Meat Quality of European Flat Oyster Cultivated at Different Distances From Finfish Cages in a Mediterranean Integrated Multi‐Trophic Aquaculture (IMTA) System

E. Batır; M. Yıldız; Ö. Metin; G. Papini; D. Pensa; M. Magdy; L. Grosso; A. Fianchini; İ. Aydın; M. Rampacci; A. Rakaj

PMC · DOI:10.1155/anu/8152399·February 23, 2026

Meat Quality of European Flat Oyster Cultivated at Different Distances From Finfish Cages in a Mediterranean Integrated Multi‐Trophic Aquaculture (IMTA) System

E. Batır, M. Yıldız, Ö. Metin, G. Papini, D. Pensa, M. Magdy, L. Grosso, A. Fianchini, İ. Aydın, M. Rampacci, A. Rakaj

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TL;DR

Oysters grown farther from fish cages in an aquaculture system had better nutritional quality due to less waste and more phytoplankton.

Contribution

The study introduces spatial planning in IMTA systems to optimize oyster nutritional quality through seasonal and spatial variability.

Findings

01

Oysters cultivated 800 m from fish cages had higher crude protein content (up to 7.81%) compared to those at 20 m.

02

Arginine and lysine were the most abundant essential amino acids, with higher arginine levels observed at the distant site during late autumn and early spring.

03

Crude lipid levels peaked in March at 1.31%, linked to prereproductive nutrient accumulation in oysters.

Abstract

This study examines the nutritional profile of European flat oysters (Ostrea edulis) cultivated in an Integrated Multi‐Trophic Aquaculture (IMTA) system in Gaeta, Italy. Oysters were deployed for 257 days, 8 July 2023–21 March 2024, at two distances from gilthead seabream (Sparus aurata) cages: 20 m, representing high exposure to fish‐derived waste, and 800 m, reflecting reduced waste influence and greater phytoplankton availability. The objective was to evaluate how proximity to fish cages, combined with seasonal variability, influences oyster nutritional quality and to identify optimal spatial arrangements in IMTA systems. Sampling occurred in summer, autumn, winter, and spring to capture seasonal changes in physiology and nutrition. Oysters at the distant site showed consistently higher nutritional quality, attributed to greater phytoplankton access and dilution of fish waste. Crude…

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Water temperature, dissolved oxygen, chlorophyl‐a, pH, and salinity values in the experimental area.

Tables6

Table 1. Proximate composition (%) and energy parameters (KJ/g) of commercial feed used in the fish farm.

Proximate composition (%) and energy (KJ/g)	% of feed
Dry matter	93.41
Crude protein	45.55
Crude lipid	15.20
Ash	8.80
Fiber	1.80
Nitrogen‐free extracts^a	22.06
Metabolizable energy (KJ/g)^b	14.00
Gross energy (KJ/g)^c	20.57

Table 2. Fatty acid composition of commercial feed used in the fish farm (g/100 g fatty acid).

Fatty acids	g/100 g fatty acids
14:0	0.74
15:0	0.10
16:0	16.60
17:0	0.15
18:0	4.80
20:0	0.25
24:0	ND
16:1n‐7	3.03
18:1n‐9	24.43
20:1n‐9	0.66
22:1n‐11	0.06
18:2n‐6	13.51
20:2n‐6	0.18
18:3n‐3	3.26
20:3n‐3	0.08
20:4n‐6	0.24
20:5n‐3	7.15
22:6n‐3	9.18
ΣSFAs^a	22.64
ΣMUFAs^b	28.18
ΣPUFAs^c	33.60
Σn‐3 HUFAs	16.41
Σn‐3	19.67
Σn‐6	13.93
n‐3/n‐6	1.41
DHA/EPA	1.28

Table 3. Amino acid composition of commercial feed used in the fish farm (g/100 g protein).

Amino acids	g/100 g protein
Essential amino acids (EAAs)
Arginine	7.23
Histidine	1.88
Isoleucine	4.60
Leucine	8.42
Lysine	4.80
Methionine	1.87
Phenylalanine	4.30
Threonine	3.72
Valine	4.02
Nonessential amino acids (NEAAs)
Alanine	6.41
Aspartic acid	9.33
Cysteine	0.23
Glutamic acid	13.03
Glycine	5.22
Proline	5.05
Serine	2.85
Tyrosine	3.85
ΣEAAs	40.84
ΣNEAAs	45.97
ΣAAs	86.81
ΣEAAs/ΣNEAAs	0.89

Table 4. Proximate composition of oyster meat and sea bream fillet (%).

Proximate composition	July		September		November		March		Sea bream fillet
Proximate composition	Distant	Near	Distant	Near	Distant	Near	Distant	Near	Fish farm
Dry matter	14.28 ± 0.12^aAB	13.32 ± 0.09^bX	13.83 ± 0.14^aB	9.11 ± 0.05^bZ	9.78 ± 0.05^C	9.33 ± 0.03^Z	14.63 ± 0.05^aA	12.50 ± 0.04^bY	30.44 ± 0.26
Crude protein	7.19 ± 0.32^aB	5.71 ± 0.13^bY	5.53 ± 0.24^aB	4.49 ± 0.21^bZ	5.48 ± 0.01^B	5.37 ± 0.07^Y	7.81 ± 0.21^aA	6.77 ± 0.12^bX	20.91 ± 0.15
Crude lipid	1.31 ± 0.05^aA	1.19 ± 0.02^bX	0.96 ± 0.03^aB	0.70 ± 0.02^bY	0.45 ± 0.01^bC	0.75 ± 0.01^aY	0.65 ± 0.01^aC	0.42 ± 0.02^bZ	7.97 ± 0.02
Ash	1.17 ± 0.03^B	1.27 ± 0.09^Z	1.26 ± 0.07^bB	1.48 ± 0.13^aZ	1.22 ± 0.02^bB	1.71 ± 0.02^aY	2.73 ± 0.04^A	2.59 ± 0.04^X	1.48 ± 0.01

Table 5. Fatty acid composition of oysters and sea bream fillet (g/100 g fatty acids).

Fatty acids	July		September		November		March		Sea bream fillet
Fatty acids	Distant	Near	Distant	Near	Distant	Near	Distant	Near	Fish farm
14:0	5.41 ± 0.12^b	5.58 ± 0.16^aY	5.64 ± 0.22	5.94 ± 0.12^Y	5.77 ± 0.42	5.37 ± 0.29^Z	5.95 ± 0.31^b	6.55 ± 0.38^aX	1.99 ± 0.05
15:0	0.87 ± 0.04^bC	1.20 ± 0.08^aY	1.01 ± 0.09^bB	1.22 ± 0.03^aY	1.24 ± 0.07^bA	1.62 ± 0.09^aX	1.33 ± 0.11^a	1.43 ± 0.06^X	0.24 ± 0.03
16:0	25.56 ± 0.86^b	25.29 ± 0.71^Y	25.49 ± 0.65^b	24.63 ± 0.37^Z	26.60 ± 1.57^a	26.42 ± 1.14^X	26.38 ± 1.34^aA	25.11 ± 1.41^bY	15.22 ± 0.59
17:0	1.82 ± 0.07^bB	1.81 ± 0.12^aY	1.78 ± 0.04^bB	1.95 ± 0.15^aY	2.24 ± 0.16^a	2.19 ± 0.16^X	2.42 ± 0.17^aA	1.98 ± 0.15^bY	0.23 ± 0.01
18:0	8.32 ± 0.29^a	8.30 ± 0.26^Y	7.78 ± 0.15^b	7.44 ± 0.34^Z	7.44 ± 0.97^b	7.52 ± 0.82^Z	7.69 ± 0.36^bB	9.26 ± 0.64^aX	3.48 ± 0.17
20:0	ND	ND	0.28 ± 0.02^b	0.25 ± 0.01^Z	0.58 ± 0.04^a	0.55 ± 0.04^X	0.27 ± 0.02^bB	0.39 ± 0.03^aY	0.31 ± 0.00
16:1n‐7	4.50 ± 0.09^c	5.10 ± 0.13^Y	5.99 ± 0.14^aA	5.49 ± 0.23^bX	5.19 ± 0.46^aB	4.53 ± 0.28^bZ	5.33 ± 0.27^aB	4.95 ± 0.41^bY	1.13 ± 0.03
18:1n‐9	7.34 ± 0.51^aA	6.77 ± 0.22^b	4.27 ± 0.08^bC	6.67 ± 0.36^a	5.69 ± 0.55^bB	6.73 ± 0.49^a	5.89 ± 0.32^bB	6.73 ± 0.37^a	4.23 ± 0.14
20:1n‐9	0.73 ± 0.03^bB	0.94 ± 0.03^aY	0.56 ± 0.02^bC	0.82 ± 0.04^aZ	1.17 ± 0.09^bA	1.30 ± 0.12^aX	0.64 ± 0.04^bB	0.99 ± 0.05^aY	30.91 ± 1.15
22:1n‐11	ND	ND	ND	ND	0.12 ± 0.02^a	0.10 ± 0.01^X	0.06 ± 0.01^b	0.07 ± 0.01^Y	1.48 ± 0.07
24:1n‐9	ND	ND	ND	ND	0.18 ± 0.03^a	0.21 ± 0.02^X	0.10 ± 0.01^b	0.13 ± 0.01^Y	0.37 ± 0.02
18:2n‐6	4.73 ± 0.32^aA	3.95 ± 0.09 b^X	2.92 ± 0.11^c	3.12 ± 0.14^Y	3.25 ± 0.12^aB	2.84 ± 0.22^bZ	2.99 ± 0.14^c	3.14 ± 0.17^Y	21.96 ± 0.52
20:2n‐6	ND	ND	0.31 ± 0.02^b	0.32 ± 0.03^Y	0.39 ± 0.06^bA	0.48 ± 0.05^aX	0.39 ± 0.03^bA	0.51 ± 0.04^aX	0.66 ± 0.02
18:3n‐3	1.58 ± 0.06^aC	1.43 ± 0.05^bZ	1.38 ± 0.06^bD	1.53 ± 0.09^aZ	1.73 ± 0.11^b	1.71 ± 0.15^Y	2.11 ± 0.11^aA	1.98 ± 0.17^bX	3.51 ± 0.10
20:3n‐3	ND	ND	0.40 ± 0.03^aA	0.22 ± 0.01^bY	0.23 ± 0.01^bB	0.37 ± 0.02^aX	0.39 ± 0.04^a	0.35 ± 0.03^X	0.19 ± 0.01
20:4n‐6	3.25 ± 0.14^bC	4.00 ± 0.21^aY	5.16 ± 0.41^aA	4.49 ± 0.23^bX	4.47 ± 0.36^b	4.20 ± 0.27^X	2.87 ± 0.16^d	2.92 ± 0.21^Z	0.45 ± 0.02
20:5n‐3	15.25 ± 0.74^aA	14.54 ± 0.62^bX	15.09 ± 0.39^aA	14.01 ± 0.53^bY	13.44 ± 0.66^c	13.50 ± 0.62^Z	14.58 ± 0.86^aB	13.96 ± 0.75^bY	4.06 ± 0.06
22:6n‐3	20.63 ± 0.89^a	21.09 ± 0.44^X	20.40 ± 0.51^a	20.84 ± 0.71^X	18.46 ± 1.10^b	18.77 ± 1.17^Y	18.74 ± 1.14^aB	17.82 ± 1.53^bZ	9.21 ± 0.09
ΣSFAs^a	41.98 ± 0.78^bB	42.18 ± 0.96^aY	41.98 ± 0.84^b	41.43 ± 0.69^Y	43.87 ± 0.54^a	43.67 ± 0.63^X	44.04 ± 0.72^a	44.72 ± 0.76^X	21.47 ± 0.17
ΣMUFAs^b	12.57 ± 0.36^aA	12.81 ± 0.33^bX	10.82 ± 0.27^bB	12.98 ± 0.38^aX	12.35 ± 0.44^a	12.87 ± 0.35^X	12.02 ± 0.27^a	12.87 ± 0.41^X	38.12 ± 0.34
ΣPUFAs^c	45.44 ± 0.93^aA	45.01 ± 0.88^bX	45.66 ± 0.79^aA	44.53 ± 0.88^bX	41.97 ± 0.67^b	41.87 ± 0.73^Y	42.07 ± 0.76^aB	40.68 ± 0.65^bZ	40.04 ± 0.31
Σn‐3 HUFAs^d	35.88 ± 0.32^a	35.63 ± 0.42^X	35.89 ± 0.35^a	35.07 ± 0.4^6X	32.13 ± 0.38^c	32.64 ± 0.48^Y	33.71 ± 0.41^aB	32.13 ± 0.29^bY	16.97 ± 0.19
Σn‐3	37.46 ± 0.42^a	37.06 ± 0.37^X	37.27 ± 0.32^a	36.60 ± 0.41^X	33.86 ± 0.28^c	34.35 ± 0.41^Y	35.82 ± 0.33^b	34.11 ± 0.39^Y	13.46 ± 0.15
Σn‐6	7.98 ± 0.13^b	7.95 ± 0.11^X	8.39 ± 0.09^a	7.93 ± 0.11^X	8.11 ± 0.07^a	7.52 ± 0.16^X	6.25 ± 0.10^c	6.57 ± 0.09^Z	23.07 ± 0.30
n‐3/n‐6	4.69 ± 0.05^b	4.66 ± 0.11^Y	4.44 ± 0.07^b	4.62 ± 0.06^Y	4.18 ± 0.06^c	4.57 ± 0.08^Y	5.73 ± 0.09^a	5.19 ± 0.06^X	0.74 ± 0.02
DHA/EPA	1.35 ± 0.03	1.45 ± 0.05^X	1.35 ± 0.05	1.49 ± 0.06^X	1.37 ± 0.03	1.39 ± 0.04^X	1.30 ± 0.04	1.28 ± 0.03^Y	2.27 ± 0.03

Table 6. Amino acid composition of oysters and sea bream fillet (g/100 g crude protein).

Amino acids	July		September		November		March		Sea bream fillet
Amino acids	Distant	Near	Distant	Near	Distant	Near	Distant	Near	Fish farm
EAAs
Arginine	8.60 ± 0.07^bB	8.91 ± 0.07^aX	8.39 ± 0.06^aB	8.07 ± 0.06^bY	9.11 ± 0.11^A	8.91 ± 0.06^X	8.96 ± 0.08^A	8.82 ± 0.07^X	6.80 ± 0.45
Histidine	3.36 ± 0.03^aA	3.10 ± 0.03^bX	1.97 ± 0.02^bB	2.99 ± 0.03^aX	1.67 ± 0.03^bC	1.78 ± 0.02^aZ	2.02 ± 0.03^aB	1.92 ± 0.03^bY	3.02 ± 0.19
Isoleucine	4.95 ± 0.04^bC	5.36 ± 0.04^aY	5.37 ± 0.04^B	5.24 ± 0.03^Y	5.77 ± 0.04^bA	5.94 ± 0.05^aX	5.50 ± 0.03^bA	5.71 ± 0.06^aX	6.12 ± 0.32
Leucine	8.83 ± 0.07^A	8.72 ± 0.07^X	8.58 ± 0.07^B	8.41 ± 0.06^XY	8.62 ± 0.09^B	8.61 ± 0.06^X	8.12 ± 0.09^C	8.25 ± 0.08^Y	7.97 ± 0.41
Lysine	9.12 ± 0.07^bC	9.82 ± 0.06^aY	9.76 ± 0.07^aB	9.27 ± 0.08^bZ	10.88 ± 0.13^A	10.71 ± 0.11^X	10.63 ± 0.15^A	10.66 ± 0.06^X	8.12 ± 0.25
Methionine	2.16 ± 0.02^B	2.21 ± 0.01^Y	2.21 ± 0.01^B	2.18 ± 0.00^Y	3.68 ± 0.02^A	3.66 ± 0.05^X	3.86 ± 0.04^A	3.87 ± 0.03^X	3.10 ± 0.11
Phenylalanine	4.86 ± 0.04^aC	4.54 ± 0.04^bY	4.14 ± 0.03^bD	4.33 ± 0.03^aY	5.52 ± 0.05^B	5.67 ± 0.07^X	6.10 ± 0.04^A	5.96 ± 0.06^X	4.41 ± 0.22
Threonine	4.43 ± 0.03^A	4.57 ± 0.03^X	4.70 ± 0.04^A	4.58 ± 0.04^X	3.81 ± 0.03^C	3.91 ± 0.03^Y	4.14 ± 0.05^B	4.03 ± 0.05^Y	5.01 ± 0.29
Valine	4.68 ± 0.04^aA	4.22 ± 0.03^bY	4.74 ± 0.04^bA	4.97 ± 0.04^aX	3.97 ± 0.05^aB	3.78 ± 0.05^bZ	3.91 ± 0.03^B	3.99 ± 0.05^YZ	4.94 ± 0.28
NEAA
Alanine	6.26 ± 0.06^A	6.14 ± 0.05^X	6.38 ± 0.05^aA	6.01 ± 0.06^bX	5.16 ± 0.04^B	5.13 ± 0.06^Y	4.79 ± 0.04^C	4.79 ± 0.03^Y	7.65 ± 0.47
Aspartic acid	13.12 ± 0.12^A	13.27 ± 0.11^X	13.28 ± 0.12^aA	12.36 ± 0.16^bY	9.15 ± 0.05^B	9.25 ± 0.11^Z	9.25 ± 0.10^B	9.05 ± 0.13^Z	10.39 ± 0.65
Cysteine	0.48 ± 0.01^A	0.42 ± 0.02^Y	0.49 ± 0.03^aA	0.36 ± 0.01^bZ	0.36 ± 0.01^B	0.40 ± 0.01^Y	0.50 ± 0.01^A	0.47 ± 0.02^X	0.27 ± 0.01
Glutamic acid	11.06 ± 0.14^aA	10.19 ± 0.10^bY	11.32 ± 0.13^bA	12.02 ± 0.09^aX	10.48 ± 0.11^B	10.42 ± 0.13^Y	9.68 ± 0.08^C	9.76 ± 0.08^Z	12.02 ± 0.97
Glycine	8.22 ± 0.06	8.06 ± 0.05^Y	8.11 ± 0.07	7.93 ± 0.06^Z	7.91 ± 0.08	7.74 ± 0.08^Z	7.96 ± 0.09^b	8.56 ± 0.09^aX	7.76 ± 0.48
Proline	1.21 ± 0.05^C	1.25 ± 0.05^Y	1.71 ± 0.06^aA	1.07 ± 0.06^bZ	1.50 ± 0.02^B	1.42 ± 0.03^X	1.55 ± 0.02^B	1.48 ± 0.03^X	4.50 ± 0.22
Serine	5.27 ± 0.04 b^B	5.77 ± 0.04^aY	5.16 ± 0.04^bB	5.71 ± 0.04^aY	6.23 ± 0.06^bA	6.42 ± 0.07^aX	6.63 ± 0.06^aA	6.17 ± 0.06^bX	4.13 ± 0.26
Tyrosine	3.28 ± 0.02^aB	3.11 ± 0.02^bZ	3.16 ± 0.02^bB	3.91 ± 0.03^aY	5.64 ± 0.07^A	5.75 ± 0.07^X	5.83 ± 0.05^A	5.87 ± 0.05^X	4.14 ± 0.21
ΣEAA	50.99 ± 0.21^B	51.45 ± 0.25^Y	49.86 ± 0.29^C	50.03 ± 0.23^Z	53.03 ± 0.38^A	52.97 ± 0.33^X	53.24 ± 0.32^A	53.21 ± 0.33^X	49.49 ± 0.42
ΣNEAA	48.90 ± 0.37^B	48.21 ± 0.41^X	49.61 ± 0.39^A	49.36 ± 0.40^X	46.43 ± 0.22^C	46.53 ± 0.28^Y	46.19 ± 0.28^C	46.15 ± 0.21^Y	49.86 ± 0.27
ΣAA	99.89 ± 0.63	99.66 ± 0.56	99.47 ± 0.58	99.39 ± 0.66	98.46 ± 0.51	99.50 ± 0.47	99.43 ± 0.51	99.36 ± 0.54	99.35 ± 0.58
ΣEAA/ΣNEAA	1.04 ± 0.01^B	1.06 ± 0.03^Y	1.01 ± 0.03^BY	1.01 ± 0.02	1.14 ± 0.05^A	1.14 ± 0.05^X	1.15 ± 0.03^A	1.15 ± 0.03^X	0.99 ± 0.03

Funding3

—Project COREFISH CUP P2022ZPMXH_002
—PO FEAMP
—Scientific Research Projects Coordination Unit of Istanbul University

Keywords

feedingIMTAnutritional profileOstrea edulisreproductionSparus aurata

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Taxonomy

TopicsMarine Bivalve and Aquaculture Studies · Aquaculture Nutrition and Growth · Meat and Animal Product Quality

Full text

1. Introduction

Mollusk aquaculture has been growing exponentially over the last decades, from 12 million tonnes in the 1990s to 19 million tonnes in 2022, a 58% growth over 22 years [1]. Production has grown by 1 million tonnes between 2020 and 2022 alone, increasing 15.6% based on soaring global demand for sustainable seafood and increasing product diversification [2–4]. In Europe, finfish are the leading aquaculture, but the second largest production group comprises bivalves such as mussels, oysters, and clams, with 553,000 tonnes [2, 5].

Despite their benefits, intensive finfish aquaculture systems have produced a range of environmental issues like nutrient enrichment, habitat alteration, and loss of biodiversity [6–8]. Integrated Multi‐Trophic Aquaculture (IMTA) has been among the most promising sustainable methods for reducing such impacts by optimizing recycling of nutrients and maintaining ecosystem balance [9, 10]. IMTA integrates species from different trophic levels, typically combining finfish with bivalves and macroalgae, where organic waste from fish farming is utilized as nutrients for filter‐feeders, mimicking natural ecosystem processes [11–13].

The European flat oyster (Ostrea edulis) is particularly valuable both ecologically and economically among the bivalves suited for IMTA. Historically an important species in coastal ecosystems and a highly prized seafood in Europe, it is currently produced mainly in France, Ireland, Spain, and the Netherlands [1, 14, 15]. O. edulis efficiently filters large volumes of water, reducing phytoplankton biomass and improving water quality, making it ideal for IMTA integration [16–18]. Moreover, this species is a rich source of essential nutrients, including proteins, amino acids, and omega‐3 fatty acids, which are critical for human health [19, 20]. Amino acids like arginine and lysine support muscle repair and immune function, while omega‐3 fatty acids such as EPA and DHA play important roles in cardiovascular health and cognitive function [21–24]. These nutrients not only support basic metabolic functions but also contribute to preventing chronic diseases, highlighting the importance of oysters as nutrient‐rich functional foods [25, 26]. In addition, research on the nutritional quality of O. edulis within IMTA remains limited, representing a critical knowledge gap.

The effectiveness of IMTA systems depends on how species at various trophic levels are arranged and interact within the space [27]. Specifically, the proximity of oysters to fish farms influences their nutritional composition and growth, which in turn affects their market value. Fish farms release organic wastes such as uneaten feed and fish excretions, which can enhance nutrient availability for filter‐feeders like O. edulis [28, 29]. However, the benefits of such nutrient enrichment depend on factors including distance from the farm, local hydrodynamics, and environmental conditions [30]. Therefore, understanding how proximity to fish farms affects oyster growth and nutrition is vital for optimizing IMTA design and management.

Seasonal changes also play a critical role in oyster meat quality by influencing environmental parameters such as water temperature, salinity, and food availability [31, 32]. Seasonal changes in primary productivity, reproductive activity, and metabolic requirements drive physiological and biochemical variations in oysters [33, 34]. For example, cold months are typically associated with higher glycogen reserves and better meat condition due to reduced spawning, whereas warmer months stimulate lipid metabolism and protein turnover linked to reproduction [35, 36].

To our knowledge, there are only a few studies on European flat oysters in IMTA systems evaluating growth, such as those by Aguado‐Giménez et al. [37] and Nikolić et al. [38]; however, this study is the first to examine meat quality within an IMTA system. The objective of this study is to evaluate the impact of proximity to a fish farm on the meat quality of European flat oysters cultivated in an IMTA system with Sparus aurata in the Tyrrhenian Sea. By comparing oysters grown at two specific distances (20 m and 800 m) from fish cages over a 257‐day period, the study aims to determine how fish farm‐derived nutrient availability influences the nutritional profile of the European flat oyster. This profile includes proximate composition, amino acid, and fatty acid content. The 20 m and 800 m distances were strategically selected based on previous literature [39] and regional hydrodynamic knowledge to cover the critical range where nutrient effects are most pronounced. The 20 m site allows for the evaluation of maximum exposure to direct fish waste, while the 800 m site is intended to capture the peak development of secondary phytoplankton blooms that occur after the initial dilution and assimilation of the cage effluent. This range allows us to identify the optimal spatial placement for oyster integration within the IMTA system, maximizing nutrient assimilation and overall system performance. The ultimate aim of the study is to not only assess the physiological response but, more critically, to quantify the market‐relevant nutritional quality (proximate composition, amino acids, and fatty acids) of O. edulis cultivated under IMTA conditions, thereby linking ecological engineering to functional food production.

2. Material and Methods

2.1. Experimental Design and Diets

This study was conducted over a 257‐day period, from July 8, 2023, to March 21, 2024, in the coastal waters of Gaeta, Italy. The experimental sites were located near the La Marea mussel farm (Mytilus galloprovincialis) (41°14′16″N, 13°35′55″E) and the P2G Sparus aurata fish farm (41°13′44″N, 13°35′58″E) (Figure 1). European flat oysters (Ostrea edulis) were deployed at two distinct locations relative to the fish farm: one at ~20 m downstream of the general water circulation (referred to as the “Near” site), and the other at ~800 m downstream (the “Distant” site). Existing infrastructure from the mussel farm was utilized to support the oyster placements. Commercial feed was used at the fish farm (Tables 1–3).

Study site in the Gulf of Gaeta.

A total of 2500 healthy oysters were prepared as an experimental group. At the beginning of the experiment, the average weight of the adult oysters (33.65 ± 2.30 g), shell length (7.00 ± 0.26 cm), and shell area (23.68 ± 1.57 cm^2^) were measured. A total of eight lantern nets (four in the Near and four in the Distant site, mesh size: 3 cm, diameter: 60 cm) of ten levels each placed at a depth of 7 m were used in the experiment. Every 2 months, oysters were collected from a single level of lantern nets, with each level spaced 25 cm apart, and transported in cold boxes to the Laboratory of Experimental Ecology and Aquaculture of the University of Rome Tor Vergata. Also, sea bream samples were bought from the P2G farm in March at the end of the experiment at commercial size (404 ± 18.20 g) and transferred to the same laboratory to determine the final commercial quality of the finfish. The sampling months of July, September, November, and March were selected to represent key seasonal periods within the 9‐month study duration. These months correspond to the following seasons in the study area: summer, autumn, winter, and spring, respectively. Samples were arranged in the laboratory, and pools were prepared with at least 5 oysters in each group. Dead oysters were not included in the study. The meat and shells of each oyster were separated, and the average meat and shell weight per group was recorded. In the meantime, fish fillet was prepared by removing the skin and bones and then portioned into 70 g appropriate sample sizes. The meat was dried in a freeze dryer (Telstar LyoQuest 62302), and the shells were dried in the oven, and their dry weights were measured on a precision balance (Orma Italy, BC 202). Then, the proximate analyses described below were conducted at the Fish Nutrition Laboratory, Faculty of Aquatic Sciences, Istanbul University. Additionally, amino acid and fatty acid analyses of the same samples were carried out at a separate accredited laboratory.

The proximate (Table 1), fatty acid (Table 2) and amino acid (Table 3) compositions of the commercial feed used in sea bream farms are given below.

2.2. Water Quality

Temperature, dissolved oxygen, and pH of the water column in the experimental area were measured with the Hach 40QD model multiparameter every 2 weeks. Salinity was measured with the YSI 3200 conductivity instrument, and chlorophyl‐a was analyzed spectrophotometrically following acetone extraction, according to the SM 10200 H method from the collected water samples.

2.3. Proximate Composition

Commercial feed, fish fillet, and oyster (samples were taken by separating meat from the shell) samples were analyzed for proximate composition (crude protein, crude lipid, ash, moisture for all samples, and fiber just for the feed) in the laboratory as three replications according to AOAC [40] at the end of the months of July 2023, September 2023, November 2023, and March 2024. Crude lipids were extracted by the Soxhlet method with diethyl ether (Velp‐Scientifica Ser, 148). Crude protein (N × 6.25) was determined using a semi‐automatic Kjeldahl (Gerhardt Vapodest, 45 s) technique. Moisture content was measured by drying samples in an oven (Nüve FN 400, TS 6073) at 105°C for 12 h. Dried samples were incinerated at 550°C for about 12 h in a muffle furnace for the determination of ash content after weight loss. Using sulfuric acid and then sodium hydroxide, 12.5% (w/w) for half an hour each, with the final residue washed with 5% HCl and water, filtered, dried, and weighed, the crude fiber was determined.

2.4. Fatty Acid Analysis

Lipid extraction from oyster, fish feed, and fish fillet samples was performed using the chloroform/methanol (2:1, v/v) method described by Folch et al. [41]. Fatty acid methyl esters (FAMEs) were prepared through transmethylation with 2 M potassium hydroxide (KOH) in methanol and n‐hexane, following Ichihara et al. [42] with minor modifications. Specifically, 10 mg of extracted oil was dissolved in 2 mL of hexane, followed by the addition of 4 mL of 2 M methanolic KOH. The mixture was vortexed for 2 min at room temperature and centrifuged at 4000 rpm for 10 min, after which the upper hexane layer was collected for gas chromatography (GC) analysis. Fatty acid composition was determined using a Shimadzu GC‐FID 2010 gas chromatograph equipped with a 60 × 0.25 mm Agilent DB‐23 capillary column. The analytical conditions were as follows: helium as the carrier gas, flame ionization detection (FID) at 250°C, injector temperature at 250°C, a split ratio of 1:0, and an oven temperature program increasing from 120°C (2 min) to 240°C (7 min) at a rate of 5°C min^−1^. FAMEs were identified by comparison with commercial standards (Sigma–Aldrich, Steinheim, Germany). All FA analyses were conducted in triplicate.

2.5. Amino Acid Analysis

Amino acid analysis for oyster, fish feed, and fish fillet samples was conducted using a high‐sensitivity triple quadrupole mass spectrometer (LCMS/8050, SHIMADZU, <0.7 u resolution, m/z 2–2000 mass range, up to 30,000  u/s scan speed, up to 555 channels/s MRM transition rate, 5 ms polarity switching time) following AOAC [40] guidelines. Samples were treated with petroleum ether for oil extraction, followed by centrifugation. Hydrolysis was performed with 6N HCl in Schott bottles, heated at 110°C for 24 h. After hydrolysis, samples were filtered and diluted 1500 times before analysis on the LCMS‐MS device. All analyses were performed in triplicate.

2.6. Data Analysis

All results are presented as means ± standard deviation (SD). Statistical analyses were conducted using SPSS version 28 statistical software. For comparing, distinguishing, and discriminating the changes in quality properties between the experimental groups, Principal Component Analysis (PCA) and Hierarchical Cluster Analysis are done. After, data were analyzed using multivariate analysis of variance (MANOVA) to assess differences across groups from two different sizes and four different seasons. When significant effects were detected, Tukey’s post hoc test was applied for pairwise comparisons. Statistical significance was set at p < 0.05.

3. Results

At the end of the experiment, oysters at the Near site reached 49.41 ± 9.76 g and those at the Distant site 52.65 ± 12.18 g [43].

3.1. Water Quality

During the experiment, water temperature fluctuated between 16.6 ± 0.65°C and 25.8 ± 0.96°C (Figure 2). Dissolved oxygen levels varied from 7.34 ± 0.95 mg/L to 9.24 ± 1.89 mg/L, peaking in November. Chlorophyl‐a concentrations ranged from 0.53 ± 0.01 µg/L to 3.23 ± 0.84 µg/L, with lower values observed during the summer months.

Water temperature, dissolved oxygen, chlorophyl‐a, pH, and salinity values in the experimental area.

3.2. Proximate Composition of Oyster and Sea Bream

The crude protein ratio of oysters varied between 4.49% in the Near September group and 7.81% in the Distant March group, with the Distant March, Near March, and Distant July groups having the highest protein content (p > 0.05). Crude lipid content was found to be lowest in the Near March group with 0.42% and highest in the Distant July group with 1.31% (Table 4). Crude lipid ratio was found to be similar in the Distant July and Near July groups and higher than other groups (p < 0.05). When dry matter ratios were examined, they varied between 9.11% in the Near September group and 14.63% in the Distant March group (p < 0.05). Ash ratios were determined as a minimum of 1.17% in the Distant July group and a maximum of 2.73% in the Distant March group. Ash ratios of the Distant March and Near March groups were statistically similar to each other and higher than other groups (p < 0.05). The Distant March had the highest values in terms of dry matter, crude protein, and ash. Crude protein, crude lipid, and ash contents of sea bream fillet were found to be 20.91%, 7.97%, and 1.48%, respectively (Table 4).

3.3. Fatty Acid Composition of Oysters and Sea Bream

In all oyster groups, the total saturated fatty acid (ΣSFA) ratios were found to be significantly higher in November and March compared to other months (p < 0.05; Table 5). Among the SFAs, palmitic acid (16:0) was the most abundant (p < 0.05). For monounsaturated fatty acids (MUFAs), the ratios ranged from 10.82% in the Distant September group to 12.98% in the Near September group (p < 0.05), with oleic acid (18:1n‐9) being the predominant MUFA (p < 0.05). When comparing total n‐6 fatty acid ratios, the highest levels were observed in the Distant September group, while the Distant March and Near March groups showed the lowest levels (p < 0.05). The main contributors to the n‐6 fatty acid profile were linoleic acid (LA, 18:2n‐6) and arachidonic acid (ARA, 20:4n‐6). Regarding total n‐3 fatty acid ratios, these were significantly lower in November and March compared to July and September (p < 0.05). The primary n‐3 fatty acids were EPA and DHA, with the DHA/EPA ratios being higher in the Near groups during July and September. In sea bream, the EPA content was determined to be 4.06%, while DHA was measured at 9.21%, resulting in a DHA/EPA ratio of 2.27.

3.4. Amino Acid Composition of Oysters and Sea Bream

Arginine and lysine were identified as the most abundant essential amino acids (EAAs) in all oyster experimental groups. Arginine levels were significantly higher in November and March, particularly in Distant sites (p < 0.05; Table 6). Phenylalanine exhibited a similar pattern. Conversely, leucine, threonine, and valine showed an opposite trend, with their levels decreasing during these months. Histidine levels began to decline after July. Significant differences between groups were observed for histidine, isoleucine, leucine, and valine, favoring the Distant site group (p < 0.05). For the other EAAs, no significant differences were detected between the Distant and Near groups, though seasonal fluctuations were noted. Overall, the total essential amino acids (ΣEAAs) reached their highest levels in March, with values of 53.24% and 53.21% in both sites. Meanwhile, total nonessential amino acids (ΣNEAAs) were highest in July and September for the Distant and Near sites.

In sea bream fillet, leucine, lysine, and arginine were the most abundant EAAs, while histidine had the lowest concentration. Among the nonessential amino acids (NEAAs), glutamic acid had the highest value. The ΣEAAs were calculated as 49.49%, while the total ΣNEAAs were slightly higher at 49.86%.

4. Discussion

This study investigated the proximate composition of oysters cultivated downstream at the Near (20 m) and Distant (800 m) conditions from a fish farm in the IMTA system. Additionally, nutritional analyses were performed on sea bream fillets from the fish farm, and water quality measures have been recorded.

Water quality in the study area showed temperatures ranging from 14°C in winter to 26°C in summer and salinity between 36 and 39 PSU, reflecting the Mediterranean’s limited water circulation and high evaporation rates (Figure 2, [44, 45]). Chlorophyl‐a concentrations, used as an indicator of phytoplankton abundance, varied seasonally from 0.53 ± 0.01 to 3.23 ± 0.84 µg/L, driven by nutrient availability, light conditions, and water mixing [46, 47]. These parameters provide a baseline overview of the environmental conditions influencing oyster growth in the region, which are near the reported optimum for oyster cultivation (15°C–20°C and 25–35 PSU; [48]).

The results revealed significant differences in the nutrient profiles of the oysters, with crude protein content ranging from 4.49% to 7.81% throughout the 6‐month experimental period. Notably, the highest protein content was observed in oysters collected from the Distant site, particularly during March and July. Seasonal variations were evident, as the nutrient differences between oysters from the Near and Distant sites were minimal due to lower metabolic activity in November and March but more pronounced in other months. These variations suggest that environmental factors, such as water quality and nutrient availability, along with differences in the nutrient sources available to oysters, play a critical role in shaping their nutritional composition [49, 50]. Crude lipid content among oysters was lowest at 0.42% in the Near site in March, while it reached the highest level at 1.31% in the Distant site in July. The subsequent elevation of lipid content in July (during the spawning window) reflects continued gonadal maturation and is crucial for the final energetic provisioning of eggs and sperm, as well as the initial metabolic recovery postspawning. Moreover, the dry matter ratio was lowest at 9.11% in September oysters in the Near site and reached its maximum at 14.63% in March oysters in the Distant site. Variability in dry matter content indicates differences in the water content of oyster samples. Higher dry matter levels suggest a greater concentration of nutrients other than water within the sample [51]. The significant peak in crude protein and dry matter observed in March signifies the biological allocation of accumulated reserves toward the demanding requirements of gametogenesis, thereby confirming the IMTA environment successfully fuels reproductive maturation while supporting prior growth findings (Batır et al., submitted for publication) and, besides increasing food availability in the water column, from a commercial perspective, translates to a superior market product with enhanced essential protein content, directly addressing consumer demand for high‐value marine protein sources. On the other hand, ash ratios associated with the presence of minerals in oysters [52] were found to be between 1.17% in the Distant July and 2.73% in the Distant March groups. As a result, oysters from the Distant site location exhibited the highest values for dry matter, protein, and ash, indicating a generally superior biochemical composition in March. However, their lipid content was slightly lower compared to the other groups. It may be attributed to the optimized growth conditions in open‐sea IMTA systems when oysters are placed ~800 m from fish farms. At this distance, oysters benefit from phytoplankton blooms stimulated by fish farm waste rather than directly ingesting the waste itself, maximizing their nutrient intake. In contrast, oysters positioned too close (20 m) experience direct exposure to waste, which can limit nutrient absorption and potentially clog their gills due to organic particle accumulation [53, 54].

The 800‐meter distance thus represents an optimal balance enhancing nutrient availability while minimizing the adverse effects of waste [55, 56]. Additionally, the observed differences in nutrient composition may be influenced by environmental factors at the growth site, a hypothesis supported by previous studies [57, 58]. Seasonal variations, including temperature fluctuations, reproductive cycles, and the continued growth of the oysters throughout the experiment, may have also played a role [59, 60].

Oysters have lower levels of nutrients in September and November when compared to March in this study. It is thought to be due to the reproduction season of oysters being between May and September [61–66], and oysters show a decrease in growth and nutrients during this period because they spend their energy on reproduction rather than storing it. Different studies have also provided information that supports this idea [67, 68]. In another study conducted in Italy, the protein content of oysters was determined as 8.10%, lipid content as 2.70%, and ash content as 3.72% [69]. Van Houcke et al. [70] calculated the nutrients of oysters based on dry weight in their study, but when the moisture content in the body was taken into account, they reported the proximate composition as 12.09% protein, 1.28% lipid, and 2.43% ash. Similarly, Pogoda et al. [71] stated in their research that they found protein to be 37.3% DW and lipid to be 9.15% DW in the summer months, and protein to be 35% and lipid to be 7.7% in the autumn months on a dry weight basis. The results of the above study regarding nutrients are consistent with the results of this study when evaluated considering moisture. These results suggest that the nutritional content of oysters may vary depending on their size and environmental conditions. The wide variation observed between protein, lipid, and ash ratios in oysters may be related to the life cycle of oysters, their feeding habits, and the environmental factors they are exposed to [72]. The determined seasonal shifts in proximate composition, particularly the high crude protein and dry matter in March (prespawning) and the elevated lipids in July (postspawning recovery), are fundamentally driven by the food availability, the oyster’s reproductive cycle, and it’s energy partitioning strategy. As detailed in the companion growth study (Batır et al., submitted for publication), the period leading up to March represented a phase of high somatic growth and tissue accumulation, followed by the onset of spawning activity in spring.

In the present study, protein, lipid, dry matter, and ash ratios were found to be 20.91%, 7.97%, 30.44%, and 1.48% in sea bream fillet, respectively. Since oysters are cultivated in proximity to the fish farm to utilize organic waste, it is essential to ensure that the integrated approach does not adversely affect the nutritional quality of the fish. When comparing the results of this study with those from other studies, the reported ranges for protein, lipid, dry matter, and ash content were consistent, measuring 18%–21%, 5%–7%, 25%–30%, and 1.2%–2%, respectively [73–76]. There is an agreement between the fillet proximate composition of the fish in the present study and the corresponding results in the other studies mentioned above. Sea bream fillets are highly nutritious, making them an important component of a sustainable and healthy diet. Proximate composition of sea bream was analyzed in this study to evaluate the potential impacts of the IMTA system on fish health and product value. With their high protein content and rich lipid profile, sea breams represent a valuable option for human nutrition.

The study also investigated the effects of the IMTA system and seasonal effects on the fatty acid composition of oysters. Previous studies have also shown that bivalve mollusks consume fish farm waste when more suitable food sources are not available in IMTA systems [55, 77–79]. In this context, the IMTA system and seasonal variations significantly influence the fatty acid profile of oysters, particularly their n‐3 fatty acid content. Oysters in this study were associated with a tendency for n‐3 HUFAs to increase with higher temperatures. In November and March, when water temperatures were lower, n‐3 HUFA levels were at their lowest, whereas in July and September, during warmer temperatures, these levels were higher. Therefore, fatty acid levels tend to increase as temperature increases. This result supports studies suggesting that high levels of n‐3 HUFAs, especially EPA and DHA, are an adaptation to the temperatures encountered in the marine environment [80, 81].

Abad et al. [82] conducted a study focusing on the seasonal variation in lipid accumulation and fatty acid composition in O. edulis, and the effects of environmental factors such as temperature, food availability, and reproductive cycles. According to the results they reached, the fatty acid profile was positively correlated with temperature but not with chlorophyl‐a because oysters store glycogen instead of lipids during phytoplankton blooms. In the same study, lipid levels were reported to be lowest in November during early gametogenesis and to increase with nutrient availability, reaching a peak at maturity in May and June. However, it has been reported that lipid levels decrease after spawning, increase again with phytoplankton blooms during the gonadal resting phase, and then decrease in oysters as lipids are used when nutrients are scarce in the autumn. It has been reported that EPA and DHA reach their highest levels in spring and early summer in connection with phytoplankton increase and the gametogenic cycle [82]. It has been reported that oysters need EPA and DHA to ensure healthy development, and research has shown that oysters contain 2.8% EPA and 13.6% DHA [82]. The authors concluded that seasonal lipid changes in oysters are more influenced by diet quality and nutrient availability than by temperature. However, temperature affects the nutrient availability in the water. While fatty acids such as EPA and DHA play a critical role in the gonads and larvae, EAAs in particular are primarily accumulated, reflecting the effect of food on reproduction and larval quality [83]. It is believed that in a warmer water environment, these fatty acids are more readily absorbed into the tissues of oysters, supporting their energy needs and overall metabolism.

In the present study, the level of n‐3 fatty acids in oysters from both sites was found to be quite high, varying between 33.86% and 37.46%. These ratios were generally consistent but were higher in the Distant site in March, toward the end of the experiment. The robust n‐3 PUFA profile (high EPA and DHA) is a direct indicator that the IMTA system successfully supports the production of an oyster that contributes significantly to the recommended human intake of these cardioprotective fatty acids. However, the level of n‐6 fatty acids was much lower on average 7.53% in favor of the Distant site. The fatty acid ratios observed in oysters from this study were consistent with those reported by Prato et al. [84] for commercial‐sized oysters, which included ΣSFA 45.14%, ΣMUFA 18.52%, EPA 8.97%, DHA 10.96%, ΣPUFA 36.34%, ΣUNSFA 54.86%, and n‐3/n‐6 1.21. Also, the fatty acid profile of adult oysters in this study was generally higher compared to the other study of younger individuals [85]. In another study, while saturated fatty acids (SFAs) increase at low temperatures, it has been stated that the rate of oleic acid, which plays a role in energy storage and membrane flexibility, increases in summer months [86], aligning with the present results.

It is believed that the higher fatty acid levels observed in oysters from the Distant site can be attributed to the dispersion of waste from nearby fish farms, which likely fueled phytoplankton growth downstream from the fish plant area. In this study, the fatty acid composition in sea bream fillets reflected that in the feed. It was also found that the fatty acid profile of commercially sized fish fillets was consistent with the ratios reported in the following literature. The fatty acid profile of commercial sea bream has been reported as follows: ΣSFA 25%–30%, ΣMUFA 35%–40%, EPA 6%–8%, DHA 11%–13%, Σn‐3 18%–20%, Σn‐6 7%–9%, an n‐3/n‐6 ratio of 2–3, and a DHA/EPA ratio generally between 1.5 and 2 in different studies [75, 87–89].

The amino acid content of oysters changes throughout their life stages, and these changes are shaped by growth, reproduction, and metabolic needs [90, 91]. In this research, analysis of the EAA profile in oysters revealed that arginine, which is linked to growth, exhibited higher levels in November and March, whereas histidine displayed an inverse trend. Histidine is also important for protein synthesis and tissue repair [92], and it is thought to fluctuate in tissue levels during growth, potentially influenced by feeding strategies [93]. Leucine, which plays a role in muscle protein synthesis and whose value can be associated with the growth performance of organisms [94, 95], decreased throughout the experimental period. Besides, lysine levels increased in November and March, when water temperatures dropped, and it is thought that this situation may provide an advantage for protein synthesis [96]. In this research, no statistical difference was found between sites in terms of the total amount of EAAs. In general, the levels of EAAs in aquatic organisms are minimally influenced by the nutrients they consume, as suggested in the literature [97, 98]. This is because these organisms primarily store the proteins and amino acids acquired through the food chain in their bodies; once they establish internal balance, they selectively utilize EAAs to meet their metabolic needs [99]. Therefore, it is believed that there are no significant differences in EAA levels among the oysters in the present study.

Notably, aspartic acid, which plays an important role in energy metabolism [100], also decreased in proportion to decreasing water temperatures. This could be attributed to reduced energy expenditure and metabolic activity in response to decreasing temperatures. Although no significant differences were observed in amino acid levels between the Distant and Near sites overall, the Distant site exhibited higher productivity for cysteine in July and September and for proline in September. This phenomenon may be associated with water conditions and the stress responses of the organisms [101]. Tyrosine is an amino acid that plays an important role in stress management and hormone synthesis [102] and remained stable across sites and seasons throughout the experimental period and is considered to play a key role in understanding responses to environmental conditions in IMTA systems in future studies. In the present work, an examination of the amino acid findings in oysters revealed seasonal variations in both essential and NEAAs. Total EAA levels were highest in November and March. Overall, no significant seasonal or site‐related differences were observed in NEAAs.

Current findings suggest that the Near site may enhance the production of certain amino acids, with statistical differences observed particularly in valine. These results suggest that the Near site may offer advantages in terms of nutritional value in the long term. The increases in certain amino acids in the Near site, compared to the Distant site, provide insights into the potential contribution of both fish and oysters to mutual nutrient cycling. Frolov and Pankov [83] reported that the amino acid composition of O. edulis adult gonads and larvae were influenced by dietary intake. They found that EAAs such as aspartic acid, serine, alanine, cysteine, tyrosine, and proline, along with certain NEAAs like threonine and lysine, were predominantly accumulated in these tissues. Overall, EAAs accounted for 48%–50% of the total amino acids. In the same study, it was highlighted that lysine, a crucial amino acid for protein synthesis and muscle development, exceeds 8% in both the gonad and larval stages, underscoring its biological significance. Additionally, lysine and leucine were found at high levels in both the gonads and larvae, whereas methionine did not show significant variation. The study suggested that these differences in leucine, lysine, and methionine levels may be attributed to the varying nutritional requirements during the larval stage, while the stable presence of methionine could be linked to its essential role in energy metabolism and reproductive processes in the gonads [103, 104].

As is known, juvenile oysters are in a rapid growth phase; they require more active protein synthesis [105]. However, adult oysters, as observed in this research, are in more advanced stages of growth, resulting in a more balanced amino acid profile and stabilized protein synthesis. Consequently, the rates of NEAAs are relatively higher. Adult oysters prioritize energy storage and protective processes, leading to elevated levels of amino acids involved in energy metabolism, such as alanine, glycine, and serine [22, 106]. Adult oysters exhibit slowed growth alongside increased reproductive activity. Consequently, their amino acid profile shifts toward those that support reproductive performance. The importance of amino acids such as histidine, threonine, and leucine becomes more pronounced in the adult stage. Additionally, it has been noted that O. edulis contains high levels of glutamic acid, aspartic acid, and alanine in both juvenile and adult stages, highlighting their dual role in biological functions and flavor enhancement [107, 108].

When the amino acid profile of the sea bream fillet in this study is examined, it is seen that it has a rich content in terms of EAAs. EAA/NEAA was calculated as 0.97, and the result shows that the sea bream fillet offers a balanced amino acid profile. When the studies conducted on commercial sea bream fillets are examined, it is seen that the values found are 30%–40% for total EAA and 35%–45% for total NEAA [109–111], supporting the results of the present study.

5. Conclusion

This study investigated the influence of proximity to a gilthead seabream farm and seasonal variations on the proximate composition, fatty acid, and amino acid profiles of European flat oysters cultivated within an IMTA system in the Tyrrhenian Sea. Seasonal trends were evident, with water temperature peaking in July–August and chlorophyl‐a concentrations reaching their maximum in March. According to our study, European flat oysters in a Mediterranean IMTA system should ideally be harvested in March, during the prespawning period, when their protein, dry matter, and overall meat quality are at their peak. Oysters grown at the Distant site (800 m from the fish farm) generally exhibited higher protein and lipid contents compared to those at the Near site (20 m), except during November when lipid levels near the farm were unexpectedly elevated, possibly due to local seawater circulation patterns. Fatty acid analysis revealed lower SFA levels in July and September, suggesting increased metabolic utilization of SFAs at higher temperatures, while n‐3 PUFAs, including HUFAs, peaked during these warmer months, likely reflecting enhanced biosynthesis to support oyster growth and reproduction. EAA concentrations followed a similar pattern, with lower levels during peak growth periods but overall peaks aligning with nutrient storage prior to reproduction. The fatty acid composition of gilthead seabream fillets corresponded closely with their commercial feed, confirming the direct influence of diet on fish nutritional quality. These findings provide important insights into how seasonal dynamics and spatial positioning within an IMTA system affect oyster nutrition and metabolism. The results highlight the critical role of reproductive cycles in modulating nutrient profiles and underscore the advantage of cultivating oysters at intermediate distances (~800 m) from fish farms to optimize nutritional quality and ecosystem benefits. This spatial recommendation has direct implications for marine spatial planning and the designation of allocated zones for aquaculture supporting sustainable multi‐trophic farming strategies in the Tyrrhenian–Mediterranean region. Limitations of the study include the focus on only two spatial points and a single geographical area, which may limit broader generalization. Future research should explore additional distances and environmental conditions across varied IMTA configurations and longer timescales to fully capture ecosystem interactions and optimize production practices. Overall, this work supports the sustainable expansion of gilthead seabream and oyster aquaculture in Mediterranean waters, contributing to improved resource management and fostering the development of resilient, environmentally responsible aquaculture systems.

Author Contributions

E. Batır: investigation, methodology, validation, formal analysis, data curation, writing – original draft preparation, visualization. M. Yıldız: conceptualization, methodology, validation, writing – review and editing, supervision, project administration, funding acquisition. Ö. Metin: investigation, methodology. G. Papini: investigation, methodology. D. Pensa: investigation, methodology. M. Magdy: investigation, methodology. L. Grosso: investigation, methodology. A. Fianchini: methodology, project administration, funding acquisition. İ. Aydın: project administration, supervision. M. Rampacci: methodology, validation, supervision. A. Rakaj: conceptualization, methodology, validation, writing – review and editing, supervision, project administration, funding acquisition.

Funding

This research is funded by the projects namely ‘Project COREFISH CUP P2022ZPMXH_002’, ‘PO FEAMP, Implementing a full scale IMTA among finfish, shellfish and sea cucumbers: production diversification and waste bioremediation (INTEG‐MED‐8/INA/19)’ and ‘Scientific Research Projects Coordination Unit of Istanbul University, IMTA applications in marine aquaculture within the scope of blue transformation (BAP‐39729)’.

Disclosure

This study is an original work and does not incorporate material from external sources.

Ethics Statement

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

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