, El Prat de Llobregat, Spain).

Identifying compounds of plant origin with antioxidant properties able to decrease zebrafish larval adiposity in vivo
To investigate the effect of CA, HT and ATX on the dynamics of zebrafish adiposity, ZOT was used as a non-invasive in vivo method for visualizing lipid droplets in living larvae by fluorescence microscopy [30]. The whole-body adiposity dynamics of each larva was expressed as a percentage of variation in Nile red fluorescence signal areas in fish previously fed on standard diet (SD) after one-day's exposure in a fasting state. The results revealed that exposure to 50 μM CA significantly decreased larval adiposity compared to CT-exposed fish (−-22.83 ± 1.02% in CA versus −16.99 ± 1.29% with CT, p 0.01) (Fig 1A). The same effect was observed with 100 μM HT (−29.72 ± 2.74% in HT versus −17.98 ± 1.97% with CT, p 0.05) ( Fig 1B). In contrast, there was no change in adiposity following exposure to 100 μM ATX ( Fig  1C).

Differential effect of CA and HT on WAT dynamics in different body parts
ZOT was used to study WAT dynamics in different body parts of zebrafish larvae, during exposure to the selected bioactive compounds. Perivisceral WAT accounted for around 60% of adiposity at these zebrafish larval developmental stages (S1 Fig). The remaining WAT in the head and tail regions represented around 30% and 10%, respectively (S1 Fig). Results showed that exposure to CA mainly affected adiposity in the head region, compared to the same region in CT fish (−10.85 ± 1.95% in CA versus −5.53 ± 0.54% with CT, p 0.05) (Fig 2A). HT also led to a significant decrease in adiposity in the head (−11.46 ± 1.03% in HT versus −7.95 ± 0.24% with CT, p 0.05) and viscera (−15.87 ± 2.23% in HT versus −7.13 ± 2.46% with CT, p 0.05) compared to CT (Fig 2F). In contrast, ATX did not have a significant antiobesogenic effect in any of the body regions ( Fig 2K). Fluorescent images of isolated or grouped adipocytes revealed a marked decrease in oil droplet size following treatment with CA or HT (Fig 2C, 2E, 2H and 2J).

Potential antagonist effect of CA and HT on PPARγ signaling pathway
In view of the significant decrease in zebrafish larvae adiposity observed after treatment with CA and HT, the next step was to elucidate whether these compounds were capable of counteracting the obesogenic effect of RGZ, a potential PPARγ agonist, in vivo. RGZ at 1 μM on an SD background diet was shown to prevent adiposity loss in fasting condition: −10.59 ± 1.35% in RGZ versus −18.1 ± 1.52% with CT, p 0.01 ( Fig 3A) and −14.23 ± 0.98% in RGZ versus −20.04 ± 0.88% with CT, p 0.05 (Fig 3B). Simultaneous exposure of the larvae to RGZ and CA revealed that the obesogenic effect of RGZ was abolished by this compound (−-19.24 ± 0.40%) (Fig 3A). A similar effect was obtained following co-incubation with RGZ and HT (−23.47 ± 2.17%) (Fig 3B).   Quantification of the PPARγ immunofluorescence signal revealed that treating adipocytes with CA or HT in combination with RGZ significantly reduced the enhanced PPARγ protein expression signal produced by RGZ alone (Fig 4A and 4B). A very similar pattern although not showing significant differences was observed concerning PPARγ protein expression by Western blot (Fig 4C and 4D). A lower level of lipid storage was observed when these cells were cultured in the presence of lipid mixture (LIP)+CA and especially in the LIP+HT condition, compared to LIP alone (0.93 ± 0.176 and 0.56 ± 0.047 in LIP+CA and LIP+HT, respectively, versus 1.34 ± 0.13 with LIP, p 0.05) (Fig 4E). Finally, qRT-PCR data revealed that LIP significantly increased the expression of pparg and cebpa transcripts compared to CT, whereas RGZ did not affect these values (Fig 4F and 4G). In addition, treatments with CA or HT slightly down-regulated pparg and cebpa mRNA levels when combined with LIP, whereas no changes were observed in combination with RGZ (Fig 4F and 4G).

In vivo effects of CA and HT on rainbow trout lipid metabolism
The next step was to clarify the role of CA and HT in an in vivo context by focusing on several key adipogenic, lipolitic and β-oxidation markers, as well as the potential cross talk between liver and WAT, using qRT-PCR and plasma parameters analyses. The data revealed that transcriptional levels of fasn in both WAT and liver were significantly decreased by HT treatment (Fig 5A and 5E), indicating a reduction in fatty acid synthesis, possibly resulting in a decrease in TAG formation and fat deposits. Moreover, pnpla2 expression in WAT but not liver decreased significantly following HT administration (Fig 5D and 5H). On the other hand, lipe1 mRNA levels in both liver and WAT were unaffected by CA or HT treatment (Fig 5C  and 5G). Following CA administration, lpl mRNA levels were unaffected in WAT, but increased in liver, suggesting a higher TAG lipase activity in this tissue (Fig 5B and 5F). Regarding β-oxidation markers, acsl1 mRNA levels presented a reverse regulation between tissues, being significantly increased by HT treatment in WAT and showing an opposite pattern but not significant in liver (Fig 6A and 6D). Aside from that, no changes were found in hoad (Fig 6B and 6E) nor pparb (Fig 6C and 6F) mRNA levels due to treatment in any tissue.
Since CA and HT were shown to modulate the expression of adipogenic, lipolytic and βoxidation genes that regulate the lipid metabolism in both WAT and liver, it was useful to obtain further information by measuring blood parameters. Analysis of selected biochemical parameters in plasma revealed a slight decrease in TAG levels in fasting rainbow trout injected with CA and a significantly lower concentration with HT compared to CT (Table 1). However, no significant effects upon treatments were observed in plasma glucose nor NEFAs.

Discussion
While mammalian models have largely contributed to improving our understanding of obesity, in recent years, teleost models have also produced excellent findings on human diseases and offer a number of advantages [41]. Despite obvious differences, teleost fish share a significant amount of genetic identity with humans, and some metabolic pathways and organ systems are also remarkably similar [42,43]. Their rapid development ex-utero and the optical semi-transparency of the embryonic and early larval stages, as well as the advances in fish cell  [14,21,41,42]. In mammals, fat deposits in specific regions differ from each other not only by localization but also by their structural and functional properties [44]. Similarly, WAT development in zebrafish is a step-wise process, with differentiated adipocytes first observed in the visceral region, then the subcutaneous area and finally, the head [27]. Moreover, zebrafish fat deposits are also mobilized sequentially, in reverse order, in response to starvation [20]; altogether highlighting the particular suitability of this species to study adiposity dynamics in an in vivo scenario.
Natural bioactive phytochemicals have recently attracted interest for their potential health benefits in preventing metabolic diseases, including obesity and insulin resistance [45]. Among these, dietary components of olive oil, red wine and marine algae are particularly interesting, due to their antioxidant properties, which are traditionally linked to longevity and reduced mortality from several diseases [46]. Furthermore, as well as obesogenic molecules, which predispose individuals to metabolic changes leading to weight gain [47], there is growing evidence that some plant compounds can improve overweight by inhibiting adipogenic culture). mRNA levels of pparg (F) and cebpa (G) were normalized to the geometric mean of the two reference genes, ef1a and ubiquitin. For gene expression analyses cells were incubated with vehicle (DM) plus CA (50 μM), HT (100 μM), RGZ (1 μM), LIP (10 μl/ml), or the indicated combination of CA or HT with RGZ or LIP, or vehicle CT alone for 24 h (day 7 of culture). Data are shown as mean ± SEM (n = 3-7 cell cultures). Significant differences (p 0.05) are indicated by different letters, using one-way ANOVA followed by Tukey's post hoc test (B, C, E, F) or the non-parametric Kruskal-Wallis test followed by paired U-Mann Whitney test (G).
https://doi.org/10.1371/journal.pone.0178833.g004 pathways or impairing lipid accumulation and may, therefore, be classified as anti-obesogenic. Recent studies have reported that some phytochemicals and dietary supplements affect the lipid metabolism in zebrafish. Adding green tea extract to a high-fat diet significantly reduced body fat storage in adults [48,49]. Other studies of very early-stage larvae reported a decrease in lipids after treatment with compounds originating from plants or algae. The protocol  consisted of a 15-day exposure to the compound on a high-fat diet background, starting at the first feeding period. For example, curcumin [50][51][52][53], indole-3-carbinol [54], baicalein [55], dieckol [56], ellagic acid [52], kaempferol [51], quercetin [56], seapolynol [57] and silibinin [58], have all been reported to inhibit lipid accumulation in these early, post-embryonic developmental stages, when the adipocyte lineage is just starting to be established [20,27]. In our work, we used the ZOT protocol at later larval developmental stages, when the animals had a well-established WAT [30]. In vivo Nile red staining of adipocyte lipid droplets and quantitative analysis of whole-mount wide-field fluorescence microscopy signals, before and after a one-day exposure to the selected chemical compounds, were used to monitor the dynamics of WAT mass linked to adipocyte droplet size rather than adipocyte cell number. In addition, this methodology discriminated between the lipid signals associated with WAT and liver and identified the potential modulation of different regional WAT fat deposits in zebrafish larvae. In this context, the aim of the present study was to investigate the anti-obesogenic properties of three antioxidant compounds of plant origin, selected for their capacity to modulate the lipid metabolism in the 3T3-L1 adipocyte cell line and rodent models [59][60][61][62][63][64]. ATX is a natural red carotenoid pigment found in a wide variety of organisms, CA is one of the most abundant hydroxycinnamic acids in the human diet, and HT is bioavailable in olive leaves and olive oil. In the present work, CA and HT caused a significant, short-term reduction in zebrafish larval adiposity, but CA exhibited stronger anti-obesogenic effects only in the head region, initiating the reversal of the fat mobilization pattern, as described above, while HT affected both the head and visceral regions. These results indicate that these compounds reduce fat mass by decreasing adipocyte size, but they may have different anti-obesogenic capacities according to the WAT location in the body. As mentioned above, it is known that adipose deposits in different parts of the body have different biological functions and biochemical profiles. Several features, such as adipocyte growth and differentiation, developmental gene expression, susceptibility to apoptosis, inflammatory capacity, and adipokine secretion, vary among deposits, as well as fatty-acid processing and WAT enlargement and loss mechanisms [65]. Nevertheless, further investigation is required to determine how CA and HT target the specific needs of the different deposits. On the other hand, ATX did not modify the adiposity level of zebrafish, even when the different regions were analyzed separately. Although several studies support the hypothesis that ATX supplementation reduces body weight and fat accumulation in mice [66], the ATX action mechanism has been demonstrated to be cell-type dependent [62]. In agreement with our data, one study in humans reported that ATX did not boost fat utilization or fat loss [67].
Adipogenesis is a complex process that typically involves the sequential activation of several transcription factors, such as PPARγ, that regulate the differentiation of pre-adipocytes into mature adipocytes, and ultimately control WAT formation promoting both, lipogenesis and fatty acid uptake. In mammals, it has been demonstrated that CA and HT exhibit a significant potential to act as anti-obesity agents by modulating the PPARγ adipogenesis pathway [64,68]. Therefore, the potential antagonism of PPARγ produced by CA and HT may also disrupt fat accumulation in teleost fish and result in a reduction of WAT mass due to lipid storage impairment. Our results revealed that CA and HT abolished the RGZ-induced obesogenic effect in zebrafish. A similar inhibitory effect has been also reported in zebrafish exposed to specific pharmaceutical antagonists of RGZ [30,69], thus supporting the idea that the compounds used in the present study may antagonize PPARγ signaling.
During the last decade, in vitro cell culture models from fish species have been widely used for studies of lipid metabolism and adipocyte functionality [31, 70,71]. Nevertheless, despite the increasing use of zebrafish as an obesity model, to our knowledge, a primary adipocyte cell culture from this species has not been developed up to date. In this regard, further investigations using the well-established primary-cultured rainbow trout adipocyte system were performed in order to have a closer look into the potential PPARγ antagonism of CA and HT. The data supported the involvement of PPARγ in the mechanism underlying the anti-obesogenic activity of CA and HT. Concomitant with the counteracting effect downregulating the PPARγ protein expression signal induced by RGZ, CA and especially HT, also reduced the adipocyte lipid content, in agreement with our previous in vivo findings in zebrafish. However, qRT-PCR analyses revealed that these two compounds did not modulate the transcript levels of typical key markers of mature adipocytes, i.e. pparg and cebpa, when these cells were cotreated with RGZ, and had only a slight effect when they were combined with LIP. These findings suggest that the anti-obesogenic effect of CA and HT may be mediated by post-transcriptional mechanisms.
PPARγ deficiency and/or disruption directly not only affects WAT development and accumulation, but also exerts an impact on the whole body metabolism [72]. The in vivo administration experiment in rainbow trout revealed that CA and HT affected the mRNA levels of adipogenic and β-oxidation genes in liver and WAT in different ways. Fat loss is usually caused by the balance of two main processes: enhancement of fat breakdown and inhibition of TAG synthesis and accumulation. This process is orchestrated by the interaction of several tissues, such as WAT, liver and skeletal muscles [73]. The decrease in WAT fasn mRNA levels produced by HT treatment was accompanied by a reduction in plasma TAG levels, possibly due to the antagonist effect of this compound on the PPARγ signaling pathway. In agreement with our results, some mammal studies have demonstrated that CA and HT regulate WAT mass by suppressing lipogenic enzyme activity and claimed that PPARγ antagonists can be used to treat hyperlipidemia [74,75]. Interestingly, HT also downregulated pnpla2 transcript abundance in rainbow trout WAT, which is indicative of decreased lipolysis, leading to a reduction in lipid turnover or compensation for excessive WAT decrease. On the other hand, CA administration increased lpl mRNA levels in the liver, also associated with the decreased TAG levels observed in plasma. Concomitant with this decrease, CA and HT also smoothly increased NEFAs levels via TAG hydrolysis. Moreover, a slight decrease was also observed in plasma glucose following treatment, in agreement with mammal studies where CA and HT have been proposed as potential hypoglycemic antidiabetic treatments [76,77]. Within this in vivo scenario it is important to notice that CA and HT are antioxidants, with proven effects upon cell viability (S2 Fig), which may have an implication in modulating fatty acid oxidation [78]. In this sense, HT administration upregulated acsl1 mRNA levels in WAT while slightly decreased its expression in liver, indicating a tissue-specific regulation as previously reported [79]. Even though enhanced acsl1 expression is traditionally believed to be essential for the synthesis of TAG, several studies in mammals revealed a possible function related to the β-oxidation of fatty acids [80,81], in agreement with our results. Nevertheless, differences were not observed concerning the mRNA levels of hoad and pparb suggesting that further studies are needed to better understand the potential involvement of CA and HT in WAT β-oxidation.

Conclusions
The findings reported here provide novel insights into the anti-obesogenic role of CA and HT antioxidants, highlighting their potential involvement in negatively modulating the PPARγ signaling pathway. Further studies using specific inhibitors of these nuclear receptors will help to understand the mechanism of action of these anti-obesogenic compounds. The teleost fish models used have been validated for studying the mode of action of these bioactive compounds on WAT, reinforcing their suitability in pharmacological and biomedical research. This study may also help the investigation of new additives to optimize adiposity in farmed animals.