Introduction

All organisms can synthesise saturated and monounsaturated fatty acids. However, the biosynthetic rate of long-chain polyunsaturated fatty acids (LC-PUFA) in vertebrates is markedly low and cannot cover physiological demands. Linoleic acid (18:2n-6, LA) and α-linolenic acid (18:3n-3, ALA) are precursors for the synthesis of omega-6 (n-6) and omega-3 (n-3) LC-PUFA series, respectively, and essential fatty acids for vertebrates, which lack Δ12/n-6 and Δ15/n-3 desaturases required to synthesise LA from oleic acid (18:1n-9c, OA) and ALA from LA (Castro et al. 2016; Tocher et al. 2019). LC-PUFA are critical components for growth and development, acting as bioactive components of membrane phospholipids, precursors of signalling molecules and modulators of gene expression. Although the specific physiological roles of n-3 LC-PUFA remain unclear, eicosapentaenoic acid (20:5n-3, EPA) and docosahexaenoic acid (22:6n-3, DHA) are thought to exert protective roles preventing atherosclerosis, stroke, obesity, type-2 diabetes, inflammation and autoimmune diseases, among others (Calder 2018; Djuricic and Calder 2021). In contrast, n-6 LC-PUFA, particularly arachidonic acid (20:4n-6, ARA), are precursors of local hormones promoting acute and chronic inflammation. ARA is a highly abundant fatty acid in the membranes of many cell types. Inflammatory stimuli releases ARA from cell membranes and let cyclooxygenase, lipoxygenase and cytochrome P450 pathways convert ARA to eicosanoids, a family of mediators and regulators of the inflammatory response that include prostaglandins, thromboxanes and leukotrienes. Therefore, many eicosanoids are linked to inflammatory diseases, although some ARA-derived metabolites also seem to be involved in the resolution of inflammation (Djuricic and Calder 2021). The role of ARA on inflammation has been widely studied in mammals. In contrast, knowledge of ARA function is still scarce in fish and cannot be directly inferred from mammalian physiology. Nevertheless, it is well-known the role of eicosanoids, mainly prostaglandins, in the regulation of fish immunity and inflammation (Xu et al. 2022). Apart from plants, fungi and some aquatic microorganisms, few other organisms such as the nematode Caenorhabditis elegans and some invertebrates can synthesise de novo n-3 and n-6 LC-PUFA in significant amounts. Vegetable oils are rich in LA and ALA, and often contain high levels of n-6 LC-PUFA, but are devoid of significant amounts of n-3 LC-PUFA, particularly EPA and DHA unless obtained from transgenic crops. Therefore, trophic transfer from microalgae and plankton to marine fish and seafood are major sources of LC-PUFA, notably n-3 LC-PUFA, in the human diet (Tocher et al. 2019; Osmond and Colombo 2019).

Shortage of n-3 LC-PUFA and increased n-6/n-3 ratio in fish fillets due to substitution of fish oil (rich in n-3 LC-PUFA) by vegetable oils (poor in n-3 LC-PUFA, but frequently rich in n-6 LC-PUFA) in aquafeeds is nowadays a major challenge for the aquaculture sector. To face this problem and increasing demands of functional food with high nutritional value, intense research is being conducted in order to improve the n-3 LC-PUFA content in farmed fish, including dietary incorporation of microalgae, genetically modified organisms (GMOs) such as yeast and algae, and plant GMO-derived oils, such as oil from false flax expressing microalgal genes (Betancor et al. 2016; Tocher et al. 2019; Osmond and Colombo 2019; Sales et al. 2021; Carvalho et al. 2022). Production in large-scale fermenters and the supply of balanced amounts of EPA and DHA constrains the use of microalgae biomass in aquafeeds (Tocher et al. 2019). Transgenesis of fish fatty acid desaturases and elongases aiming to increase EPA and DHA levels was assayed in zebrafish (Alimuddin et al. 2007, 2008; Cheng et al. 2015). However, fish desaturases and elongases act on both n-3 and n-6 fatty acid series and generally do not substantially change the n-6/n-3 ratio (Pang et al. 2014). Efficient conversion of n-6 PUFA into n-3 PUFA in transgenic mice expressing Caenorhabditis elegans n-3 fatty acid desaturase fat-1 (FAT-1), an n-3 fatty acid desaturase absent in vertebrates (Kang et al. 2004), led to use synthetically humanised and fish codon-optimised C. elegans FAT-1 to generate transgenic zebrafish (Pang et al. 2014), common carp (Zhang et al. 2019), channel catfish (**ng et al. 2023), and other vertebrates, including mice, cattle, pigs and sheep (Lai et al. 2006; Ji et al. 2009; Chen et al. 2013; Liu et al. 2016, 2017; Li et al. 2018; Tang et al. 2019; Luo et al. 2020; Sun et al. 2020; You et al. 2021). Transgenesis of C. elegans fat-1 in zebrafish, common carp, channel catfish, pigs and sheep efficiently increases EPA and DHA, while decreases the n-6/n-3 ratio (Lai et al. 2006; Pang et al. 2014; Li et al. 2018; Zhang et al. 2019; Luo et al. 2020; **ng et al. 2023). The effect is potentiated in zebrafish and pigs by double transgenesis with codon-optimised C. elegans Δ12 fatty acid desaturase fat-2 (FAT-2), a Δ12 desaturase that converts OA into LA and which is also absent in vertebrates (Pang et al. 2014; Tang et al. 2019).

Given that there are major concerns on environmental risk, sustainability, fish welfare, food safety as well as consumer perception and acceptance of GMOs (Tocher et al. 2019; Osmond and Colombo 2019), in recent years we developed an alternative methodology to GMO generation based on the production of chitosan-tripolyphosphate (TPP)-DNA nanoparticles for transient modification of the expression of target genes in the liver of gilthead seabream (Sparus aurata) (González et al. 2016; Gaspar et al. 2018; Silva-Marrero et al. 2019). Chitosan is a cationic polymer of glucosamine and N-acetylglucosamine derived from chitin by deacetylation. Chitosan is increasingly used as carrier for delivering nucleic acids in vivo due to its well-known mucoadhesion, low toxicity, biodegradability and biocompatibility (Wu et al. 2020).

With the aim to promote sustained production of n-3 LC-PUFA in S. aurata, in the present study chitosan-TPP nanoparticles encapsulating plasmids expressing fish codon-optimised Caenorhabditis elegans FAT-1 and FAT-2 were intraperitoneally administered every 4 weeks to S. aurata (3 doses in total). Seventy days post-treatment, the effect of chitosan-TPP-DNA nanoparticles was assessed on growth parameters, intermediary metabolism and fatty acid content in the liver and skeletal muscle of S. aurata.

Materials and methods

Animals

S. aurata juveniles (7.7 g ± 0.2, mean weight ± SEM) were obtained from Piscicultura Marina Mediterranea (AVRAMAR Group, Burriana, Spain) and maintained at 20 °C in 250-L aquaria supplied with running seawater in the aquatic animals facility of the Scientific and Technological Centers of the Universitat de Barcelona (CCiTUB) as described (Silva-Marrero et al. 2017). Fish were fed with commercial diet (Dibaq Microbaq 165, Dibaq, Segovia Spain), containing 52% protein, 18% lipids, 12% carbohydrates, 10% ash, 8% moisture and 21.3 kJ/g gross energy. For the acclimation regime, fish were fed twice daily (9:00 and 17:00) at a ration of 5% body weight (BW). Two weeks before experimental treatments, the ration was adjusted and kept to 3% BW until the end of the experiment. Fish were weighted every 2 weeks to readjust the feed amount. To study the long-term effect of fish codon-optimised C. elegans fat-1 and fat-2 expression, 4 groups of fish were intraperitoneally injected up to 3 times (once every 4 weeks) with chitosan-TPP nanoparticles complexed with pSG5 (empty plasmid, control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2. Every single administration consisted of 10 μg plasmid per gram BW. Fourteen days after the last injection and 24 h following the last meal, fish were sacrificed by cervical section, blood was collected and the liver, intestine, skeletal muscle and brain were dissected out, frozen in liquid nitrogen and kept at − 80 °C until use. To prevent stress, fish were anesthetised by tricaine methanesulfonate (MS-222; 1:12,500) before handling.

Preparation and characterisation of chitosan-TPP-DNA nanoparticles

Fish codon-optimised FAT-1 and FAT-2 cDNA sequences (GenBank accession nos. ON374024 and ON374025, respectively) were synthesised based on C. elegans FAT-1 and FAT-2 using GeneArt Instant Designer (Thermo Fisher Scientific, Waltham, MA, USA) and ligated into pSG5 (Agilent Technologies, Palo Alto, CA, USA). The resulting constructs (pSG5-FAT-1 and pSG5-FAT-2) were verified by cycle sequencing on both sides. Chitosan-TPP nanoparticles encapsulating pSG5 (control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 were prepared by ionic gelation (González et al. 2016). For each experimental condition, 1 mg of plasmid was mixed with 4 mL of 0.84 mg/mL TPP (Sigma-Aldrich, St. Louis, MO, USA). Chitosan-TPP-DNA nanoparticles were formed upon dropwise addition of the TPP-DNA solution into 10 mL of 2 mg/mL low molecular weight chitosan (Sigma-Aldrich, St. Louis, MO, USA)-acetate buffer (pH 4.4) solution. Nanoparticles were sedimented by centrifugation at 36,000 g for 20 min at 15 °C and resuspended in 2 mL of 2% w/v mannitol, which acted as cryoprotector during lyophilisation. Nanoparticles were subjected to a freeze–drying process at − 47 °C. Particle size and Z potential were determined by dynamic light scattering and laser Doppler electrophoresis, respectively, using Zetasizer Nano ZS fitted with a 633 nm laser (Malvern Instruments, Malvern, UK). Chitosan-TPP-DNA nanoparticles were resuspended in 0.9% NaCl before intraperitoneal administration to S. aurata.

Body composition

For moisture determination, fish were dried at 85 °C until constant weight was reached (Busacker et al. 1990; Lucas 1996). Moisture was calculated as [wet weight (g) − dry weight (g)]*100/wet weight (g). Dried samples were further used for assaying nitrogen (N), lipid and ash. N content was determined with FlashEA 1112 analyser (Thermo Fisher Scientific, Waltham, MA, USA) and was subsequently used to estimate crude protein by multiplying N content by a factor of 6.25. Crude lipid was extracted with petroleum ether using a Soxhlet extractor. For ash determination, samples were incinerated in a Hobersal 12PR/300 muffle furnace (Hobersal, Caldes de Montbui, Spain) at 550 °C for 12 h (Busacker et al. 1990; Lucas 1996). Crude protein, lipid and ash are expressed as percentage of dry weight.

Growth parameters

Specific growth rate (SGR), feed conversion ratio (FCR), hepatosomatic index (HSI), protein retention (PR), lipid retention (LR) and protein efficiency ratio (PER) were calculated according to the following equations:

$${\text{SGR}} = \left( {{\text{lnW}}_{{\text{f}}} {-}{\text{ lnW}}_{{\text{i}}} } \right)*{1}00/{\text{T}};$$

where Wf and Wi are mean final and initial body fresh weight (g) and T is time (days)

$$\begin{gathered} {\text{FCR}} = {\text{dry}}\;{\text{feed}}\;{\text{intake }}\left( {\text{g}} \right)/{\text{wet}}\;{\text{weight}}\;{\text{gain }}\left( {\text{g}} \right) \hfill \\ {\text{HSI}} = {\text{liver}}\;{\text{fresh}}\;{\text{weight }}\left( {\text{g}} \right)*{1}00/{\text{fish}}\;{\text{body}}\;{\text{weight }}\left( {\text{g}} \right) \hfill \\ {\text{PR}} = {\text{body}}\;{\text{protein}}\;{\text{gain }}\left( {\text{g}} \right)*{1}00/{\text{protein}}\;{\text{intake }}\left( {\text{g}} \right) \hfill \\ {\text{LR}} = {\text{body}}\;{\text{lipid}}\;{\text{gain }}\left( {\text{g}} \right)*{1}00/{\text{lipid}}\;{\text{intake }}\left( {\text{g}} \right) \hfill \\ {\text{PER}} = {\text{weight}}\;{\text{gain }}\left( {\text{g}} \right)/{\text{feed}}\;{\text{protein}}\;{\text{provided }}\left( {\text{g}} \right) \hfill \\ \end{gathered}$$

Enzyme activity assays and metabolites

Enzyme activity assays and metabolites were spectrophotometrically determined at 30 °C in a Varioskan LUX multimode microplate reader (Thermo Fisher Scientific, Waltham, MA, USA). Liver crude extracts were obtained by homogenisation of powdered frozen tissue (1:5, w/v) in 50 mM Tris-HCl (pH 7.5), 4 mM, EDTA, 50 mM NaF, 0.5 mM phenylmethylsulfonyl fluoride, 1 mM dithiothreitol and 250 mM sucrose, 30 s at 4 °C using PTA-7 Polytron (Kinematica GmbH, Littau-Luzern, Switzerland). Following centrifugation at 10,000 g for 30 min at 4 °C, the supernatant was collected for enzyme activity assays. Reaction mixtures for 6-phosphofructo-1-kinase (Pfkl), fructose-1,6-bisphosphatase (Fbp1) and total protein were as previously described (Metón et al. 1999b). Enzyme activities were expressed as specific activity (U/g protein). One unit of Pfkl activity was considered the amount of enzyme needed to oxidise 2 μmol of NADH per min. One unit of Fbp1 activity of was defined as the amount of enzyme necessary for transforming 1 μmol of substrate per min. Serum glucose, triglycerides and cholesterol were measured with commercial kits (Linear Chemicals, Montgat, Spain).

Reverse transcription coupled to quantitative real-time PCR (RT-qPCR)

Total RNA from S. aurata tissues was isolated using HigherPurity Tissue Total RNA Purification Kit (Canvax, Cordoba, Spain) and reverse-transcribed with Moloney murine leukaemia virus reverse transcriptase (Life Technologies, Carlsbad, CA, USA) according to manufacturer's instructions. The mRNA expression levels of genes listed in Table 1 were determined using QuantStudio 3 Real-Time PCR System (Thermo Fisher Scientific, Waltham, MA, USA). The reaction mixture contained 0.4 μM of each primer (Table 1), 5 μL of SYBR Green (Thermo Fisher Scientific, Foster City, CA, USA), 0.8 μL of diluted cDNA and sterilized milli-Q water to final volume of 10 μL. The amplification cycle was 95 °C for 10 min, followed by 40 cycles at 95 °C for 15 s and 62 °C for 1 min. For each gene, standard curves for determining efficiency of the amplification reaction were generated with serial dilutions of control cDNA. Amplification of single products was confirmed by checking dissociation curves after each experiment. Amplicon size was checked by agarose gel electrophoresis. S. aurata ribosomal subunit 18S (18s), β-actin (actb) and elongation factor 1 alpha (eef1a) were used as endogenous controls to normalise the mRNA levels for genes of interest in liver samples. For tissue distribution, normalisation was performed against S. aurata 18s expression. The standard ΔΔCT method was used to calculate variations in gene expression (Pfaffl 2001).

Table 1 Primer sequences used for RT-qPCR in the present study
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Fatty acid methyl ester (FAME) analysis

Fatty acid profiles of liver and muscle were analysed by gas chromatography with flame ionisation detection as previously described (Silva-Marrero et al. 2019), using GC-2025 (Shimadzu, Kyoto, Japan) with capillary column BPX70, 30 m × 0.25 mm × 0.25 μm (Trajan Scientific and Medical, Ringwood, Australia). Oven temperature started at 60 °C for 1 min and then it was raised to 260 °C (rate: 6 °C/min). Injector (AOC-20i, Shimadzu, Japan) and detector temperatures were 260 °C and 280 °C, respectively. Sample (1 μL) was injected with helium as carrier gas and split ratio 1:20. Supelco 37 Component FAME Mix (Sigma-Aldrich, St. Louis, MO, USA) was used as reference for identifying fatty acids.

Statistics

To identify significant differences between treatments, the SPSS Version 25 software (IBM, Armonk, NY, USA) was used to submit experimental data to one-way analysis of variance followed by the Duncan post-hoc test (> 2 groups). Statistical significance was considered when P < 0.05.

Results

Delivery of chitosan-TPP complexed with pSG5-FAT-1 and pSG5- FAT-2 increases fish codon-optimised FAT-1 and FAT-2 mRNA levels in S. aurata

To assess the metabolic effects resulting from expression of C. elegans fat-1 and fat-2 in the liver of S. aurata, we designed fish codon-optimised C. elegans FAT-1 and FAT-2 cDNA sequences for further ligation into pSG5 and prepared chitosan-TPP nanoparticles complexed with empty pSG5 (control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 by ionic gelation. Particle size and Z potential of naked chitosan-TPP, expressed as mean ± SEM (n = 3), was 214.6 nm ± 20.2 and 37.5 mV ± 0.6, respectively. Incorporation of plasmid DNA to chitosan-TPP did not significantly modify particle size, which was 262.7 nm ± 74.0 (mean ± SEM, n = 3), but decreased Z potential to 12.0 mV ± 0.8 (mean ± SEM, n = 3).

For long-term sustained expression of fish codon-optimised fat-1 and fat-2 in the liver of S. aurata, each experimental group of fish received every 4 weeks up to 3 intraperitoneal injections of chitosan-TPP complexed with 10 μg/g BW of the corresponding plasmid (pSG5, pSG5-FAT-1, pSG5-FAT-2 or pSG5-FAT-1 + pSG5-FAT-2). The dosing schedule was based on preliminary studies showing that 28 days post-administration of chitosan-TPP-pSG5-FAT-1 and chitosan-TPP-pSG5-FAT-2 (10 μg/g BW of plasmid) to S. aurata increased the hepatic mRNA levels of fish codon-optimised fat-1 and fat-2 to levels even higher than those found at 72 h post-treatment. Expressed as mean ± SEM (n = 3), fold increase over control values at 72 h and 28 days post-treatment were 31.6 ± 4.1 and 74.8 ± 29.0, respectively, for fat-1 mRNA levels, while for fat-2 mRNA levels fold increase was 20.3 ± 2.8 and 70.2 ± 7.0, respectively. Seventy days after the beginning of the experiment (14 days following the last injection), the mRNA levels of fish codon-optimised fat-1 and fat-2 were determined by RT-qPCR in several tissues of treated fish, including the liver, intestine, skeletal muscle and brain (Fig. 1).

Fig. 1
figure 1

Effect of long-term treatment with chitosan-TPP nanoparticles complexed with pSG5 (control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 on the mRNA levels of fish-codon optimised C. elegans FAT-1 and FAT-2 in S. aurata tissues (brain, skeletal muscle, liver and intestine). Fourteen days after the last injection and 24 h following the last meal, exogenous fat-1 (a) and fat-2 (b) expression was assayed by RT-qPCR, normalised to the S. aurata 18 s mRNA levels and represented as mean ± SEM (n = 4). For each tissue, homogeneous subsets for the treatment are shown with different letters (P < 0.05)

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When compared with control fish, chitosan-TPP nanoparticles complexed with pSG5-FAT-1 and pSG5-FAT-2 significantly increased the mRNA levels of fat-1 and fat-2, respectively, in the liver and intestine of S. aurata. Specifically, fat-1 mRNA abundance in the liver of fish administered with pSG5-FAT-1 was 201.8-fold higher than in control fish, while treatment with pSG5-FAT-2 upregulated fat-2 297.4-fold. For the intestine, pSG5-FAT-1 and pSG5-FAT-2 upregulated 10.6-fold fat-1 and 24.7-fold fat-2, respectively. Nanoparticle administration did not exert effects on the skeletal muscle and brain.

Effect of fish codon-optimised FAT-1 and FAT-2 expression on whole-body composition, growth performance and serum metabolites in S. aurata

Sustained expression of fish codon-optimised FAT-1 + FAT-2 in the liver of S. aurata caused a moderate but significant 7.6% decrease of whole-body crude protein values observed in control fish. No effect was observed in moisture, ash and crude lipid body composition (Table 2). Analysis of growth performance parameters showed significantly increased weight gain values in fish expressing fat-1 (18% of increase) and fat-1 + fat-2 (26% of increase) compared to control fish. No significant difference was found between controls and treatment with FAT-2. Similarly, the highest SGR was found in fish treated with FAT-1 + FAT-2, followed by fish treated with FAT-1, controls and fish treated with FAT-2. Fish expressing fat-2 also presented the lowest PER. HSI significantly decreased in fish treated with FAT-1 and FAT-1 + FAT-2 to 72% of control values. No significant differences were observed in PR and LR.

Table 2 Growth performance, nutrient retention and body composition of S. aurata after intraperitoneal injection of chitosan-TPP complexed with empty vector (pSG5, control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2
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Serum glucose, triglycerides and cholesterol were also determined in 70-day treated S. aurata. Any of the treatments assayed affected blood glucose levels. However, co-expression of fat-1 + fat-2 significantly decreased 1.8-fold triglycerides and 1.5-fold cholesterol compared to control levels (Fig. 2).

Fig. 2
figure 2

Effect of long-term treatment with chitosan-TPP nanoparticles complexed with pSG5 (control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 on serum glucose (a), triglycerides (b) and cholesterol (c) in S. aurata. Fourteen days after the last injection and 24 h following the last meal, fish were sacrificed and the blood was collected. Values are represented as mean ± SEM (n = 6–7). Homogeneous subsets for the treatment are shown with different letters (P < 0.05)

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Effect of fish codon-optimised FAT-1 and FAT-2 on the fatty acid profile in the liver and skeletal muscle

The effect of long-term expression of fat-1 and fat-2 was analysed on the fatty acid profile of the liver and skeletal muscle of S. aurata. Table 3 shows the fatty acid composition in the liver of S. aurata long-term treated with chitosan-TPP nanoparticles complexed with pSG5 (control), pSG5-FAT-1, pSG5-FAT-2, and pSG5-FAT-1 + pSG5-FAT-2. Among 30 different fatty acids identified in this study, treatment with FAT-1 and FAT-1 + FAT-2 significantly increased EPA (1.5-fold and 1.6-fold, respectively), DHA (2.4-fold and 2.3-fold, respectively) and total n-3 fatty acids (1.7-fold in both cases). The n-6/n-3 ratio significantly decreased in fish expressing fat-1 (to 60.3% of control values), fat-2 (66.9%) and fat-1 + fat-2 (63.7%). A moderate 1.2-fold increase of cis-10-heptadecenoic acid (17:1n-7) was also observed in FAT-1 + FAT-2 treated fish, while expression of fat-1 decreased palmitoleic acid (16:1n-7) to 58.4% of control levels.

Table 3 Effect of chitosan-TPP complexed with empty vector (pSG5, control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 on the fatty acid profile of S. aurata liver
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The effect of long-term hepatic expression of fat-1, fat-2 and fat-1 + fat-2 on the fatty acids profile in the skeletal muscle is shown in Table 4. Twenty-one out of 29 fatty acids identified in the skeletal muscle were significantly affected by co-expression of fat-1 and fat-2. Total saturated fatty acids significantly decreased to 57.1% of control levels, mostly resulting from the low content in myristic acid (14:0; 34.4% of controls), palmitic acid (16:0; 56.3% of controls) and margaric acid (17:0; 50.0% of controls). In addition, treatment with FAT-1 and FAT-2 also decreased margaric acid to 50.0% of control values. In contrast, longer saturated fatty acids (with more than 17 carbons) presented increased values than in controls. Thus, stearic acid (18:0) increased 1.2-fold, while arachidic acid (20:0), behenic acid (22:0), tricosylic acid (23:0) and lignoceric acid (24:0) rised from non-detectable levels in control fish to low but detectable levels in fish treated with FAT-1 + FAT-2.

Table 4 Effect of chitosan-TPP complexed with empty vector (pSG5, control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 on the fatty acid profile of S. aurata skeletal muscle
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Monounsaturated, PUFA and total n-3, n-6 and n-9 fatty acids increased 1.3-fold, 1.3-fold, 2.2-fold, 1.1-fold and 1.5-fold, respectively, in the skeletal muscle of fish expressing fat-1 + fat-2. As a result of greater effect on n-3 series than on n-6 fatty acids, the n-6/n-3 ratio significantly decreased to 52.0% of control levels. Considering unsaturated fatty acids with a content greater than 1% for any assayed treatment, expression of fat-1 + fat-2 significantly increased EPA (1.7-fold) and DHA (3.0-fold) from the n-3 series, LA (18:2n-6c; 1.1-fold) and OA (18:1n-9c; 1.5-fold), while decreased palmitoleic acid to 61.5% of control levels. For less abundant unsaturated fatty acids (content between 0.1 and 1%), treatment with FAT-1 + FAT-2 also resulted in significant increases of cis-10-heptadecenoic acid (1.5-fold), gondoic acid (20:1n-9; 3.2-fold), erucic acid (22:1n-9; 5.2-fold), eicosadienoic acid (20:2n-6; 5.5-fold) and ARA (20:4n-6; 1.4-fold).

Effect of fish codon-optimised FAT-1 and FAT-2 on the expression of key genes in de novo lipogenesis and fatty acid oxidation in the liver

The effect of chitosan-TPP-DNA nanoparticles expressing fat-1 and fat-2 was also assessed on the hepatic expression of genes involved in de novo lipogenesis and fatty acid oxidation. As shown in Fig. 3, treatment with FAT-1 significantly decreased the mRNA levels of elongation of very long chain fatty acids protein 4a (elovl4a; to 43.3% of control values), elongation of very long chain fatty acids protein 4b (elovl4b; to 45.4%), elongation of very long chain fatty acids protein 5 (elovl5; to 62.3%), sterol regulatory element-binding protein 1 (srebf1; to 41.3%) and peroxisome proliferator-activated receptor alpha (ppara; to 46.0%), while treatment with FAT-2 decreased elovl5 (to 43.6%) and ppara (to 56.3%) mRNA levels (Fig. 3f–h, k, l). Co-expression of fat-1 and fat-2 also significantly downregulated acetyl-CoA carboxylase 1 (acaca; to 31.4% of control values), acetyl-CoA carboxylase 2 (acacb; to 65.0%), acyl-CoA 6-desaturase (fads2; to 69.4%), elovl4b (to 59.1%), elovl5 (to 41.8%), 3-hydroxy-3-methylglutaryl-coenzyme A reductase (hmgcr; to 64.2%) and srebf1 (to 41.9%) (Fig. 3a, b, e, g, h, j, k). No significant differences were found for fatty acid synthase (fasn), stearoyl-CoA desaturase-1a (scd1a) and carnitine O-palmitoyltransferase 1, liver isoform (cpt1a) (Fig. 3c, d, i).

Fig. 3
figure 3

Effect of long-term treatment with chitosan-TPP nanoparticles complexed with pSG5 (control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 on the expression of key genes in de novo lipogenesis and fatty acid oxidation in the liver of S. aurata. (a-l) Fourteen days after the last injection and 24 h following the last meal, fish were sacrificed and the liver were collected. Data are means ± SEM (n = 6). Expression data were normalised by the geometric mean of S. aurata 18 s, actb and eef1a mRNA levels. Homogeneous subsets for the treatment are shown with different letters (P < 0.05)

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Effect of fish codon-optimised FAT-1 and FAT-2 on glycolysis-gluconeogenesis, the pentose phosphate pathway, hnf4a and nr1h3 in the liver

Figure 4a–h shows the effect of long-term expression of fish codon-optimised fat-1 and fat-2 on the hepatic expression of rate-limiting enzymes in glycolysis-gluconeogenesis. Due to the pivotal role of the enzymes that control the flux through the fructose-6-phosphate/fructose-1,6-bisphosphate cycle in the regulation of glycolysis-gluconeogenesis, we measured both the mRNA levels and the enzyme activity of Pfkl and Fbp1. Gene expression of pfkl and fbp1 was not significantly affected by the treatments (Fig. 4d, e). However, when considering the Pfkl/Fbp1 activity ratio, fish treated with FAT-1 + FAT-2 exhibited a significant increased glycolytic flux (29.9%) compared to control fish (Fig. 4f). Similarly, treatment with FAT-2 showed a trend to increase the Pfkl/Fbp1 activity ratio (22.7%).

Fig. 4
figure 4

Effect of long-term treatment with chitosan-TPP nanoparticles complexed with pSG5 (control), pSG5-FAT-1, pSG5-FAT-2 and pSG5-FAT-1 + pSG5-FAT-2 on the expression of key genes in glycolysis-gluconeogenesis and the pentose phosphate pathway, hnf4a and nr1h3 in the liver of S. aurata. (a–k) Fourteen days after the last injection and 24 h following the last meal, fish were sacrificed and the liver were collected. Hepatic mRNA levels and enzyme activity of Pfkl and Fbp1 are presented as mean ± SEM (n = 6). Expression data were normalised by the geometric mean of S. aurata 18 s, actb and eef1a mRNA levels. Homogeneous subsets for the treatment are shown with different letters (P < 0.05)

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In regard of other glycolytic-gluconeogenic enzymes, expression of fat-2 and fat-1 + fat-2 significantly upregulated 1.5-fold and 1.8-fold, respectively, the mRNA levels of liver pyruvate kinase (pklr) and 1.4-fold those of 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase (pfkfb1) (Fig. 4c). Compared to controls, the expression levels for glucokinase (gck), glucose-6-phosphatase catalytic subunit (g6pc1) and phosphoenolpyruvate carboxykinase (pck1) were not affected (Fig. 4a, b, h).

In addition, expression of fat-2 also significantly increased the hepatic mRNA levels (2.0-fold) of glucose-6-phosphate dehydrogenase (g6pd), the rate-limiting enzyme in the oxidative phase of the pentose phosphate pathway (Fig. 4i), while treatment with FAT-2 and FAT-1 + FAT-2 upregulated hepatocyte nuclear factor 4-alpha (hnf4a; 1.5-fold) and oxysterols receptor LXR-alpha (nr1h3; 1.6-fold) mRNA levels, respectively (Fig. 4j–k).

Discussion

With the aim to induce sustained production of n-3 LC PUFA in S. aurata, the ionotropic gelation technique was used to obtain chitosan-TPP nanoparticles complexed with plasmids expressing fish codon-optimised C. elegans fat-1 and fat-2. C. elegans fat-1 and fat-2 were chosen to improve n-3 LC PUFA biosynthesis in S. aurata on the basis of their functionality in transgenic vertebrates, including fish. In fact, methyl-end desaturases from nematodes, including C. elegans fat-1 and fat-2, are considered one of the three main clades in the evolution of animal methyl-end desaturase genes, together with those from two distinct gene lineages of cnidarians and some lophotrochozoans and arthropods (Kabeya et al. 2018). Even though knowledge on pathways of PUFA biosynthesis in non-vertebrate animals is still limited, increasing availability of genomic data and functional characterisation of methyl-end desaturases from marine invertebrates will contribute to a better understanding of n-3 LC-PUFA biosynthesis in marine ecosystems (Monroig et al. 2022).

Monthly intraperitoneal administration of 3 doses of chitosan-TPP-DNA nanoparticles allowed long-standing high expressional levels of the exogenous proteins in the liver, mild expression in the intestine and barely detectable levels in the skeletal muscle and brain. Biodistribution of fish codon-optimised fat-1 and fat-2 expression shows that the particle size of chitosan-TPP-DNA complexes used in the present study favoured liver retention in S. aurata, which confirms previous reports where we analysed the acute effect of expressing exogenous SREBP1a and silencing of endogenous cytosolic alanine aminotransferase and glutamate dehydrogenase (González et al. 2016; Gaspar et al. 2018; Silva-Marrero et al. 2019). Possibly, discontinuous endothelia of the intestine enables chitosan-TPP-DNA nanoparticle absorption and transportation to the liver through portal circulation (Hagens et al. 2007), while the tight morphology of capillary endothelium in the muscle and brain may limit the transfer of nanoparticles and result in the scarce levels of transcript (Kooij et al. 2005). The feasibility of implementing chitosan-TPP-DNA administration in aquaculture relies on the fact that chitosan is recognised as safe by the U.S. Food and Drug Administration, on one hand, and that a procedure similar to that currently used to administer commercialised DNA vaccines for fish, could be applied to induce endogenous production of n-3 LC PUFA, on the other.

Transgenesis of fat-1, a gene that facilitates the conversion of n-6 to n-3 fatty acids, scarcely affected body growth in zebrafish, common carp, mice, pig and lamb (Bhattacharya et al. 2006; Ji et al. 2009; Liu et al. 2016; Zhang et al. 2018, 2019; Sun et al. 2020). However, in the present study long-term hepatic expression of fat-1 and fat-1 + fat-2 significantly increased weight gain but not lipid content in S. aurata. A trend to increase SGR was observed in fish co-expressing fat-1 and fat-2, while SGR values in control fish were in agreement with those reported for Sparus aurata under similar experimental conditions but non-treated with chitosan nanoparticles (Caballero-Solares et al. 2015; Sáez-Arteaga et al. 2022). Our findings suggest that increased levels of n-3 LC-PUFA and decreased n-6/n-3 fatty acid ratio resulting from expression of fish codon-optimised fat-1 and fat-2 may contribute to increased growth performance in S. aurata. In support of this hypothesis, substitution of fish oil (rich in n-3 LC-PUFA) by vegetable oil enhances the n-6/n-3 ratio and lowers n-3 LC-PUFA and weight gain in S. aurata (Houston et al. 2017), as well as in other marine fish such as cobia (Trushenski et al. 2012) and the anadromous Atlantic salmon (Qian et al. 2020). However, dietary fish oil does not significantly affect growth in other fish species such as zebrafish (Meguro and Hasumura 2018), common carp (Ljubojević et al. 2015), red hybrid tilapia (Al-Souti et al. 2012) and rainbow trout (Richard et al. 2006). Different adaptative responses to dietary n-3 LC-PUFA and the specific ability for converting n-3 and n-6 C18 PUFA into highly unsaturated long-chain fatty acids may result at least in part from functional diversification of Fads2 activity among teleosts, which in turn may have been influenced by a variety of factors such as phylogeny, trophic level, habitat (marine vs. freshwater) and trophic ecology (Castro et al. 2016; Garrido et al. 2019). Better growth performance of S. aurata submitted to sustained expression of fish codon-optimised fat-1 and fat-1 + fat-2 may also result from improved health condition due to increased n-3 LC-PUFA and decreased n-6/n-3 ratio. Consistently, fat-1 transgenesis prevents liver steatosis and lipid deposition in the abdominal cavity of zebrafish by a mechanism involving hepatic downregulation of lipogenic-related genes and upregulation of steatolysis-related genes (Sun et al. 2020). Moreover, fat-1 transgenesis prevents glucose intolerance, insulin resistance, non-alcoholic fatty liver disease and allergic airway responses in mice (Bilal et al. 2011; Kim et al. 2012; Romanatto et al. 2014; Boyle et al. 2020), and exerts protective vascular effects on pigs and cattle by reducing inflammatory factors and improving the immune system (Liu et al. 2016, 2017). Accordingly, S. aurata treated with FAT-1 and FAT1 + FAT-2 showed decreased HSI levels, which therefore may essentially result from lower lipid deposition in the liver of fish expressing fat-1.

Body fatty acid composition is affected by multiple factors, including de novo fatty acid synthesis, physiological requirements and dietary fatty acid profile. Single-gene expression of either fat-1 or fat-2 enhanced fatty acid desaturation and, consequently, n-3 LC-PUFA synthesis in transgenic mice (Pai et al. 2014), pig (Tang et al. 2019), zebrafish (Pang et al. 2014), and common carp (Zhang et al. 2019). Similarly, S. aurata long-term treated with chitosan-TPP-DNA nanoparticles expressing either fat-1 or fat-2 showed a general trend to increase liver and muscle EPA, DHA, and total n-3 fatty acids and PUFA, while decreased the n-6/n-3 ratio and saturated fatty acids, conceivably by conversion into unsaturated fatty acids. Most of these effects were potentiated by hepatic co-expression of fat-1 and fat-2. Combined activities of FAT-1 and FAT-2 decreased saturated fatty acids such as 14:0, 16:0 and 17:0, while increased unsaturated fatty acids, particularly n-3, and to a lesser extend n-9 and n-6. Thus, co-expression of fat-1 and fat-2 promoted a synergistic effect that favoured liver production of n-3 LC-PUFA and its accumulation in the muscle, particularly EPA and DHA. Given that fat-1 and fat-2 mRNA levels were scarcely detected in the skeletal muscle of S. aurata, changes in the muscle fatty acid profile between treatments may ascribe to hepatic fat exportation forming part of very low density lipoproteins (VLDL).

In contrast to most animals where fat-1 transgenesis generally results in a significant decrease of the n-6 fatty acid series (Kang et al. 2004; Lai et al. 2006; Liu et al. 2016, 2017; Li et al. 2018; You et al. 2021), the effect of fat-1 transgenesis in fishes seems species-dependent. Similarly as in S. aurata, transgenesis of fat-1 did not affect total n-6 fatty acids in the muscle of channel catfish (**ng et al. 2023), while a slight decrease was observed in zebrafish muscle (Pang et al. 2014). In contrast, fat-1 transgenesis largely decreased total n-6 PUFAs in common carp muscle (Zhang et al. 2019). The effect of fat-2 transgenesis in the fatty acid profile also depended on the fish species. Thus, fat-2 transgenesis in channel catfish decreased palmitic acid (16:0) and, as in S. aurata, led to a not significant trend to increase total n-6 and n-3 series in the muscle (**ng et al. 2023). However, fat-2 increased total n-6 PUFAs while did not affect the n-3 series in zebrafish muscle (Pang et al. 2014). Divergent effects of C. elegans fat-1 and fat-2 among fish species may be attributed to the interaction between exogenous enzymes and the diversity of endogenous desaturases and elongases that are distributed within different teleost taxonomic groups, and which in turn are responsible for different specific abilities for converting n-3 and n-6 C18 PUFA into highly unsaturated LC-PUFAs in teleosts (Monroig et al. 2022).

Among biochemical parameters improved by fat-1 transgenesis, reduced circulating levels for triglycerides and cholesterol were reported in mice (Romanatto et al. 2014), pigs (Liu et al. 2016), and cattle (Liu et al. 2017), while decreased hepatic triglycerides and cholesterol ester were found in zebrafish (Sun et al. 2020). Similarly, hepatic co-expression of fat-1 and fat-2 reduced serum triglycerides and cholesterol in S. aurata. In this regard, the present study showed that long-term co-expression of fat-1 and fat-2 promoted a general decrease of the expression of key enzymes for de novo lipogenesis in the liver of S. aurata. In addition, decreased serum triglycerides may also be attributed in part to the increase of n-3 LC-PUFA in the liver, where n-3 fatty acids are generally thought to reduce the production of VLDL and induce fatty acid β-oxidation (Shearer et al. 2012).

Treatment with FAT-1 + FAT-2 significantly downregulated hmgcr, which encodes the rate-limiting enzyme in cholesterol synthesis, and key genes in fatty acid synthesis, such as acaca and acacb, which catalyse conversion of acetyl-CoA into malonyl-CoA in the cytosol and mitochondrion, respectively, fatty acid elongases (elovl4b and elovl5) and fads2 desaturase. Although no significant, the expression of scd1a, which catalyses the insertion of a cis double bond at the Δ9 position into saturated C16 and C18 fatty acyl-CoA (Wang et al. 2005), also showed a trend to decrease in fish treated with nanoparticles expressing fat-1 + fat-2. Furthermore, the expression of the three fatty acid elongases herein analysed (elovl4a, elovl4b and elovl5) strongly decreased by sustained expression of fat-1. In transgenic animals, the effect of FAT-1 seems to depend on a variety of factors including the species, environmental conditions and dietary lipid content. Similarly as in S. aurata, a high-fat diet (13.4% of crude lipid versus 18.0% used in the present study) downregulated the hepatic expression of acaca, fasn and scd1 in fat-1 transgenic zebrafish. However, a low-fat diet (3.1% of crude lipid) caused the opposite effects, upregulating the expression of the three genes (Sun et al. 2020). In line with our findings, fat-1 transgenesis in mice decreased the levels of phosphorylated ACACA and FASN (Romanatto et al. 2014). However, fat-1 transgenic common carp showed upregulation of fads2, elovl5 and elovl2 in the liver, and transgenic pigs co-expressing fat-1 and fat-2 also presented increased expression levels of elovl5 and elovl2 in the muscle, skin and fat (Zhang et al. 2019; Tang et al. 2019).

Downregulation of acacb in S. aurata treated with FAT-1 + FAT-2 suggests a limited synthesis rate of mitochondrial malonyl-CoA. Any of the treatments affected the mRNA abundance of cpt1a, which is essential for the mitochondrial uptake of long-chain fatty acids and their subsequent β-oxidation in the mitochondrion. However, given that malonyl-CoA is a potent allosteric inhibitor of CPT1A (Saggerson 2008), our data suggest that in addition to decrease de novo lipogenesis, sustained co-expression of fat-1 and fat-2 may increase fatty acid oxidation in the liver of S. aurata fed medium- or high-fat diets. Similar results were reported for zebrafish fed a high-fat diet, where fat-1 transgenesis stimulated lipolysis-related genes and mitochondrial energy metabolism-related genes while downregulated the hepatic expression of genes related with lipogenesis and lipid deposition (Sun et al. 2020). Accordingly, upregulation of hepatic fatty acid oxidation-related genes by fat-1 transgenesis was also reported in common carp (Zhang et al. 2019), mice (Romanatto et al. 2014; Boyle et al. 2020) and goat cells (Fan et al. 2016), as well as in fat-1 and fat-2 double transgenic zebrafish (Pang et al. 2014).

In mammals, alternate promoters in the srebf1 gene generate SREBP1a and SREBP1c, which constitute transcription factors with a major role in de novo lipogenesis activation. SREBP1c primarily transactivates genes required for fatty acid and triglyceride synthesis while SREBP1a is a potent activator of all SREBP-responsive genes, including genes associated with cholesterol synthesis. Consistent with the role of srebf1 in the transcription of lipogenic genes both in fish and mammals (Carmona-Antoñanzas et al. 2014; Silva-Marrero et al. 2019), downregulation of srebf1 in the liver of S. aurata submitted to long-term expression of fish codon-optimised fat-1 and fat-1 + fat-2 led to a trend to decrease the expression of genes involved in cholesterol synthesis (hmgcr) and fatty acid synthesis (acaca, acacb and fasn), desaturation (scd1a and fads2) and elongation (elovl4a, elovl4b and elovl5). In agreement with our findings, transgenic zebrafish expressing fat-1 (when feeding a high-fat diet), fat-2 and fat-1 + fat-2 and double transgenic pigs for fat-1 and fat-2 also showed downregulated expression levels of srebf1 (Pang et al. 2014; Tang et al. 2019; Sun et al. 2020). Since DHA suppresses srebf1 expression and enhances its protein degradation (Jump 2008), increased levels of DHA seem the main responsible for decreased expression of srebf1 and de novo lipogenic genes in the liver of S. aurata expressing fat-1 and fat-1 + fat-2. Consistently, substitution of fish oil, rich in DHA, by vegetable oil leads to upregulation of srebf1 and fatty acid synthesis-related genes in S. aurata (Ofori-Mensah et al. 2020).

In the present study, FAT-1 and FAT-2 downregulated ppara in the liver of S. aurata. PPARA is a nuclear receptor activated by a wide range of ligands including fatty acids and fatty acid metabolites, such as eicosanoids. In the mammalian liver, PPARA controls the expression of genes involved in fatty acid uptake, intracellular transport, acyl-CoA formation and fatty acid mitochondrial and peroxisomal oxidation, ketogenesis and lipoprotein metabolism (Bougarne et al. 2018). Supplementation of fish oil to rodents enhances ppara expression in the liver (Hein et al. 2010; Kamisako et al. 2012), possibly as a result of increased availability of n-3 LC-PUFA, particularly EPA. However, the effect of dietary fish oil on ppara expression in fish depends on the species. Similarly as in S. aurata expressing fat-1 and fat-2, fish oil was shown to decrease the hepatic mRNA levels of ppara in S. aurata and juvenile turbot (Peng et al. 2014; Ofori-Mensah et al. 2020), while the opposite effect was reported in large yellow croaker and lean, but not fat, Atlantic salmon (Morais et al. 2011; Du et al. 2017). As pointed out by Peng et al. (2014), fatty acid-derived factors other than EPA-mediated activation may contribute to species-specific regulation of ppara expression in fishes.

In spite of limited knowledge of the effect of C. elegans FAT-1 and FAT-2 on glucose metabolism, fat-1 transgenesis was reported to improve glucose homeostasis by lowering hepatic gluconeogenesis and decreasing blood glucose and insulin in mice (Romanatto et al. 2014). Similarly, reduced plasma glucose was found in fat-1 transgenic cattle (Liu et al. 2017). In the present study, we found that long-term expression of fish codon-optimised C. elegans FAT-2 and FAT-1 + FAT-2 stimulated glycolysis and the expression levels of hnf4a and nr1h3 in the liver of S. aurata. By controlling the flux through the fructose-6-phosphate/fructose-1,6-bisphosphate substrate cycle, pfkl and fbp1 exert critical roles in hepatic glycolysis-gluconeogenesis. Although the mRNA levels of pfkl and fbp1 were not significantly affected by any of the treatments, expression of fat-2 and fat-1 + fat-2 promoted higher levels of Pfkl/Fbp1 activity ratio, possibly as a result of pfkfb1 upregulation. The bifunctional enzyme pfkfb1 catalyses the synthesis and degradation of fructose-2,6-bisphosphate, which is a major regulator of glycolysis–gluconeogenesis through allosteric activation of Pfkl and inhibition of Fbp1 (Okar et al. 2004). We previously showed that refeeding and high carbohydrate diets upregulate pfkfb1 and the kinase activity of the bifunctional enzyme in the liver of S. aurata, leading to a concomitant increase in fructose-2,6-bisphosphate levels (Metón et al. 1999a, 2000). As in mammals, fructose-2,6-bisphosphate is an allosteric activator of S. aurata PFKL (Mediavilla et al. 2008). Therefore, our results suggest that pfkfb1 upregulation in the liver of fish expressing fat-2 and fat-1 + fat-2 may be a key step favouring the glycolytic flux through the fructose-6-phosphate/fructose-1,6-bisphosphate substrate cycle, which in turn will increase the hepatic content of fructose-1,6-bisphosphate, an allosteric activator of Pklr.

The nuclear receptor HNF4A is a master regulator of liver metabolism through transcriptional regulation of target genes involved in glucose metabolism, lipid metabolism and hepatocyte differentiation (Meng et al. 2016). In mammals, HNF4A transactivates both glycolytic and gluconeogenic genes. Thus, HNF4A-binding to the gene promoter is required for insulin-stimulated upregulation of gck and pklr in the fed state, while a synergistic action of HNF4A and FOXO1 enhances the transcription of g6pc1 and pck1 during fasting (Hirota et al. 2008; Ganjam et al. 2009). Furthermore, HNF4A was previously shown to induce the expression of nr1h3 (Theofilatos et al. 2016), which encodes LXR-alpha, a nuclear receptor stimulated by insulin that is also involved in glucose and lipid metabolism (Zhao et al. 2012). Indeed, LXR-alpha was shown to upregulate pklr mRNA levels in mice (Cha and Repa 2007), and behave as a key regulator of pfkfb expression in humans by binding and transactivating the gene promoter of the bifunctional enzyme (Zhao et al. 2012). Therefore, increased hnf4a and nr1h3 mRNA abundance in S. aurata expressing fat-2 and fat-1 + fat-2 may enhance hepatic upregulation of pfkfb1 and pklr expression, and thus increase the glycolytic flux in the liver. Consistent with HNF4A-dependent enhancement of glycolysis in S. aurata, hnf4a expression was previously shown to increase in S. aurata under glycolytic conditions versus gluconeogenic conditions such as fasting and treatment with streptozotocin (Salgado et al. 2012). Increased levels of n-3 LC-PUFA may be a key factor leading to hnf4a and nr1h3 upregulation in the S. aurata liver. In agreement, fat-1 transgenic mice presented increased hepatic mRNA levels of hnf4a and to a lesser extend nr1h3 (Kim et al. 2012). Similarly, dietary supplementation with dried marine algae, rich in n-3 LC-PUFA (particularly DHA), induced hnf4a expression in the pig liver (Meadus et al. 2011). Furthermore, fish oil upregulated nr1h3 in S. aurata adipocytes (Cruz-Garcia et al. 2011), and in the liver of juvenile turbot, Nile tilapia and mice (Kamisako et al. 2012; Peng et al. 2014; Ayisi et al. 2018).

To our knowledge, the effect of fat-1 and fat-2 transgenesis on the pentose phosphate pathway was not previously addressed. In the present study, long-term expression of fish codon-optimised fat-2 and fat-1 + fat-2 promoted higher expression levels of g6pd, which encodes the rate-limiting enzyme for the production of NADPH in the oxidative phase of the pentose phosphate pathway. Previous reports indicated that dietary carbohydrates are a key factor that enhances G6pd activity in the liver of S. aurata (Metón et al. 1999b). Nevertheless, our findings support that fatty acid composition, particularly the n-3/n-6 ratio, also seems to regulate the hepatic expression of g6pd. In agreement with g6pd upregulation by n-3 LC-PUFA, fish oil stimulated G6PD activity in the rat liver (Yilmaz et al. 2004), and dietary supplementation with n-3 PUFA increased g6pd mRNA levels in the pig muscle (Vitali et al. 2018). Furthermore, n-6 PUFA, particularly LA, decreased g6pd mRNA levels in rat hepatocytes (Kohan et al. 2011). Species-specific regulation of g6pd expression by fatty acid composition may occur in other fishes. In this regard, total replacement of fish oil by vegetable oil did not affect G6pd activity but increased the mRNA levels in the liver of Nile tilapia (Ayisi et al. 2018), while enhanced G6pd activity in the liver of Atlantic salmon (Menoyo et al. 2005). Bearing in mind a general trend to downregulate de novo hepatic lipogenesis in S. aurata co-expressing fat-1 and fat-2, NADPH resulting from g6pd upregulation by n-3 LC-PUFA may reinforce cellular protection from oxidative stress.

Considered together, the metabolic effects of periodical administration of chitosan-TPP-DNA nanoparticles expressing fish codon-optimised C. elegans fat-1 and fat-2 included hepatic downregulation of de novo lipogenesis-related genes, leading to reduced circulating levels of triglycerides and cholesterol, stimulation of fatty acid oxidation and upregulation of glucose oxidation via glycolysis and pentose phosphate pathway without affecting glycemia. Although co-expression of C. elegans fat-1 and fat-2 in S. aurata reduced blood triglycerides, the action of fat-1 and fat-2 may allow protein sparing in the liver of a carnivorous teleost, such as S. aurata, by a mechanism involving increased glucose and fatty acid oxidation to obtain energy in fish fed medium- or high-fat diets.

Conclusion

The present study shows that long-term treatment with chitosan-TPP nanoparticles complexed with plasmids expressing fish codon-optimised C. elegans fat-1 and fat-2 allowed efficient expression of exogenous FAT-1 and FAT-2 desaturases in the liver of S. aurata, which in turn elevated the n-3 LC-PUFA content, particularly EPA and DHA, and decreased the n-6/n-3 ratio both in the liver and the skeletal muscle. Co-expression of fish codon-optimised fat-1 and fat-2 promoted the highest weight gain, n-3 LC-PUFA accumulation in the muscle, and had metabolic effects that included downregulation of lipid biosynthesis and stimulation of fatty acid and glucose oxidation in the liver. Expression of fish codon-optimised fat-1 and fat-1 + fat-2 downregulated the hepatic expression of srebf1 and as a consequence, the mRNA levels of key genes in de novo lipogenesis, while fat-2 and fat-1 + fat-2 upregulated hnf4a, nr1h3 and glucose oxidation through glycolysis and the pentose phosphate pathway. Our findings support that chitosan-TPP-DNA nanoparticles co-expressing fish codon-optimised fat-1 and fat-2 can alleviate the effect of fish oil replacement with vegetable oil currently occurring in aquafeeds and enable production of functional fish rich in EPA and DHA for human consumption. Future studies will help to determine dosage and the optimal developmental stage to administer fat-1 and fat-2 chitosan-TPP-DNA nanoparticles in cultured fish.

Availability of data

Data are available from the corresponding author upon reasonable request.