# Dietary DHA limitation did not affect swimming and metabolic performance, but reduced growth in wild European sea bass

Mickaël Péron, Philippe Soudant, Fabienne Le Grand, David Mazurais, Victor Simon, Christel Lefrançois, Marie Vagner

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Mickaël Péron, Philippe Soudant, Fabienne Le Grand, David Mazurais, Victor Simon, et al.. Dietary DHA limitation did not affect swimming and metabolic performance, but reduced growth in wild European sea bass. Biochimie, In press, 10.1016/j.biochi.2025.07.005 . hal-05175430

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Dietary DHA limitation did not affect swimming and metabolic
performance, but reduced growth in wild European sea bass

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Mickaël Péronª, Philippe Soudantª, Fabienne Le Grandª, David Mazuraisª, Victor Simonª,
Christel Lefrancoisᵇ, Marie Vagnerª

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a Univ Brest, CNRS, IRD, Ifremer, LEMAR, F-29280 Plouzane, France

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b Littoral Environnement et Sociétés (LIENSs), UMR 7266, CNRS-Université de La Rochelle, 2 rue Olympe de Gouges, F-17042 La Rochelle, France

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Corresponding author: mickael.peron.sci@gmail.com

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**Abstract**

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Long-chain polyunsaturated fatty acids (LC PUFA), particularly docosahexaenoic acid (DHA), are essential for cell membrane structure and function, impacting overall fish performance. These molecules, produced primarily by phytoplankton, are transferred up the trophic chain; however, climate change is predicted to modify phytoplankton communities with a cascading effect on the global DHA production and thus availability for consumers such as fish. This study aimed to evaluate the effects of dietary DHA limitation on i) the fatty acid composition in fish tissues ii) somatic growth, swimming performance, and metabolic rates, and iii) the activation of biosynthetic pathways at the molecular level, by measuring gene expression involved in DHA synthesis. We conditioned wild-caught European sea bass (*Dicentrarchus labrax*) juveniles for five months on a DHA-depleted or control diet. Dietary DHA limitation led to selective retention or synthesis of DHA in fish tissues (liver, brain, and muscle), a reduced growth and an up-regulation of DHA biosynthetic pathways without compensating for DHA deficiency in tissues. Fish fed the low DHA diet showed increased biosynthetic activity, suggesting that this pathway may be energetically costly, as high tissue DHA correlated with reduced growth. Alternatively, the lower tissue DHA levels in these fish might cause slower growth. However, metabolic rates and swimming performance were not affected by dietary treatment. Inter-individual variability was observed across all variables, highlighting underlying trade-offs when facing DHA limitation. This work provides insight into the physiological consequences of dietary DHA reduction due to global change and the mechanisms fish employ to mitigate its effects.
30 **Keywords:** Critical swimming speed; metabolic rate; n-3 LC PUFA; *fads2* gene expression;
31 Specific Growth Rate; dietary DHA

32 **Author contribution**

33 MP: Conceptualization, Investigation, Methodology, Data curation, Writing – original draft, –
34 review & editing.

35 PS: Conceptualization, Investigation, Data curation, Methodology, Supervision, Writing –
36 review & editing.

37 FLG: Conceptualization, Data curation, Investigation, Methodology, Supervision, Writing –
38 review & editing.

39 DM: Conceptualization, Data curation, Investigation, Methodology, Supervision, Writing –
40 review & editing

41 VS: Methodology, Investigation.

42 CL: Methodology, Investigation, Writing – review & editing.

43 MV: Conceptualization, Data curation, Investigation, Methodology, Supervision, Writing –
44 review & editing, Funding

45 **Funding**

46 University of Western Brittany and the Doctoral School of Marine and Coastal Sciences
47 funded M.P. This study is a contribution to the project ADJUST supported by the French
48 National program EC2CO (Ecosphère Continentale et Côtière), the Scientific Council of the
49 European Institute for Marine Sciences, and the Interdisciplinary graduate school for the
50 blue planet (ANR-17-EURE-0015).

51 **1. Introduction**

52 Long chain polyunsaturated fatty acids (LC PUFA), particularly omega-3 (n-3) LC PUFA are
53 essential components of cell membranes in most organisms including fish [1]. Among them,
54 docosahexaenoic acid (DHA, 22:6n-3) and eicosapentaenoic acid (EPA, 20:5n-3) are
55 important fatty acids (FA) found in high proportions in membrane polar lipids [2]. In some
56 tissues like brain or retina DHA is usually the major FA and can reach up to 50% of the
57 membrane FA composition [3]. Consequently, n-3 LC PUFA are crucial for cell function and
58 tissue integrity, influencing the overall biological performance of the organism [1,4]. In fish,
59 EPA is involved in the modulation of immune and inflammatory responses through the
60 production of eicosanoids such as prostaglandins and leukotrienes [5]. DHA is essential for
61 maintaining membrane fluidity and neural function, thereby supporting sensory and
62 cognitive abilities, stress responsiveness or visual acuity [3].

63 In natural environment, phytoplankton are the primary producers of n-3 LC PUFA, which are
64 subsequently transferred to higher trophic levels [6,7]. The availability of these FA at the
65 base of the food web is crucial for organisms like marine fish, which have a limited ability to
66 synthesize them due to a lack of key enzymes (desaturases and elongases) required for n-3
67 LC PUFA biosynthesis [1,8]. Environmental changes such as ocean acidification, rising
68 temperatures, and hypoxia have been shown to alter marine organisms' physiology,
69 including phytoplankton, and shift their community structure [9–11]. These shifts may
70 favour microalgae species less efficient in n-3 LC PUFA production while reducing the
71 abundance of those that are more productive, leading to a decrease in n-3 LC PUFA
72 availability [10,12–14].

73 The effects of reduced dietary n-3 LC PUFA on fish performance have been extensively
74 studied in aquaculture, particularly in relation to somatic growth, survival, and lipid
75 composition of tissue [15–17]. However, few studies have explored the impact of reduced
76 dietary n-3 LC PUFA on ecologically relevant parameters like aerobic metabolism and
77 swimming performance [18–20].

78 For instance, adult Atlantic salmon (*Salmo salar*) fed a diet rich in n-3 LC PUFA for 112 days
79 exhibited reduced maximum aerobic swimming speed (Ucrit) compared to fish fed on a
80 canola oil-enriched diet lower in n-3 LC PUFA [21]. However, other authors found that post-
81 juvenile Atlantic salmon fed high n-3 LC PUFA levels for 31 weeks had higher Ucrit than those
82 fed a poultry fat-enriched diet, lower in n-3 LC PUFA [22]. Similarly, Arctic charr (*Salvelinus
83 alpinus*) fed a high n-3 LC PUFA diet had greater Ucrit values than those fed rapeseed and
84 palm oil-enriched diets, which were lower in n-3 LC PUFA [23]. In golden grey mullet (*Chelon
85 auratus*) juveniles, dietary n-3 LC PUFA content did not affect Ucrit, but it did increase oxygen
86 consumption during swimming and impaired recovery capacities [20].

87 However, these studies examined the effects of a combined dietary limitation of EPA and
88 DHA. It was reported that the reduction in DHA production by microalgae is expected to be
89 greater than for other n-3 LC PUFA [10] potentially decreasing DHA availability for higher
trophic levels, including fish [9,10,24,25]. Some studies have shown that DHA alone (without EPA) can meet the n-3 LC PUFA requirements for growth and survival in juvenile cobia (*Rachycentron canadum*) [26]. DHA has been extensively studied for its benefits on vertebrate cardio-vascular function, brain development, neurological functions and cognition (Lauritzen et al., 2001; Sargent et al., 1999; Tocher, 2003). In teleost fish, DHA is involved in muscle fiber development by influencing mitochondrial quality control as well as improving mitochondrial function [30–32]. Additionally, DHA may improve carbohydrate utilization, which is vital for maintaining growth and energy efficiency [33]. Understanding the effects of sole dietary DHA limitation on the physiology of organisms with limited ability to synthesize it is therefore critical to apprehend how a dietary DHA decrease may impact fishes in the wild.

The European sea bass is an active and opportunistic predator foraging mostly small fish and crustaceans, and subject to predation by bigger fish or avian predators at juvenile stage [34]. Given its active nature, swimming performance may serve as a proxy for individual fitness in natural environments [35]. To date, only one study has explored the relationship between dietary fatty acids and metabolic or swimming performance in European sea bass [36]. This study found that a dietary limitation in n-3 LC PUFA reduced growth rates, while higher n-3 LC PUFA levels decreased metabolic rates. However, the results of this study were based on domesticated fish, which may have genotypic differences affecting phenotypic traits (e.g., metabolism or swimming performance) compared to their wild counterparts to ensure ecological relevance and accurately capture the impacts of global environmental changes on natural aquatic ecosystems. Farmed fish are often selectively bred for traits like fast growth or disease resistance and are raised in controlled environments over several generations. This can result in genetic and behavioral differences compared to wild populations, potentially altering their physiological or ecological responses to stress [37,38]. Using wild fish reduces these confounding factors and provides a more accurate representation of how natural populations may be affected. In addition, wild fish exhibit a larger inter individual variability in traits, which is essential to understand how global change may impact natural populations.

In this context, the aim of this study was to use wild caught sea bass brought to the laboratory to experimentally assess the effects of a dietary DHA limitation on (i) growth,
swimming performance, and associated metabolic rates, using the critical swimming speed ($U_{crit}$) step protocol [39], and (ii) the activation of biosynthetic pathways at the molecular level to compensate for DHA limitation by measuring the expression of genes involved in fatty acid biosynthesis. We hypothesized that a five-months dietary DHA limitation will stimulate the biosynthetic pathways at the molecular level [40,41], inducing a metabolic cost, that would consequently alter growth and metabolic performances. This study will enhance our understanding of the underexplored consequences of global change on an ecologically relevant species.

## 2. Material and Methods

### 2.1. Ethical Statement

All of the experiments were performed with respect of the regulations of the Animal Care Committee of France (ACCF). All experiments were conducted at the Ifremer facilities (Plouzané, France, Agreement number: B29-212-05) and approved by the ethical committee (DAP30512_2021031811505740). All precautions were taken to minimize fish suffering.

### 2.2. Fish Maintenance

#### 2.2.1. Fish collection

110 juvenile European sea bass were captured in the Seine estuary during the NOURDEM survey in August 2021 [42]. Fish were then immediately transported to Ifremer rearing facilities (Plouzané, France) into an oxygenated (> 120 % of air saturation) 400 L tank containing the water of the environment in which they were caught.

#### 2.2.2. Rearing

Fish (n = 110) were acclimated in quarantine during 66 days in a single 2000 L tank with open circuit and natural temperature that followed the natural variation of the bay of Brest. During this period, they were fed a mix (from 25 to 100%) of frozen seafood and commercial pellets (NeoStart, Le Gouessant, Lamballe, France) in order to acclimate them to pellets as described in previous studies [20,43,44]. After 30 days, fish were then fed exclusively on commercial pellets for 36 days. After these 66 days of quarantine, fish (n=101; 18.9 ± 0.2 cm, 73.6 ± 2.4 g; nine fish died at the beginning of the quarantine period) were transferred to six experimental 400 L tanks (n=16-17 individuals per tank, total n = 101) in open circuit,
exposed to a 12h light / 12h dark photoperiod cycle, and natural temperature cycle (mean = 14.5°C ± 0.14, min-max = 10.3°C-18.7°C). Before being transferred to their tanks, fish were anesthetized using MS-222 (tricaine methanesulfonate, 0.1 g L⁻¹), individually weighed, measured and individually identified with pit-tags (M120; biolog-id, Réseaumatique, Bernay, France). Fish were then progressively (from 25 to 100% of experimental diet in 2 weeks) fed with two experimental diets, i.e. either a control or a DHA-depleted diet (Table 1). These diets were formulated to be iso-lipidic, iso-proteic and iso-caloric, but with a contrasting amount of DHA, by adding either fish oil or microalgae oil (*Nannochloropsis* spp.), respectively. The oil composition was the only parameter that differed between the two diets. The final protein and lipid contents ranged within the needs defined for Atlantic sea bass (e.g Boujard et al., 2004; Peres and Oliva-Teles, 2001; Skalli and Robin, 2004). Both diets were manufactured at INRAE (Donzacq, France). The two conditions (hereafter called Control and Depleted, respectively) were reared in triplicate (n= 51 fish for the Control condition and n= 50 for the Depleted condition). Fish were fed this diet once a day (0.6-2 % of total tank biomass, depending on water temperature) for a minimum of 163 days and a maximum of 192 days. The oxygen saturation in the tanks was daily controlled and was never below 90%. Fish were monitored daily and their feed intake was checked by visual observation. Every month, the diet quantity was adjusted by monitoring fish growth. For that, 24h-starved fish were anesthetized with MS-222 (0.1 g.L⁻¹) and then weighed (± 0.1 g) and measured (± 0.1 cm).

<table><caption>Table 1: Formulation and fatty acid composition (% of total lipids) of the two experimental diets (Control and Depleted). ARA: Arachidonic acid; DHA: Docosahexaenoic acid; EPA: Eicosapentaenoic acid; FA: Fatty acids; LC PUFA: Long chain polyunsaturated fatty acids; MUFA: Monounsaturated fatty acids; PUFA: Polyunsaturated Fatty acids; SFA: Saturated Fatty acids. See in the next section of the material and methods section for fatty acid analysis protocol.</caption><thead><tr><th>Ingredients (g 100g<sup>-1</sup>)</th><th>Control diet</th><th>Depleted diet</th></tr></thead><tbody><tr><td>Fish Meal</td><td>20</td><td>20</td></tr><tr><td>Wheat gluten</td><td>50</td><td>50</td></tr><tr><td>Canola oil</td><td>3</td><td>5</td></tr><tr><td>Fish oil</td><td>7</td><td>0</td></tr><tr><td><i>Nannochloropsis</i> spp. Oil</td><td>0</td><td>5</td></tr><tr><td>Starch</td><td>6</td><td>6</td></tr><tr><td>Vitamins</td><td>4</td><td>4</td></tr><tr><td>Minerals</td><td>2</td><td>2</td></tr><tr><td>Cellulose</td><td>5</td><td>5</td></tr><tr><td>Betaine</td><td>3</td><td>3</td></tr></tbody></table>
Proximal composition (g 100g⁻¹ ± SD)

<table><tr><td>Proteins</td><td>59.2 ± 1.8</td><td>59.5 ± 1.8</td></tr><tr><td>Lipids</td><td>13.8 ± 1.0</td><td>13.1 ± 1.0</td></tr><tr><td>Nitrogen</td><td>9.5 ± 0.3</td><td>9.5 ± 0.3</td></tr></table>

FA composition (% of total FA ± SD)

<table><tr><td>14:0</td><td>4.9 ± 0.5</td><td>2.2 ± 0.5</td></tr><tr><td>15:0</td><td>0.3 ± 0.5</td><td>0.2 ± 0.5</td></tr><tr><td>16:0</td><td>15.1 ± 1.2</td><td>12.8 ± 1.0</td></tr><tr><td>17:0</td><td>0.1 ± 0.5</td><td>0.1 ± 0.5</td></tr><tr><td>18:0</td><td>3.2 ± 0.5</td><td>1.2 ± 0.5</td></tr><tr><td>20:0</td><td>0.4 ± 0.5</td><td>0.4 ± 0.5</td></tr><tr><td>22:0</td><td>0.2 ± 0.5</td><td>0.2 ± 0.5</td></tr><tr><td>Σ SFA</td><td>24.8 ± 1.0</td><td>17.9 ± 1.5</td></tr><tr><td>16:1n-7</td><td>4.8 ± 0.5</td><td>9.4 ± 0.8</td></tr><tr><td>16:1n-9</td><td>0.1 ± 0.5</td><td>0.5 ± 0.5</td></tr><tr><td>18:1n-7</td><td>2.8 ± 0.5</td><td>2.0 ± 0.5</td></tr><tr><td>18:1n-9</td><td>22.7 ± 1.8</td><td>30.0 ± 2.4</td></tr><tr><td>20:1n-9</td><td>1.5 ± 0.5</td><td>0.6 ± 0.5</td></tr><tr><td>Σ MUFA</td><td>35.7 ± 2.3</td><td>43.8 ± 2.6</td></tr><tr><td>18:2n-6</td><td>12.6 ± 1.0</td><td>17.0 ± 1.4</td></tr><tr><td>18:3n-3</td><td>2.9 ± 0.5</td><td>5.1 ± 0.5</td></tr><tr><td>20:4n-6</td><td>0.8 ± 0.5</td><td>1.5 ± 0.5</td></tr><tr><td>20:5n-3</td><td>10.7 ± 0.9</td><td>13.2 ± 1.1</td></tr><tr><td>22:5n-3</td><td>1.5 ± 0.5</td><td>1.2 ± 0.5</td></tr><tr><td>22:6n-3</td><td>5.3 ± 0.5</td><td>0.5 ± 0.5</td></tr><tr><td>Σ PUFA</td><td>39.6 ± 2.5</td><td>39.5 ± 2.4</td></tr><tr><td>Σ LC PUFA</td><td>18.6 ± 1.2</td><td>15.7 ± 1.0</td></tr><tr><td>Σ n-3</td><td>22.7 ± 0.5</td><td>19.9 ± 1.0</td></tr><tr><td>Σ n-6</td><td>14.0 ± 1.1</td><td>19.3 ± 1.2</td></tr><tr><td>EPA/DHA</td><td>2.0 ± 0.1</td><td>26.4 ± 0.5</td></tr><tr><td>ARA/EPA</td><td>0.07 ± 0.0</td><td>0.11 ± 0.0</td></tr></table>

Analyzed by UpScience
(St Nolff, France)

Proximal composition: Fish meal: LT70 Sopropêche; Wheat gluten: Roquette; Canola oil: Oléandes; FishOil: Polaris Omegavie®; Nannochloropsis spp oil: Polaris; Starch: Roquette; Vitamin premix (IU or mg kg⁻¹ diet): DL-a tocopherol acetate 60 IU, sodium menadione bisulphate 5 mg, retinyl acetate 15,000 IU, DL-cholecalciferol 3000 IU, thiamin 15 mg, riboflavin 30 mg, pyridoxine 15 mg, B12 0.05 mg, nicotinic acid 175 mg, folic acid 500 mg, inositol 1000 mg, biotin 2.5 mg, calcium pantothenate 50 mg, choline chloride 2000 mg (UPAE, INRAE); Mineral premix (g or mg kg⁻¹ diet): calcium carbonate (40% Ca) 2.15 g, magnesium oxide (60% Mg) 1.24 g, ferric citrate 0.2 g, potassium iodide (75% I) 0.4 mg, zinc sulfate (36% Zn) 0.4 g, copper sulfate (25% Cu) 0.3 g, manganese sulfate (33% Mn) 0.3 g, dibasic calcium phosphate (20% Ca, 18% P) 5 g, cobalt sulfate 2 mg, sodium selenite (30% Se) 3 mg, KCl 0.9 g, NaCl 0.4 g (UPAE, INRAE).

2.3. Swimming challenge
186 2.3.1. Experimental setup

187 Swimming and metabolic performances were acquired on 18 fish per experimental
188 condition, using two swim tunnel respirometers in parallel (Loligo Systems, Tjele, Denmark)
189 filled with the same water as those in the fish tanks (Figure S1). For each tunnel, the
190 respirometer (10L) was composed of a swim chamber (40Lx10Hx10W cm) and a hydraulic
191 system that created a laminar flow thanks to a honeycomb structure placed upstream of the
192 swim chamber. An external motor generated the flow with a propeller and the experimenter
193 controlled the speed. Both motors were calibrated using a current meter before the
194 experiments. In both tunnels, a flush pump was used to maintain the oxygen level above
195 85% in the swim chamber. Black shelter on the side of the swim chamber and mild lightning
196 was used to avoid any disturbance of the animal during the experiment. Experiments were
197 continuously monitored using webcamts (Logitech C920s PRO) placed above the swim
198 chamber that were displayed live on a computer. Oxygen levels were measured using
199 oxygen probes (FireSting O₂, PyroScience, Germany) calibrated before each experiment and
200 placed inside the swim chamber. The probes monitored oxygen level and temperature every
201 second and data were acquired using the software PyroScience Workbench (V1.3.9.2140).

202 2.3.2. Swimming protocol

203 Before each experiment, two fish (one per experimental condition) were electronically
204 identified, isolated from their rearing tanks to be fasted for a minimum of 48h [35,48].
205 Following this fasting period, i.e. the day before the swimming test, after 4:00 pm, fish were
206 anesthetized with MS-222, measured in a bucket filled with seawater to further adjust the
207 current velocity in body length per second (BL s⁻¹). Fish were then transferred from the
208 acclimation tank to the swim chamber of the swim tunnel in a plastic bag filled with
209 seawater (without exposing the gills to the air) [49]. After a few minutes (~10 min) recovery
210 of the fish from the anaesthesia, fish were trained in the swim chamber for 10 min by
211 progressively increasing the velocity of the current. After training, the fish remained
212 overnight in the swim tunnel at a low velocity (0.5 BL s⁻¹). During all night, oxygen
213 consumption and temperature were recorded, and the oxygen level in the swim chambers
214 was maintained thanks to the flushing pump with cycles of 15 min of measurement and 5
215 min of flushing. The following morning, a step protocol experiment [39] was conducted to
measure swimming performance. Swimming performance was assessed by measuring the critical swimming speed ($U_{crit}$; the maximum velocity a fish can sustain aerobically; Brett, 1964) and the associated oxygen consumption. With intervals of 20 min, the current velocity was increased for 5 min following this protocol: increments of $1.5 \text{ BL s}^{-1}$ from 0.5 to $3.5 \text{ BL s}^{-1}$, then increments of $0.75 \text{ BL s}^{-1}$ until fish exhaustion.

During each swimming step of the protocol (20 min), the flushing pump was turned off and decreasing oxygen level corresponding to fish oxygen consumption was measured. During the speed-up steps (5 min), the flushing pump was turned on, allowing the restoration of oxygen levels that were never under 75%. Fish were considered exhausted when they failed to swim off the grid for 15 seconds after light stimulation. When the exhaustion was reached, the velocity of the current was immediately reduced to the recovery speed ($0.5 \text{ BL s}^{-1}$). The fish then entered a recovery period during which the oxygen consumption was measured, as previously described (i.e. 20 min measurement, 5 min flush) for 1 hour (= 3 measurements), to assess the fish recovery capacity in terms of excess post-oxygen consumption (EPOC). EPOC quantifies the energy and time required to restore tissue and cell storage of oxygen, metabolite balance, such as lactate and glycogen, and other metabolic functions such as ionic and osmotic balance [50].

After the recovery period, the fish was taken out of the swim chamber with a net, and immediately euthanized with MS-222 ($0.4 \text{ g L}^{-1}$). Its width, height, weight and length were measured, and white muscle (sampled on the left side, just anterior to the head), liver (without gallbladder), and brain were immediately sampled and flash frozen in liquid nitrogen for further fatty acid and gene expression analyses. Before and after each experiment, the background microbial respiration was measured for 1h in the swim chamber without fish. At the end of each experiment, the swim tunnel was fully cleaned with bleach tabs and rinsed three times with fresh water. The tunnels were then rinsed with and filled again with UV-treated sea water. During the swimming challenge, three fish (2 in the Control condition and 1 in the Depleted condition), apparently healthy, died overnight, during the recovery period before the swimming test. At the end of the experiment, 16 fish remained in the Control condition and 17 fish remained in the Depleted condition.

## 2.4. Calculation of metabolic and swimming performance variables
246 2.4.1. Growth performance

247 For each individual, the Specific Growth Rate (SGR; % per day; Lugert et al., 2016) was
248 calculated over the 5 months experimental period using the following equation (1):

$$
SGR = 100 \times \left( \frac{\log(W_{final}) - \log(W_{initial})}{D} \right) (1)
$$

249 Where W<sub>final</sub> is the weight at the end of the experiment, W<sub>initial</sub> is the weight at the beginning
250 of the experiment and D is the number of days between the disposal into experimental
251 condition and the end of the experiment (162-192 days).

252 Fulton condition factor (CF, arbitrary unit) [52] was also calculated over the 5 months of
253 conditioning using the following equation (2):

$$
CF = 100 \times \left( \frac{W}{TL^3} \right) (2)
$$

254 Where W is the weight (g) and TL is the total length (cm) of the fish.

255 2.4.2. Critical swimming speed

256 U<sub>crit</sub> (BL s<sup>-1</sup>) was calculated based on the following equation (3) [39]:

$$
U_{crit} = U_{max} + \frac{t_1}{t} \times U_1 (3)
$$

257 Where U<sub>max</sub> (BL s<sup>-1</sup>) is the highest velocity reached by the individual during a whole
258 swimming step (i.e. 20 min), t<sub>1</sub> (min) is the time the individual spent at the exhaustion
259 velocity, t (min) is the time of a complete swimming phase (i.e. 20 min), and U<sub>1</sub> (BL s<sup>-1</sup>) is the
260 increment of velocity before exhaustion of the fish (i.e. 1.5 or 0.75 BL s<sup>-1</sup>).

261 Correction for solid blocking effect was made for all fish [53], and the maximum cross
262 section area of the fish ranged from 7 to 14% of the swim section area for length ranging
263 from 17.6 to 23.4 cm.

264 2.4.3. Oxygen consumption

265 The MO₂ (mg O₂ kg⁻¹ h⁻¹) was calculated based on the equation (4) [54].
$$M_{O_2 \text{ meas}} = \left[ \left( \frac{(\Delta[O_2])}{\Delta t} - \frac{\Delta O_2}{\Delta t_{\text{bact}}} \right) \right] \times \left( \frac{v}{m} \right) \quad (4)$$

Where $\Delta[O_2]$ (mg O₂ L⁻¹) is the decrease in oxygen concentration relative to oxygen consumption measured in the swim chamber over time $\Delta t$ (hours), $v$ is the volume of the swim tunnel (10 L) minus the volume of the fish, and $m$ is the fish weight (kg). $\Delta[O_2]/\Delta t$ is obtained by plotting $[O_2]$ versus time to adjust a linear regression (Graphical Analysis 3.4, Beaverton, OR, USA). Only slopes with a coefficient of regression (R²) above 0.95 were considered. The slope corresponding to the microbial background respiration was calculated before and after each experiment, and the $\Delta[O_2]/\Delta t_{\text{bact}}$, which is the mean of these two measurements, was subtracted from the $MO_2$ measured with fish.

As the weight of an animal influences its respiratory metabolism, $MO_2$ was standardized with a standard fish mass of 100 g ($MO_2$) [55,56], with the following equation (5):

$$M_{O_2} = M_{O_2 \text{ meas}} \times \left( \frac{m}{m_{\text{corrected}}} \right)^{1-A} \quad (5)$$

Where $MO_{2meas}$ is the calculated oxygen consumption obtained with the equation (5), $m$ is the fish weight (kg), and $m_{corrected}$ is the standard mass (100 g). $A$ is the allometric exponent of the relationship between metabolic rate and fish mass. We used a value of 0.8 as it has been previously used for *Dicentrarchus labrax* [35], and commonly used in teleost species [57].

### 2.4.4. Metabolic rates

Standard Metabolic rate (SMR; mg O₂ kg⁻¹ h⁻¹) was extrapolated at the intercept (6) (i.e. $MO_2$ when $U = 0$ BL s⁻¹; Brett, 1964):

$$M_{O_2} = SMRe^{bU} \quad (6)$$

Where *SMR* is the intercept, *b* is a constant and *U* is the swimming speed (BL s⁻¹).

Maximal Metabolic Rate (MMR; mg O₂ kg⁻¹ h⁻¹) was estimated by taking the highest value of $MO_2$ during the swimming challenge [58].

Aerobic scope (AS; mg O₂ kg⁻¹ h⁻¹) was then calculated with the following equation (7) [54]:

$$AS = MMR - SMR \quad (7)$$
EPOC (mg O₂ kg⁻¹) was calculated by (1) subtracting the baseline value of MO₂ (i.e. the value of MO₂ measured just before starting the experiment) from the three MO₂ values calculated for the post exercise phase, then (2) integrating the area bounded between this new MO₂ values and time using Graphical Analysis vs 3.4 (Beaverton, OR, USA) [50]. EPOC is the energy and time needed to return metabolic processes to their resting state, following intense exercise [50].

The net cost of transport at maximum exercise (NCOT; mg O₂ kg⁻¹ m⁻¹) was calculated as in [59] following equation (8):

$$NCOT = \frac{(MMR - SMR)}{U_{crit}} \quad (8)$$

Where $U_{crit}$ is expressed in m min⁻¹, MMR and SMR in mg O₂ kg⁻¹ min⁻¹.

## 2.5. Fatty acid analysis

The fatty acid composition of liver, muscle and brain of all individuals tested in the swim tunnel ($n_{Control}=15$; $n_{Depleted}=16$) was measured. For each tissue, total lipids were extracted by grounding flash frozen tissues (whole liver, whole brain, and about 100 mg of muscle) into a homogeneous powder in liquid nitrogen, and by diving ~50 g of each tissue into a 6 mL mixture of CHCl₃/MeOH (2:1, v/v; Folch, 1957). The rest of the grounded liver was preserved at -80°C for further gene expression measurement (see 2.6). Although no initial sampling was conducted on the fish from this study, the fatty acid composition of juveniles from the same site and age class was characterized the previous year (Péron et al., 2024).

### 2.5.1. Lipid class separation

For each total lipid sample, extracted lipids were separated into neutral lipid (NL, corresponding to the reserve lipids) and polar lipid (PL, corresponding to the membrane lipids) fractions following the method described by Le Grand et al., (2014), and using an aliquot (¼) of the total lipid extract. Briefly, the lipid extract aliquot was evaporated to dryness under nitrogen, recovered with three washings of 0.5 mL of chloroform/methanol (98:2, v/v), and deposited at the top of a silica gel column (40 mm × 4 mm, silica gel 60 A 63–200 µm rehydrated with 6% H₂O (70–230 mesh)). NL were eluted with 10 mL of chloroform/methanol (98:2, v/v) and collected in a glass vial containing 2.3 µg of internal
315

standard (tricosanoic acid, C23:0). PL were then eluted with 20 mL of MeOH and collected in the same way as NL. Samples from both fractions were evaporated to dryness using a rotating evaporator (Genevac).

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### 2.5.2. Transesterification of neutral and polar lipid fractions

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NL and PL fractions were processed to obtain Fatty Acid Methyl Esters (FAME), following a protocol modified from Mathieu-Resuge et al., (2019). 1 mL of KOH/MeOH (0.5M, 5.6:94.4; v/v) was added. Samples were then heated at 80°C during 30 min. After cooling, 800 µL of H₂SO₄/MeOH (3.4%, v/v) was added and heated at 100°C for 10 min in a dry bath. After cooling, 800 µL of hexane and 1.5 mL of hexane-saturated distilled water were added to the mixture. After a homogenization and centrifugation at 738 g during 1 min at 20 °C, a removal of the aqueous phase was realized, the hexane phase, containing FAME, was washed two more times with hexane-saturated distilled water.

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### 2.5.3. FAME analysis

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FAME were analyzed in a Varian CP8400 gas chromatograph (GC; Varian Inc., CA, USA) coupled with a flame-ionization detector (FID) as described by Mathieu-Resuge et al., (2019). FAME were injected in split less-mode in parallel on two different columns (DBWAX 30 m × 0.25 mm ID x 0.2 µm and DB5 30 m × 0.25 mm ID x 0.2 µm, Agilent). Identification of FAME was realized by comparison of their retention time using those of commercial standards (Supelco 37 Component FAME Mix, PUFA No.1 and No.3, and Bacterial Acid Methyl Ester Mix, Sigma, MO, USA). Internal standard allowed to calculate FA concentration (µg mg⁻¹ WW). Fatty acid mass fraction was defined as the mass of each fatty acid to the total fatty acid content. We also calculated the fish tissue (percentage in PL)/diet ratio (percentage in total lipids) for DHA and EPA [63].

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## 2.6. Gene expression analysis

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Total RNA were extracted from the frozen liver powder (n=36) using Extract-all reagent (Eurobio; Courtaboeuf, Essonne, France) coupled with purification steps on a Nucleospin RNA column as described by Mazurais et al., (2020). The extraction protocol included one step of DNase treatment (Macherey-Nagel, Düren, Germany). Concentrations and purity of extracted RNA were measured using a ND-1000 NanoDrop spectrophotometer
(ThermoScientific Inc., Waltham, MA, USA). A 4150 TapeStation System (Agilent Technologies Inc, Santa Clara, CA, USA) was used to evaluate the RNA integrity (RIN) and only samples having a RIN ≥ 8 were used (Control n= 16; Depleted n = 14). For each sample, two positive and one negative reverse transcription (RT) reactions for cDNA synthesis were performed using iScript cDNA Synthesis kit (Bio-Rad Laboratories Inc., Hercules, CA, USA), as described in Mazurais et al., (2020). The levels of relative expression of five genes of interest were investigated by qPCR using a CFX96 Touch Real-Time PCR Detection system (Bio-Rad Laboratories Inc.). The relative expression levels of five genes involved in fatty acid metabolism were investigated by qPCR using a CFX96 Touch Real-Time PCR Detection System (Bio-Rad Laboratories Inc.). We chose five different genes of interest in this study. First, fatty acid desaturase 2 (fads2), elongation of very long-chain fatty acid 6 (elovl6) and stearoyl-CoA desaturase (scd1b) were chosen for their role in the biosynthesis of LC PUFA, though they are not specific in biosynthesising DHA (Castro et al., 2016; Geay et al., 2010; Ntambi and Miyazaki, 2004). Secondly, Fatty acid synthase (fas) is involved in lipogenesis and lipolysis processes and is known to be influenced by DHA/EPA ratio [67] and lipoprotein lipase (lppl) plays a role in the regulation of body lipid content [68].

Expression levels were normalized to three reference genes (elongation factor 1-alpha (ef1), beta-actin (actin), and ribosomal protein L13a (l13a)) which met the stability requirements of the CFX Maestro software (average M value = 0.68). Primers were designed from available *D. labrax* mRNA sequences on GenBank to obtain PCR products ranging from 150 to 200 bp using Primer3 software (Table S1). The specificity of amplicons was confirmed via melting curve analysis. Relative transcript levels were calculated using the ΔΔCt method [69] with the control samples automatically selected by the software.

## 2.7. Statistical analysis

All analyses were performed using RStudio (v.4.2.1). The effect of dietary treatment on growth performance, metabolic variables (SMR, MMR, AS, EPOC and NCOT), FA mass fraction, gene expression were tested with linear mixed models (*lme4* package, *lmer* function). If necessary (i.e non-normal residuals), data were transformed to log (gene expression data). In all linear models, we used dietary treatment (i.e. two levels: Depleted and Control) as a fixed factor, and the tank number as a random factor. The significance of models was tested using type II ANOVA after ensuring that the assumptions were met
(package performance, check_model function). Finally, the correlations between DHA mass fraction and growth performance were tested using Spearman's correlation. Results with p-values less than 0.05 were considered statistically significant and led to rejection of the null hypothesis.

## 3. Results

### 3.1. Growth performance

The results concerning growth performance are summarized in Table 2. No visual differences in feeding behaviour were observed between groups throughout the experiment. The SGR calculated over the 162-192 days of experiment was significantly two times lower in the depleted groups compared to the Control groups (Table 2). Three individuals from the Depleted condition had negative SGR but none in the Control condition. However, neither TL nor weight, all calculated at the end of the experiment, were significantly affected by the dietary treatment (p>0.05). Fulton's K showed a non-significant trend (p=0.05). In the depleted group, the SGR was negatively correlated with the DHA mass fraction in muscle, but not with those in liver or brain (Figure 1).

<table><caption>Table 2: Growth performance and condition factor of European seabass juveniles fed the Control diet (n=15) or the Depleted diet (n=16). SGR: Specific Growth Rate (% per day) calculated over the experimental period (i.e. 162-192 days); CF: Condition Factor (Fulton's K, Arbitrary Unit AU), TL: Total Length (cm) and final weight (g) calculated at the end of the experimental period (i.e. at 162-192 days). Data are presented as mean ± Standard Error of the Mean (SEM). Statistical difference is expressed by the p-value obtained from an Anova on the mixed linear models.</caption><thead><tr><th></th><th>Control group</th><th>Depleted group</th><th>p-value</th><th>Df</th><th>F value</th></tr></thead><tbody><tr><td>SGR (% per day)</td><td>0.1 ± 0.0</td><td>0.05 ± 0.0</td><td>0.04</td><td>1</td><td>4.6</td></tr><tr><td>Final CF (AU)</td><td>1.2 ± 0.0</td><td>1.1 ± 0.0</td><td>0.05</td><td>1</td><td>4.1</td></tr><tr><td>Final TL (cm)</td><td>21.1 ± 0.3</td><td>20.8 ± 0.4</td><td>0.59</td><td>1</td><td>0.3</td></tr><tr><td>Final Weight (g)</td><td>114.9 ± 5.6</td><td>105.9 ± 6.9</td><td>0.32</td><td>1</td><td>1.0</td></tr></tbody></table>
<figure><img src="image_3.png" /><figcaption>Figure 1: Spearman correlation between Specific Growth Rate (SGR) and proportion of DHA in polar lipids (PL) of muscle. The grey regression line represents the Control condition (n=15) and the black regression line represents the Depleted condition (n=16).</figcaption></figure>

## 3.2. Fatty acid composition

### 3.2.1. Fatty acid composition in PL

In PL of all tissues (Table 3), the main FA groups (SFA, MUFA, PUFA) were present in similar quantities in both Control and Depleted dietary groups ($p>0.05$).

The DHA quantities in PL were lower only in the liver of the Depleted fish compared to the Control fish. In the liver, DHA quantities were 37% lower in the Depleted group compared to the control group. DHA concentrations tended to be lower in the brain and muscle of Depleted fish, but the differences were not statistically significant. However, when considering the proportion of DHA (% of DHA on total fatty acids), all organs of Depleted fish showed lower DHA proportions compared to the control (Table S2, S3, and S4). The EPA proportions in PL were higher in the Depleted group compared to the Control group only in the liver. This resulted in a twofold higher EPA/DHA ratio in the liver of the Depleted group compared to the Control group (Table 3).
412 The arachidonic acid (ARA, 20:4n-6) proportions were higher in the Depleted group in both
413 the muscle and the liver, but not in the brain. However, 18:2n-6 proportions were similar
414 between groups despite the dietary differences (Table 3).

415 3.2.1. Fatty acid composition in NL

416 Concerning NL (Table 4), the fatty acid concentrations in the muscle showed no significant
417 difference and displayed a higher variability compared to PL. On the contrary, the DHA
418 proportions in NL were very reflective to those of the diet (Table S5).

419 Although DHA was present at only 0.5% in the Depleted diet, it was present in the NL and PL
420 of the three tissues, with higher proportions in the PL than in the NL.

421 The ratio of DHA and EPA between tissue and diet is shown in Figure 3. A ratio > 1 indicates
422 that the FA is selectively retained or synthesised in the tissue, reflecting that the dietary
423 availability of this FA was below the physiological requirement [63,70]. A ratio equal to one
424 indicates that the supply of this FA meets the physiological requirement. For both
425 experimental groups the tissue/diet DHA ratio was >1 and was more than 30 times higher for
426 the Depleted group compared to the Control group (Figure 3A). For the Depleted group, the
427 brain had the highest ratio and liver had the lowest, but these ratios were still very high
428 (37.9 and 32.8, respectively). EPA ratio was > 1 in the liver and muscle, but < 1 in the brain
429 (Figure 3B).

<table><caption>Table 3: Fatty acid profiles of Polar Lipids (µg of FA per mg of tissue) in the liver, muscle and brain of European
sea bass juveniles fed the Control (n=15) and the DHA Depleted (n=16) diets. PL: polar lipids. Data are
presented as mean ± SEM. Statistical significance is indicated by values in bold.</caption><thead><tr><th rowspan="2">PL FA</th><th colspan="2">Liver</th><th colspan="2">Muscle</th><th colspan="2">Brain</th></tr><tr><th>Control</th><th>Depleted</th><th>Control</th><th>Depleted</th><th>Control</th><th>Depleted</th></tr></thead><tbody><tr><td>16:0</td><td>1.69 ± 0.05</td><td>1.65 ± 0.09</td><td>0.86 ± 0.05</td><td>0.93 ± 0.02</td><td><strong>5.10 ± 0.12</strong></td><td><strong>4.68 ± 0.15</strong></td></tr><tr><td>18:0</td><td>1.09 ± 0.04</td><td>1.17 ± 0.1</td><td>0.32 ± 0.01</td><td>0.32 ± 0.01</td><td>2.73 ± 0.05</td><td>2.53 ± 0.06</td></tr><tr><td>24:0</td><td>-</td><td>-</td><td>-</td><td>-</td><td>0.50 ± 0.02</td><td>0.50 ± 0.02</td></tr><tr><td>Σ SFA</td><td>2.94 ± 0.11</td><td>2.97 ± 0.14</td><td>1.24 ± 0.05</td><td>1.31 ± 0.03</td><td>8.66 ± 0.20</td><td>8.0 ± 0.22</td></tr><tr><td>16 :1n-7</td><td>0.17 ± 0.02</td><td>0.21 ± 0.02</td><td>0.06 ± 0.01</td><td>0.7 ± 0.01</td><td>0.73 ± 0.03</td><td>0.70 ± 0.03</td></tr><tr><td>16 :1n-9</td><td>0.08 ± 0.0</td><td>0.08 ± 0.0</td><td>-</td><td>-</td><td>0.44 ± 0.01</td><td>0.43 ± 0.01</td></tr><tr><td>18 :1n-7</td><td>0.26 ± 0.01</td><td>0.27 ± 0.01</td><td>0.10 ± 0.0</td><td>0.10 ± 0.0</td><td>0.44 ± 0.01</td><td>0.41 ± 0.01</td></tr></tbody></table>
<table><tr><td>18 :1n-9</td><td>1.61 ± 0.09</td><td>1.75 ± 0.11</td><td>0.61 ± 0.03</td><td>0.62 ± 0.02</td><td>5.93 ± 0.19</td><td>5.60 ± 0.18</td></tr><tr><td>20 :1n-9</td><td>0.08±0.01</td><td>0.09±0.01</td><td>0.03±0.0</td><td>0.03±0.0</td><td>0.40 ± 0.02</td><td>0.39 ± 0.01</td></tr><tr><td>24 :1n-9</td><td>-</td><td>-</td><td>-</td><td>-</td><td>1.39 ± 0.05</td><td>1.40 ± 0.05</td></tr><tr><td>Σ MUFA</td><td>2.26 ± 0.11</td><td>2.48 ± 0.14</td><td>0.84 ± 0.04</td><td>0.85 ± 0.02</td><td>9.62 ± 0.31</td><td>9.21 ± 0.29</td></tr><tr><td>18:2n-6</td><td>0.46 ± 0.02</td><td>0.54 ± 0.05</td><td>0.22 ± 0.02</td><td>0.24 ± 0.01</td><td>0.16 ± 0.01</td><td>0.15 ± 0.01</td></tr><tr><td>18:3n-3</td><td>0.07 ± 0.0</td><td>0.09 ± 0.0</td><td>0.03 ± 0.0</td><td>0.04 ± 0.0</td><td>0.02±0.0</td><td>0.02±0.0</td></tr><tr><td>20 :4n-6</td><td><b>0.37 ± 0.02</b></td><td><b>0.56 ± 0.03</b></td><td><b>0.11 ± 0.01</b></td><td><b>0.15 ± 0.01</b></td><td>0.98 ± 0.03</td><td>0.99 ± 0.03</td></tr><tr><td>20 :5n-3</td><td><b>1.15 ± 0.04</b></td><td><b>1.59 ± 0.08</b></td><td>0.60 ± 0.04</td><td>0.71 ± 0.03</td><td>1.06 ± 0.04</td><td>1.07 ± 0.04</td></tr><tr><td>22 :5n-3</td><td>0.17 ± 0.01</td><td>0.15 ± 0.01</td><td>2.4 ± 0.0</td><td>2.0 ± 0.1</td><td>0.24 ± 0.01</td><td>0.29 ± 0.01</td></tr><tr><td>22 :6n-3</td><td><b>2.35 ± 0.06</b></td><td><b>1.71 ± 0.10</b></td><td>0.88 ± 0.05</td><td>0.84 ± 0.04</td><td>6.26 ± 0.19</td><td>5.51 ± 0.22</td></tr><tr><td>Σ PUFA</td><td>4.93 ± 0.13</td><td>4.96 ± 0.23</td><td>2.10 ± 0.11</td><td>2.22 ± 0.05</td><td>9.38 ± 0.3</td><td>8.70 ± 0.27</td></tr><tr><td>Σ LC PUFA</td><td>4.19 ± 0.11</td><td>4.11 ± 0.19</td><td>1.74 ± 0.09</td><td>1.85 ± 0.05</td><td>8.73 ± 0.24</td><td>8.08 ± 0.25</td></tr><tr><td>Σ n-6</td><td><b>1.0 ± 0.03</b></td><td><b>1.26 ± 0.07</b></td><td>0.40 ± 0.02</td><td>0.47 ± 0.01</td><td>1.21 ± 0.03</td><td>1.27 ± 0.04</td></tr><tr><td>Σ n-3</td><td>3.83 ± 0.11</td><td>3.59 ± 0.17</td><td>1.64 ± 0.09</td><td>1.71 ± 0.05</td><td>7.68 ± 0.23</td><td>6.98 ± 0.26</td></tr><tr><td>Σ DMA</td><td>-</td><td>-</td><td>-</td><td>-</td><td>2.25 ± 0.12</td><td>2.37 ± 0.11</td></tr><tr><td>EPA/DHA</td><td><b>0.5 ± 0.0</b></td><td><b>1.0 ± 0.1</b></td><td><b>0.70 ± 0.04</b></td><td><b>0.88 ± 0.06</b></td><td>0.17 ± 0.0</td><td>0.2 ± 0.0</td></tr></table>

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<table><caption>Table 4: Fatty acid profiles of Neutral Lipids (µg of FA per mg of tissue) in the muscle of European sea bass juveniles fed the Control (n=15) and the DHA Depleted (n=16) diets. NL: neutral lipids. Data are presented as mean ± SEM. Statistical significance is indicated by the p-value.</caption><thead><tr><th rowspan="2">NL FA</th><th colspan="2">Muscle</th></tr><tr><th>Control</th><th>Depleted</th></tr></thead><tbody><tr><td>14:0</td><td>0.37 ± 0.08</td><td>0.22 ± 0.04</td></tr><tr><td>16:0</td><td>2.53 ± 0.58</td><td>1.88 ± 0.32</td></tr><tr><td>18:0</td><td>0.51 ± 0.10</td><td>0.39 ± 0.07</td></tr><tr><td>Σ SFA</td><td>3.52 ± 0.79</td><td>2.56 ± 0.44</td></tr><tr><td>16 :1n-7</td><td>0.79 ± 0.21</td><td>0.69 ± 0.11</td></tr><tr><td>18 :1n-7</td><td>0.42 ± 0.10</td><td>0.30 ± 0.05</td></tr><tr><td>18 :1n-9</td><td>4.18 ± 1.03</td><td>3.27 ± 0.56</td></tr><tr><td>20 :1n-9</td><td>0.23 ± 0.05</td><td>0.18 ± 0.03</td></tr><tr><td>Σ MUFA</td><td>5.96 ± 1.45</td><td>4.67 ± 0.78</td></tr><tr><td>18 :2n-6</td><td>1.33 ± 0.31</td><td>0.96 ± 0.18</td></tr><tr><td>18 :3n-3</td><td>0.23 ± 0.05</td><td>0.18 ± 0.03</td></tr><tr><td>20 :4n-6</td><td>0.07 ± 0.01</td><td>0.07 ± 0.01</td></tr><tr><td>20 :5n-3</td><td>0.88 ± 0.19</td><td>0.70 ± 0.12</td></tr><tr><td>22 :5n-3</td><td>0.14 ± 0.03</td><td>0.08 ± 0.02</td></tr><tr><td>22 :6n-3</td><td>0.54 ± 0.12</td><td>0.29 ± 0.06</td></tr><tr><td>Σ PUFA</td><td>3.83 ± 0.87</td><td>2.67 ± 0.47</td></tr><tr><td>Σ LC PUFA</td><td>1.75 ± 0.38</td><td>1.20 ± 0.20</td></tr><tr><td>Σ n-6</td><td>1.52 ± 0.35</td><td>1.13 ± 0.21</td></tr><tr><td>Σ n-3</td><td>1.99 ± 0.43</td><td>1.35 ± 0.23</td></tr><tr><td>EPA/DHA</td><td>1.61 ± 0.11</td><td>2.5 ± 0.18</td></tr></tbody></table>
<figure><img src="image_4.png" /></figure>

Figure 2: Ratio of FA between tissue and diet (± SEM) for DHA (A) and EPA (B) of European sea bass juveniles fed either a control or a DHA depleted diet. The grey bars represent the Control condition (n=15) and the black bars represent the Depleted condition (n=16). The horizontal dotted line represents the 1:1 ratio between the tissue and the diet. A ratio greater than 1 indicates that the FA is selectively retained or synthesised in the tissues, indicating that the dietary availability of this FA was below the physiological requirement. When this ratio is equal to one, it indicates that the supply of this FA meets the physiological requirement.

442 3.3. Swimming and metabolic performance

443 As expected, the MO₂ increased exponentially with the swimming speed for both dietary

444 treatments (Figure 4). The metabolic variables (SMR, MMR, AS, EPOC and NCOT) and Ucrit

445 were not affected by the dietary treatment (p>0.05; results summarized in Table 5).

446
<figure><img src="image_5.png" /><figcaption>Figure 3: Oxygen consumption (MO₂, mg O₂ kg⁻¹ h⁻¹, ± SEM) as a function of swimming speed (body length BL s⁻¹) for both dietary groups for European sea bass juveniles fed either a control or a DHA depleted diet. The Control group is represented by circles in grey (n=14) and the Depleted group is represented by the triangles in black (n=16). Both lines are exponential curves fitted on MO₂ as a function of swimming speed. (Control group curve: y=32.833e⁻⁰.³⁰⁵⁹x; r² = 0.99; Depleted group curve: y=46.094e⁻⁰.²³⁵⁶; r²=0.99).</figcaption></figure>

<table><caption>Table 5: Metabolic and swimming performance for European sea bass juveniles fed either a Control diet or a DHA Depleted diet. Data are standard metabolic rate (SMR, mg O₂ kg⁻¹ h⁻¹), maximum metabolic rate (MMR, mg O₂ kg⁻¹ h⁻¹), aerobic scope (AS, mg O₂ kg⁻¹ h⁻¹), excess post-oxygen consumption (EPOC, mg O₂ kg⁻¹), critical swimming speed (Ucrit, body length BL s⁻¹), and net cost of transport (mg O₂ kg⁻¹ m⁻¹). Data are presented as mean ± SEM, n is the number of individuals statistically tested.</caption><thead><tr><th>Dietary Condition</th><th>SMR</th><th>MMR</th><th>AS</th><th>EPOC</th><th>U<sub>crit</sub></th><th>NCOT</th></tr></thead><tbody><tr><td>Control</td><td>27.1 ± 0.4<br>(n=14)</td><td>165.8 ± 1.8<br>(n=14)</td><td>138.6 ± 1.7<br>(n=14)</td><td>1623.5 ±<br>41.7 (n=14)</td><td>4.8 ± 0.1<br>(n=14)</td><td>2.3 ± 0.1<br>(n=14)</td></tr><tr><td>Depleted</td><td>27.0 ± 0.5<br>(n=16)</td><td>155.3 ± 1.6<br>(n=16)</td><td>128.2 ± 1.5<br>(n=16)</td><td>1463.4 ±<br>32.8 (n=16)</td><td>4.7 ± 0.1<br>(n=16)</td><td>2.2 ± 0.1<br>(n=16)</td></tr></tbody></table>
<table><tr><td>p-value</td><td>0.81</td><td>0.38</td><td>0.26</td><td>0.43</td><td>0.49</td><td>0.55</td></tr><tr><td>F value</td><td>0.07</td><td>0.9</td><td>1.4</td><td>0.62</td><td>0.48</td><td>0.02</td></tr></table>

459

460

### 3.4. Gene expression

461

462

463

464

The relative expression of *fads2*, *fas* and *lpl* transcripts measured in the liver were significantly higher in the Depleted fish than in the Control fish (p<0.01, Figure 5). However, the relative gene expression of *elovl6* and *scd1b* were not significantly different between the two dietary groups (Figure 5).

465

466

467

Significant negative correlations were found between the hepatic DHA proportions and the relative expressions of *fads2*, *fas*, *lpl* and *scd1b* for the Depleted group, but not for the Control one (Figure 6).

<figure><img src="image_6.png" /></figure>

468

469

470

471

<caption>Figure 4: Relative expression of genes (± SEM) coding for enzymes involved in lipid metabolism and measured in the liver of European sea bass juveniles fed either a Control (n=16) or a DHA Depleted (n=14) diet. *elovl6*: fatty acid elongase 6, *fads2*: fatty acid desaturase 2, *fas*: fatty acid synthase, *lpl*: lipoprotein lipase, *scd1b*:</caption>
472 stearoyl-CoA desaturase. Asterisks represent statistical significance between the two dietary conditions. Linear

473 mixed model. ** p<0.01.

<figure><img src="image_7.png" /><figcaption>Figure 5: Spearman correlation between the DHA content in polar lipid (PL) of liver and the relative transcript expressions of *fads2* (A), *fas* (B), *lpl* (C) and *scd1b* (D). The grey regression line represents the Control condition (n=16) and the black regression line represents the Depleted condition (n=14).</figcaption></figure>

474

475

476

477

478

## 4. Discussion

479 We investigated the effects of a long-term (5 months) dietary DHA limitation on wild-caught

480 European sea bass juveniles for somatic growth, FA composition in tissue, swimming

481 performance and metabolic rates. The DHA proportions in the tissues of DHA Depleted fish

482 were significantly reduced compared to the Control fish, although they remained high

483 considering the extreme DHA limitation in their diet. The dietary DHA limitation negatively

484 affected growth, but did not modulate swimming performance nor metabolic rates. We

485 found high inter-individual variations in all measured traits, suggesting phenotypic plasticity.

486 4.1. Consequences of dietary DHA limitation on tissue FA proportion, gene

487 expression and fish growth

488
Over the six-month conditioning period, no mortality was observed, and the condition index increased or remained stable, reaching 1.2 in the control group and 1.1 in the DHA-depleted group—values higher than those typically reported in wild individuals (≈1.0; Péron et al., 2024). Growth performance was consistent with known values for this species at comparable temperatures [72]. The fatty acid composition of liver mirrored that of the diets, confirming efficient assimilation of dietary lipids. DHA levels in muscle and liver of control fish were within the range reported for wild individuals [71], while brain DHA in the depleted group was 11% lower than in the same cohort sampled the previous year but remained within the inter-estuarine range observed in wild populations. The muscle and liver DHA proportions found in the Depleted fish was 17 and 40% lower, respectively than what has been found in wild fish from the same population which suggests that our DHA depleted diet effectively induced tissue DHA modulation [71].

The fatty acid composition of fish tissue reflected that of their diet, especially for LC PUFA. In particular, the dietary DHA deficiency was reflected in the concentrations of the liver.

In brain, results show that despite slightly more elevated ARA levels and lower DHA proportions, the overall LC PUFA content in PL remained relatively stable, suggesting that brain lipid homeostasis might be more resistant to dietary depletion than liver and muscle [3].

The calculation of the PL tissue/diet DHA ratio, which was well above one in all tissues of the depleted group, suggests that DHA requirements were not met in the fish fed the Depleted diet. When the value of this ratio is above 1, it suggests that the availability of these FA were below the physiological needs of fish and that it occurs either a selective retention in the tissue, or a synthesis of these FA. Hence, it suggests that a selective retention or an internal synthesis of DHA occurred in their tissues in order to compensate for the dietary deficiency [70]. However, this ratio does not account for the tissue growth. Since fish tissues accumulated lipids previously under an unknown and presumably DHA-rich diet in the wild, the elevated tissue/diet DHA ratio observed in the Depleted group may reflect both selective retention of DHA and the carry-over effects of previous dietary exposure, rather than de novo synthesis alone. This is particularly evident in muscle, where DHA concentrations were similar between Depleted and Control fish despite severe differences in diet, suggesting a
physiological regulation and possible redistribution from other organs. The ratio was also above one in the Control group, suggesting that this diet may have marginally undersupplied DHA relative to physiological demands under the given experimental conditions (e.g., lower temperature, wild origin), compared to established requirements from aquaculture trials [47]. In addition, a notable inter-individual variability in DHA proportions and concentrations within each condition group was observed. This suggests that individual fish differ in their capacity to retain or synthesize DHA, despite experiencing the same dietary constraints, pointing to underlying physiological plasticity. Some individuals from the Depleted condition had higher tissue DHA proportions and quantities than some individuals in the Control group, despite a tenfold lower DHA level in their diet. This inter-individual variability indicates a complex regulation of DHA in the tissue membranes that could relate on a multitude of levels such as retention, synthesis or a difference in the pre-experimental diet. High individual variations of n-3 LC PUFA proportions in membrane were previously observed in muscle of golden grey mullets fed high (17.5%) and low (1.2%) levels of EPA+DHA [73]. Contrary to DHA, tissue/diet EPA ratio showed a tissue specific response. In both experimental groups, it was equal or above 1 in muscle and liver, but was below 1 (at 0.5) in the brain. This suggests that EPA supply in both diets met the fish requirements and that this FA did not need to be retained as preferentially as DHA in the targeted organs. This result could also suggest that EPA might be used to synthesize DHA in the brain [63] or catabolized, as observed in mammals [74].

The hypothesis of DHA synthesis in Depleted fish is supported by the upregulation, in liver, of the expression of *fads2*, involved in n-3 LC PUFA metabolism, as well as by the negative correlation between gene expressions and hepatic DHA content observed in this group. However, this correlation may be driven primarily by a single point at the lower range of the DHA proportion. Notably, *fads2* encodes the delta6 desaturase, which is the first enzyme involved in the biosynthesis of n-3 LC PUFA and is considered as a limiting step of further biosynthesis [75]. Even if *fads2* is not specific to DHA synthesis, this upregulation could potentially indicate an internal conversion of DHA from precursors in the liver to maintain the levels in the tissue. However, despite the fact that upregulation of *fads2* has already been observed in European sea bass fed low levels of n-3 LC PUFA, this upregulation was not correlated with FADS2 enzymatic activity (Geay et al., 2010), which makes it unlikely that
significant DHA have been synthesized in our study. The lower DHA levels yet observed in the liver of Depleted fish compared to Control fish could suggest either (i) that FADS2 activity and fads2 expression were not necessarily related, or (ii) that FADS2 activity resulting from fads2 upregulation was not sufficient to induce compensating fatty acid synthesis in the Depleted fish. The first hypothesis could be explained by a regulation at the post-transcriptional level, involving notably microRNA, inhibiting LC PUFA biosynthesis as it was recently shown in Rabbit fish *Signatus canaliculatus* [76]. The same assumption can be applied to FAS activity, as it remained stable regardless of dietary variations [77,78]. The endogenous biosynthesis of DHA could be further confirmed by using stable isotopes as it was used in previous studies [79,80].

In addition to be associated with an upregulation of *fads2*, the DHA depleted diet was associated with a lower growth rate and induced a higher inter-individual variability in growth rate between individuals. This pattern was not uniform across individuals, highlighting that growth responses to DHA depletion are modulated by individual-level variability in physiological adjustments. While the ingestion rate has not been measured in this study, fish were daily observed during their feeding time, and no differences of ingestion rates, that could partly explain difference in growth rates, were visually observed between the two groups. Interestingly, in the depleted group, higher the DHA proportion in muscle, lower the growth rate (SGR). Yet, DHA has been shown to play a key role in fish growth [81]. Thus, this counterintuitive relationship between growth and DHA content would suggest an energetic cost of the physiological acclimation of individuals to a low dietary DHA content [70, 82, 83]. The upregulation of biosynthesis pathways observed at molecular level in the depleted group to synthetize DHA trying to counteract the dietary depletion may be too costly and prevent fish to allocate energy to grow. Such divergent patterns of growth and tissue composition further support the presence of trade-offs in energy allocation that may vary across individuals.

## 4.2. Consequences of dietary DHA depletion on swimming and metabolic performance

Despite the lower growth rate observed and the hypothesized higher energy expenditure in the Depleted fish to counteract the dietary deficiency, metabolic rates and maximum aerobic swimming performance were similar between the two dietary groups. This suggests
that some cellular mechanisms may mask or offset the expected energetic costs. Similar MMR between fish fed either a low n-3 LC PUFA diet or a control diet has been recently reported in Atlantic salmon [70]. The authors suggested that, even though the deficiency in n-3 LC PUFA decreased mitochondrial ATP production, mitochondrial metabolism could show a resilience to low levels of n-3 LC PUFA as this did not affect the MMR of fish [70]. In addition, as mentioned earlier, our study showed that some individuals from the Depleted condition had DHA levels in the range of those measured in Control individuals. This might result from the ability of some individuals to physiologically adjust their performance at the metabolic level without obvious relationship with the activation of the DHA biosynthesis pathways. Indeed, the metabolic rate is an integrative trait depending on multiple factors (ventilation, oxygen transport, mitochondrial performance), that could have been affected without being directly visible on the measured oxygen consumption. Individuals might also use different energy sources to fuel their needs. For example, fish fed the Depleted DHA diet might have used MUFA to sustain their swimming performance, as these FA were higher in the depleted diet [36,84]. While the high EPA level in the DHA Depleted diet did not avoid the decreased growth of Depleted fish, this might have balanced out the differences in metabolic rates. Indeed, no study investigated the effects of EPA and DHA separately but it is known that a diet rich in EPA+DHA can lower fish SMR [84]. Therefore, there might be an influence of EPA (and not DHA) on the activity of cellular ionic pumps that could regulate fish metabolic rates [85,86]. Altogether, our findings suggest that the preserved swimming performance in DHA-depleted fish may result from a complex interplay of compensatory mechanisms that buffer the expected functional consequences of altered DHA metabolism. This includes the potential role of alternative FA substrates such as MUFA or EPA, and inter-individual variability in the capacity to maintain cellular function despite dietary DHA limitations.

## 5. Conclusions

To our knowledge, this study is the first to examine, in an ecological context, the effects of a dietary limitation specifically in DHA, not combined with EPA, on the metabolic and swimming performance of wild-caught fish. DHA dietary limitation resulted in selective DHA retention or synthesis in fish tissues, the upregulation of biosynthetic pathways for DHA
synthesis at the molecular level, and a reduced growth rate of wild European sea bass
611
juveniles. However, the activation of these biosynthetic pathways did not fully compensate
612
for the DHA deficiency, suggesting a metabolic cost associated with DHA synthesis. The
613
strong association between higher DHA proportions and lower growth rates supports the
614
hypothesis that DHA is required primarily for anabolic processes during somatic growth,
615
whereas swimming and respiration, being catabolic processes, likely do not directly rely on
616
DHA availability.
617

Despite the dietary DHA limitation, metabolic rates and swimming performance remained
618
unchanged between groups, indicating that the isocaloric nature of the diets was sufficient
619
to sustain instantaneous metabolic activity regardless of n-3 LC PUFA availability. This
620
highlights the resilience of European sea bass to dietary changes, potentially due to
621
metabolic plasticity and the ability to mobilize alternative lipid sources for energy
622
production. Finally, considerable inter-individual variability was observed across all
623
measured variables (somatic growth, metabolic rates, Ucrit), further supporting the existence
624
of strong phenotypic plasticity. Investigating the underlying energetic trade-offs at the
625
individual level and their implications for fish growth and performance would provide
626
further insights into the adaptive responses to dietary DHA limitation.
627

628 Acknowledgment:

Authors thank Emmanuel Dubillot for his help to set the swim tunnels, Polaris for the
629
*Nannochloropis* spp. oil supply, the NOURDEM team (M. Drogou, S. Martin, L. Le Ru from
630
Ifremer Brest) for their help in fishing individuals in the Seine estuary and José Luis
631
Zambonino Infante for the formulation of the diets.
632

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