Reviving Oyster Aquaculture in Romania: A Scientifically Driven Study Aiming at Obtaining Missing-Link Data for Successful Production

Abstract

Oyster aquaculture offers promising opportunities for diversifying marine production in the Romanian Black Sea, where favorable environmental conditions and recent regulatory developments support shellfish farming. This study aimed to generate baseline data for the cultivation of the Pacific oyster, Magallana gigas, through an in situ experimental trial conducted off Mamaia Bay, Romania. A 50 m experimental long-line system was deployed at 13.5 m depth, and triploid oysters were cultured for one year at two depth horizons (3 m and 6 m). The growth performance, meat yield, Condition Index, microbiological quality, environmental parameters, and epibiotic communities were monitored monthly. Cultivation depth significantly influenced oyster growth, with individuals reared at 6 m consistently achieving a greater wet weight and shell length than those at 3 m. Growth rates peaked during spring, and meat yield values indicated good commercial quality. Environmental monitoring showed strong seasonal variability, with high summer temperatures and reduced dissolved oxygen associated with increased mortality. Microbiological analyses revealed higher bacterial loads during warm months. The Condition Index classified the oysters as generally “fine” to occasionally “special”. Overall, the results demonstrate that Black Sea conditions can support successful cultivation of M. gigas, although seasonal environmental stress and epibiosis require appropriate farm management.

1. Introduction

Most marine stocks are overfished or will be in the near future, while demand for seafood is constantly growing [1]. Governments, industry, and consumers are seeking to increase aquaculture production (including fish, mollusks, crustaceans, and aquatic plants) to balance wild extraction and seafood demand. In response, aquaculture has grown globally at an annual rate of about 9% and, in 2022, it surpassed wild capture for the first time, and it is estimated to meet about 60% of the world’s aquatic food needs by 2030 [2].

Magallana gigas (Thunberg, 1793) (formerly Crassostrea gigas, commonly known as the Pacific or Japanese oyster) is one of the most widely farmed oysters globally due to its fast growth, adaptability, and high market demand [3]. At species level, cupped oysters were listed among the most farmed, with about 6.2 million tonnes in production in 2022, closely following whiteleg shrimp (Penaeus vannamei), with 6.8 million tonnes produced [2]. China is by far the largest producer, with more than 5.8 million tonnes produced [4]. Europe, France, Spain and Italy are the main importers of oysters, while France is the main exporter [5]. France continues to be the EU’s top oyster producer, responsible for over 80% of total output, primarily along its Atlantic coastline, strongly recovering after the drop in 2019–2020 caused by a norovirus outbreak [6].

The Pacific oyster is a non-native species in the Black Sea, having been introduced for aquaculture purposes and subsequently established in various coastal regions of Bulgaria, Romania, Türkiye and Ukraine [7]. The species was first translocated in 1980 along the Crimean coast [8]. Since then, M. gigas has been documented in natural habitats [9], exhibiting a high degree of ecological plasticity and rapid adaptation to local environmental conditions, particularly along the Romanian coast [10].

Between 1997 and 1999, a mariculture initiative was implemented in Romania under the Black Sea Environmental Programme (BSEP), with funding from the PHARE program. As part of this project, a pilot hatchery was established at NIMRD Constanța [11]. Later on, during 2001–2003, in collaboration with a private company, experimental work focused on developing breeding and grow-out techniques for the Pacific oyster. The long-line culture system was identified as the most suitable method under Romanian coastal conditions [12].

Despite the attempts of a private operator, the further development of marine aquaculture in Romania was encumbered by administrative and legislative hindrances: the lack of microbiological classification of the Black Sea waters, as required by Regulation (EC) No. 627/2019, and the impossibility of leasing the water surface for the setup of farms at sea [12]. The microbiological survey of the areas proposed for classification was completed in the fall of 2020, with all three live bivalve mollusk production and relaying areas in the Romanian sector of the Black Sea (Perișor—Chituc, Mamaia Bay and Agigea—Mangalia) being rated as class A [13]. Moreover, the first allocated zones for aquaculture (AZAs) at the Romanian coast were identified by applying an internationally endorsed methodology which calculates the Degree of Compatibility (DC) both for finfish and shellfish [14]. Finally, water concession was settled in 2024 with the issuance of a legal act and now 28 lots are open for lease in inland maritime waters and territorial sea [15], which opens up immense opportunities for marine aquaculture on the Romanian coast.

Thus, Japanese oyster aquaculture presents a promising, albeit challenging, opportunity along the Romanian coastline. Environmental parameters, particularly salinity and seawater temperature, are generally conducive to the species’ growth. Although salinity levels are lower than those observed in open-ocean environments, the southern sector of the Romanian coast maintains relatively stable conditions, with average salinity values ranging between 17 and 18‰ [16], which are suitable for M. gigas cultivation. The thermal regime of the Black Sea, especially in its southern and western coastal zones, typically falls within the species’ tolerance range of 10–30 °C [17], with an annual mean temperature of approximately 14–15 °C [16]. In addition, the high trophic status of the region—driven primarily by nutrient inputs from the Danube River—results in elevated chlorophyll a concentrations, averaging over 7 μg/L [18], which further supports the growth of filter-feeding bivalves. Collectively, these conditions suggest that M. gigas is a viable candidate for aquaculture development in the area.

In this context, the present study aimed to generate essential baseline data for the development of Japanese oyster (M. gigas) aquaculture in the Black Sea through an in situ experimental trial conducted in the Mamaia Bay, Constanța, Romania. The primary objectives included assessing oyster growth performance and overall quality (meat yield, Condition Index), evaluating the effectiveness of different cage types, and identifying key operational challenges. Particular attention was given to the structural resilience of the long-line system under local hydrodynamic conditions, as well as to issues related to epibiosis and microbiological factors that may influence cultivation outcomes.

2. Materials and Methods

2.1. Study Area and Long-Line System Design

For the purposes of this trial, an experimental long line was assembled and deployed in September 2024 (Supplementary Material S1). A 50 m long-line installation was set up off the Mamaia Bay, in the northern part of Constanta, Romania, at a 3.5 km distance from the shore (coordinates of the central point: 44°14′14.92″ N, 28°40′31.43″ E) (Figure 1). The site is characterized by a sedimentary (sandy) substrate and a water depth of 13.5 m. The system was anchored using two 300 kg concrete bars (decommissioned railway beams) at each end. The headrope was made of Ø 32 mm polyethylene rope, while for the anchorages Ø 20 mm polyethylene rope was used. The lower part of the anchorage to the concrete beams was made of stainless steel chain to prevent breakage. The floatability of the entire installation was provided using seven inflatable navigation buoys. Subsequently, the experimental baskets/trays were connected to the headrope and launched at the desired depths.

2.2. Experimental Set-Up

The triploid M. gigas spat was provided by the hatchery Naturedulis s.r.l. (Goro, Italy) in two initial size classes: juveniles (J) (average 1 cm shell length) and half-grown (HG) oysters (average shell length 3.5 cm). Initial subsamples were measured and weighted to create a baseline for growth assessment (Supplementary Material S2). Upon arrival (October 2024), the J cohort was placed in protective net sacks (0.5 cm mesh size), to prevent any loss of individuals. Oysters were initially stocked in four Hexcyl^® baskets (Hexcyl Systems Pty Ltd., Ceduna, South Australia) (Supplementary Material S3) fitted with 2 kg ballast weights to ensure the maintenance of the position underwater. The stocking density was 700 individuals/basket for the HG oysters and 320 individuals/basket for the J spat. Two depth horizons were considered (two baskets at 3 m and two baskets at 6 m, respectively) for the deployment of the baskets and both the J and HG cohorts were introduced at these different depths, to allow a comparative assessment of the growth of Japanese oysters under local conditions. After three months of grow-out (October 2024–January 2025), the Hexcyl^® baskets were replaced with four Ostriga^® trays (ACQUA&CO S.r.l., Cadelbosco Sopra (RE), Italy) (Supplementary Material S4), similarly deployed at a 3 m and 6 m water depth to accommodate the increased biomass, dividing the oysters evenly among the tiers of each tray. Later on (June 2025), as the oysters started to grow steadily, the biological material was subsequently split, adding two more Ostriga^® trays (one at 3 m and another at 6 m) and at this point combining the J and HG cohorts.

2.3. Regular Maintenance and Monitoring

Regular overall monitoring was performed monthly, from October 2024 to October 2025. During each on-site check, the installation integrity was assessed, cages cleaned and oysters culled (Supplementary Material S5).

Growth rate (length, weight measurements, meat-to-shell ratio calculation) was assessed on random subsamples from each treatment (n = 30 individuals per treatment) until June 2025 (when the HG and J cohorts were combined), to ensure statistical consistency and reliability. For meat yield (MY) calculation, only HG oysters from the deepest horizon (6 m) were subsampled, as only these individuals had reached a size suitable for reliable shucking and accurate determination of meat and shell weights. MY measurements were conducted monthly between February–June 2025.

Total shell length (mm) and shell width (mm) were measured using a vernier caliper, and a Kern PBJ-N top-loading precision balance with readability d ≥ 0.01 g was utilized for total wet weight (g) determination.

Growth performance was evaluated calculating the specific growth rate, according to Ricker (1975) [19]:

$$SGR (\%/\mathrm{d}\mathrm{a}\mathrm{y})={\displaystyle \frac{lnWf-lnWi}{t \left(\mathrm{d}\mathrm{a}\mathrm{y}\mathrm{s}\right)}} x\times 100$$

(1)

where

Wf = oyster final mean wet weight (g);

Wi = oyster initial mean wet weight (g);

t = time of farming (days).

In order to detect seasonal growth dynamics, SGR was calculated relative to the biometry recorded at the timepoint immediately prior.

The meat yield (MY), as an indicator for commercial quality, was calculated according to the formula [20]:

$$Meat Yield \left(\%\right)=\left({\displaystyle \frac{Wet weight of oyster meat}{Oyster weight in shell}}\right)x\times 100$$

(2)

2.4. Environmental Condition Data

Temperature and salinity from the pilot long-line area were measured monthly using various techniques, including a reversible thermometer, titration, and a CastAway CTD multiparameter probe (SonTek Cast-Away-CTD, San Diego, CA, USA). Dissolved oxygen (DO) was measured by the Winkler method, a titrimetric technique for measuring dissolved oxygen in water [21].

For chlorophyll a, the trichromatic method was used. Seawater samples were collected using a bucket. One liter of seawater was stored in opaque containers and immediately filtered in NIMRD’s laboratory. Filtration was performed using mixed cellulose ester (MCE) membrane filters with a porosity of 0.45 microns. Once filtration was complete, the filter was folded with the biological material inside, blotted with absorbent paper to remove excess water, and placed in a clean and appropriately labeled plastic centrifuge [22]. The filters were then stored at −20 °C in the dark until analysis. Once the filters were thawed, 10 mL of 90% acetone was added to each plastic centrifuge tube. The tubes were then shaken vigorously using a Biosan vortex (Biosan Latvija SIA Rīga, Latvia) to ensure proper mixing. After this step, the filters were left overnight at 2–4 °C to facilitate pigment extraction. The following day, prior to spectrophotometric measurements, the filters were centrifuged at 4000 rpm for 20 min to obtain a less turbid extract. The sample extract was transferred from the 15 mL plastic centrifuge tube to the cuvette by careful pipetting. Then, the absorbance of the sample extract was measured at 750, 663, 645 and 630 nm against a 90% acetone blank. The chlorophyll a concentration was calculated according to the equations in the SCOR—UNESCO Report of Working Group 17 [23]:

$${\displaystyle \frac{\left(11.64\times A_{(750-663)}-2.16\times A_{\left(750-645\right)}+0.10\times A_{(750-630)}\right)\times v}{V}}\mu \mathrm{g}/\mathrm{L}$$

(3)

where

The molar extinction coefficients are 11.64, 2.16, and 0.10;

v is the volume of the extract in 90% acetone;

V is the volume of the seawater sample used in the analysis.

2.5. Microbiological Investigations

We investigated the following biological quality parameters in HG oysters over five months (June to October): total number of germs (TNG), total coliforms (TCs), fecal coliforms (FCs), Escherichia coli, Vibrio spp., Pseudomonas spp. and Salmonella spp. The total number of germs (TNG) was determined by inoculating culture medium in Petri dishes and incubating for 48 h at 37 °C, in accordance with ISO 4833-1:2013 [24] and the Bacteriological Analysis Manual [25].

The total number of coliforms (TC), fecal coliforms (FCs), and E. coli were determined using the most probable number (MPN) detection technique, in accordance with International Organization for Standardization (ISO) standard EN ISO 16649-3:2015 [26] and the Bacteriological Analysis Manual [27]. The tubes containing 9 mL of double-concentrated sodium lauryl sulfate broth medium were incubated at 37 °C for 48 h. If gas appeared in the lactose broth, it indicated that coliform bacteria were present in the samples studied. The characteristic number was obtained from the number of positive tubes in each of three successive, decimal dilution sets. Correlating this number with the McCrady table determined the number of microorganisms per gram of sample (according to ISO-7218:2024) [28], which was then multiplied by 100 to express the number of germs per 100 g of oyster meat and intervalvular liquid, in accordance with EN ISO 16649-3:2015 [26]. From these positive tubes, Levine EMB agar plates (SANIMED INTERNATIONAL IMPEX S.A., Bucharest, Romania) were inoculated to identify E. coli, and the plates were then incubated for 24 h at 44 °C.

To identify Vibrio bacteria, the meat and intravalvular fluid from the oysters were inoculated into alkaline peptone water (APW) and incubated at 37 °C for 48 h. Cultures that showed bacterial growth in APW were then replicated on agar plates with thiosulfate, citrate, bile salts, and sucrose (TCBS). The TCBS plates were inoculated from serial APW dilutions of the samples and incubated at 37 °C for 24 h. Following incubation, the plates were examined to identify colonies exhibiting the characteristic morphology of Vibrio species, in accordance with ISO 21872-1:2017 [29].

To identify Pseudomonas bacteria in oyster meat and intervalvular fluid, samples were inoculated into nutrient broth and incubated at 37 °C for 48 h. Cultures showing turbidity or visible growth were then replicated on MacConkey (MK) plates (SANIMED INTERNATIONAL IMPEX S.A., Bucharest, Romania). MK plates were inoculated using serial dilutions from the cultures that had tested positive and were then incubated at 37 °C for 24 h. Colonies exhibiting characteristic morphology were identified as Pseudomonas in accordance with ISO 13720:2010 [30].

The detection of Salmonella spp. was carried out in accordance with EN ISO 6579-1:2017 [31]. Samples of 25 g of meat and intravalvular fluid were pre-enriched in buffered peptone water at 37 °C for 18 h, followed by selective enrichment in Rappaport–Vassiliadis soy broth (RVS) at 41.5 °C for 24 h. Suspect colonies were isolated on Xylose Lysine Deoxycholate (XLD) agar and confirmed by biochemical tests. The results are expressed as the presence or absence of Salmonella spp. in 25 g of meat and intravalvular fluid.

Bacterial colonies grown for total germ counts, E. coli, Vibrio spp., Pseudomonas spp., were counted using the Sphere Flash automated colony counter. Colored smears were made from the specific colonies grown on the selective media using the Gram method.

Specific biochemical tests were performed on each bacterium of the group. The confirmation of all identified bacterial groups was carried out using the following biochemical tests: the oxidase test, motility test, indole production, H₂S production, and urea hydrolysis on MIU (Motility Indole Urease) medium; sugar reactions on TSI (Triple-Sugar Iron) medium; the malonate reaction in malonate broth; MILF (Motility Indole Lysine Fermentation) medium (SANIMED INTERNATIONAL IMPEX S.A., Bucharest, Romania) for the simultaneous determination of motility, indole production, lysine decarboxylation, and glucose fermentation; and indole reaction strips. The interpretation of the results was performed according to the specific biochemical characteristics of each identified bacterial genus.

For all groups investigated, the annotation “abs.” indicates the absence of characteristic colonies after incubation.

2.6. Condition Index and Proximate Composition

For proximate composition analysis, a random oyster subsample (N = 30, HG) was collected in August 2025 from the HG cohort at 6 m depth, as only these individuals had sufficient tissue mass for drying, ash content, and organic matter calculations. Oysters were manually shucked and the meat was separated from the shells with a scalpel. Their wet tissue mass (g) and shells (g) were weighed on an electronic balance (KERN KB 3600-2N) (KERN & SOHN GmbH, Balingen, Germany). The meat was dried in an oven at 105 °C for 24 h and the shells for 6 h until constant mass was achieved.

The Condition Index (CI) of each oyster was calculated according to Lucas and Beninger (1985) [32], as follows:

$$CI=(dry meat weight/dry shell weight)\times 100$$

(4)

The moisture percentage of the meat was calculated for each individual oyster according to the following formula:

Moisture (%) = [(vtotal weight − dry weight)/total weight] × 100

(5)

Ash was analyzed using the direct ashing-furnace method at 540 °C and referenced to the wet weight of the oysters. The organic substance content was calculated by subtracting the ash content from the dry meat weight.

2.7. Associated Epifauna Identification

The identification of the epibiont build-up on the cages, ropes and oyster shells was performed on a monthly basis, during the regular maintenance and cleaning, by removing the epibionts by soft brushing and gentle scraping under freshwater flow. The washed material was sieved through 90 μm mesh, fixed, and preserved in 4% formalin. Subsequently, all epibionts from each sample were identified and taxonomically quantified under an Olympus SZ61 stereomicroscope (Manufactured by Olympus Corporation, Tokyo, Japan) to the lowest taxonomic level possible [33]. Polychaetes were identified according to Surugiu [34,35], and the accuracy of taxonomy for each species was verified against the World Register of Marine Species [36].

2.8. Statistics

To evaluate the effect of cultivation depth on oyster growth performance, the data were analyzed separately for the two developmental stages (half-grown and juveniles). Differences among experimental groups were evaluated using Analysis of Variance (ANOVA), followed by Tukey’s Honest Significant Difference (HSD) post hoc test for specific pairwise comparisons. All statistical analyses were conducted using the R statistical software environment, version 4.3.2 [37]. The linear correlations between environmental parameters and overall mortality were assessed using Pearson’s correlation coefficient r in Excel (version 16x for Microsoft 365).

3. Results

3.1. Overall Resilience of the Long-Line Design and Associated Epifauna

The regular monitoring of the pilot facility revealed a sound structural design, adapted to the harsh local conditions. The long line withstood the difficult hydro-meteorological conditions of the Black Sea (in December 2024, waves of over 6 m were recorded on-site), demonstrating that proper design ensures functionality.

As a consequence of the characteristic high trophicity in the area, epibiosis development on the cages was one of the main issues requiring regular maintenance and cleaning (Supplementary Material S6). An overview of the associated epibiotic fauna is given in

Table 1. Overview of the associated epifauna that settled on the experimental long line.

Species	Settlement Location	Presence	Time
Amphibalanus improvisus (Darwin, 1854)	Baskets/trays, oyster shells	Constant	More abundant during the cold season (November–December)
Membranipora membranacea (Linnaeus, 1767)	Baskets/trays, oyster shells	Constant	Permanent
Ascidiella aspersa (Müller, 1776)	Oyster shells	Occasional	August–September
Calyptospadix cerulea Clarke, 1882	Ropes, trays, oyster shell	Seasonal (warm)	August–October
Mytilus galloprovincialis Lamarck, 1819	Ropes, oyster shells	Seasonal (warm)	June–October
Anadara kagoshimensis (Tokunaga, 1906)	Ropes, oyster shells	Occasional	August–September
Hediste diversicolor (O.F. Müller, 1776)	Oyster shells	Occasional	August–September
Polydora spp.	Oyster shells	Occasional	August–September
Cyanobacteria	Ropes, baskets/trays, oyster shells	Seasonal (cold)	March–May
Hippocampus guttulatus Cuvier, 1829	Trays	Occasional	August–September

below. The dark-colored Hexcyl^® baskets were rapidly colonized by barnacles (Amphibalanus improvisus) and bryozoans (Membranipora membranacea), being invaded by epibionts from the immersion (November–December). During January–February, the trays and ropes were mostly clean, with a cyanobacteria film starting to develop in March, with no major modification until late spring. In June, however, a significant black mussel (Mytilus galloprovincialis) spat settlement was observed mainly on the ropes, following the May spawning. Later during the summer months, young mussels started settling on trays and oyster shells as well. On the yellow Ostriga^® trays, the epibiotic organisms attached themselves as the water warmed, with the hydrozoan Calyptospadix cerulea being dominant during the peak development period (August–October 2025). Other epibionts that settled on the oyster shells in moderate quantities were juvenile bivalves Anadara kagoshimensis, and, occasionally, ascidians (Ascidiella aspersa). Additionally, during the warm months, polychaete worms (Hediste diversicolor) were sporadically identified on the shells, as well as scarce Polydora spp. mud blisters (but no actual worms were identified). The oyster aquaculture installation also provided attachment support for vertebrates—the long-snouted seahorse Hippocampus guttulatus.

A massive epibiosis outbreak occurred in September 2025: from barnacles, bryozoans and hydrozoans, to a new wave of mussel spat from a late-summer spawning, which required a thorough cleaning of the cages and individual oysters.

3.2. Oyster Growth Performance

The absolute growth parameters (wet weight and shell length) were analyzed separately for the two initial developmental stages: half-grown oysters (HG, 6 months old at seeding) and juveniles (J, 3 months old at seeding). For both cohorts, cultivation depth emerged as a highly significant factor driving biomass accumulation.

The HG oysters exhibited a robust growth trajectory throughout the 246-day trial. Individuals deployed at the 6 m depth horizon consistently outperformed those at 3 m. By June 2025 (Day 246), the HG cohort at 6 m reached a significantly higher final mean wet weight of 56.07 ± 1.45 g compared to 40.81 ± 1.94 g at 3 m (ANOVA followed by Tukey’s HSD, p < 0.05) (Figure 2a). A corresponding significant trend was observed for shell length, with the 6 m HG cohort achieving 74.83 ± 3.69 mm, compared to 68.67 ± 1.15 mm at the shallower depth (Figure 2b).

A similar depth-dependent growth pattern was recorded for the J cohort. Despite starting from a much smaller baseline, the J oysters deployed at 6 m showed a distinct growth advantage over the 3 m cohort, especially during the rapid spring growth phase. At the end of the monitoring period, the J oysters at 6 m reached a mean weight of 15.11 ± 3.06 g and a length of 46.17 ± 3.33 mm, significantly outperforming the 3 m group, which reached 9.85 ± 1.76 g and 43.08 ± 3.96 mm (Tukey’s HSD, p < 0.001) (Figure 3a,b).

The monthly evolution of the specific growth rate (SGR, %/day) highlights the relative performance of all four experimental treatments over time (Figure 4a,b). As expected from typical bivalve allometry, the J cohort maintained inherently higher SGR values compared to the HG oysters throughout the trial. However, the SGR curves clearly show that, within both developmental stages, the 6 m depth consistently supported higher relative daily growth. The SGR fluctuated seasonally, peaking during the highly productive spring months before stabilizing in early summer, reflecting the physiological adaptation of M. gigas to the local thermal and trophic regimes of the Black Sea.

During the experimental trial, the meat yield (meat-to-shell ratio, %) of the subsampled HG oysters ranged within 18.59% ± 0.44 and 27.70 ± 1.95, recording the peak value in May, when the biological material reached the age of 13 months (Figure 5). This is in line with growth rate results, indicating an excellent quality for commercialization of the oysters cultured under Black Sea conditions (Supplementary Material S7).

3.3. Environmental Parameter Evolution and Mortality

The environmental parameter monitoring spanned one year to provide a full seasonal cycle for interpreting oyster performance under Black Sea conditions (Figure 6).

As such, chlorophyll a (µg/L) measurements indicated a wide variation, between 0.75 µg/L (November) and 8.78 µg/L (February). The lowest values were recorded in autumn–winter, with a major peak in February and moderate levels in late spring–early summer. While the February peak suggests a winter phytoplankton bloom, summer levels were moderate despite the high temperature. There was a weak correlation with mortality (r ≈ −0.15), thus food availability does not appear to be a mortality driver for the experimentally reared oysters.

Temperature varied within a wide range, from 3.5 °C (February) to 26.0 °C (August). There was a gradual decrease from November (10.8 °C) to the winter minimum of February (3.5 °C), followed by a rapid warming from March onward. There was a heat peak in July–August (23–26 °C), with water cooling again in October (15.6 °C). The summer temperatures recorded approach thermal stress thresholds for M. gigas. Additionally, there was a strong positive correlation with overall mortality (r ≈ 0.83), indicating temperature is a major driver of summer mortality.

Salinity, on the other hand, was rather constant, ranging from 14.59‰ (November) to 19.75‰ (August). After a lower salinity in late autumn, there was a gradual increase through winter and spring, reaching a peak in summer, thus reflecting the seasonal hydrological balance between freshwater input (from rainfall and the Danube) and evaporation. There was only a moderately strong positive correlation with overall oyster mortality (r ≈ 0.64).

Dissolved oxygen (DO) saturation at the experimental long-line site ranged widely throughout the monitored period, from 24.9% to 75.1%. High oxygen levels were recorded in winter–early spring, with a decline during late spring and summer, with a severe minimum in July (only 24.9% saturation), which coincided with the peak oyster mortality. As such, there was a strong negative correlation with mortality (r ≈ −0.67), which suggests hypoxia is a critical stress factor during summer.

Oyster mortality was assessed globally throughout the entire period, as the HG and J oysters followed a similar pattern, regardless of depth horizon, with a minimum of 10 individuals in February and a maximum of 87 in July. There was a moderate mortality at the beginning of the trial, in November (45), with the lowest values in winter (10–16). An increase started in June (28), with a severe peak in July (87) and August (84). This mortality peak clearly coincides with the highest temperatures and the lowest dissolved oxygen values recorded on-site.

3.4. Bacterial Load

During the five months of bacterial load monitoring, the total number of germs ranged from 10² to 7 × 10³ CFU/g (

Table 2. Bacterial contamination of oyster meat and intravalvular fluid.

Bacteria Group/Period	June	July	August	September	October
Total number of germs (TNG) (CFU/g)	102	5 × 103	7 × 103	3 × 102	102
Total coliforms (MPN/100 g)	1.2 × 102	103	3 × 103	102	102
Fecal coliforms (MPN/100 g)	1.2 × 102	1.7 × 102	1.7 × 102	70	50
Escherichia coli (MPN/100 g)	abs.	50	1.3 × 102	20	abs.
Vibrio spp. (CFU/g)	102	103	104	102	50
Pseudomonas spp. (CFU/g)	50	102	103	102	abs.
Salmonella spp. (25 g)	abs.	abs.	abs.	abs.	abs.

Allowable limits as per EU legislation [38]: E. coli = 230 MPN/100 g of flesh and intravalvular fluid; fecal coliforms = 300 MPN/100 g of flesh and intravalvular fluid; Salmonella spp. = absence in 25 g.

). The minimum contamination was recorded in June and October (10² CFU/g), while increased contamination levels were observed in July (5 × 10³ CFU/g) and August (7 × 10³ CFU/g). Total coliforms ranged from 1.2 × 10² to 3 × 10³ MPN/100 g, with the highest levels recorded in July and August, with a decrease in contamination levels observed in the autumn months (10² MPN/100 g).

The contamination with fecal coliforms ranged from 20 to 170 MPN/100 g, with the highest values in July and August (170 CFU/100 g) and a decrease observed from September to October. E. coli reached its highest level of contamination in August (130 CFU/g), with no detectable presence during the autumn months. EU legislation levels were not detected to be exceeded throughout the sampling [38].

Between June and October, Vibrio spp. was identified in oyster meat and intravalvular fluid, with values ranging from 50 to 10⁴ CFU/g. The highest level of contamination was observed in August (10⁴ CFU/g), after which there was a decrease in subsequent months. Pseudomonas spp. was present from June to September, showing values between 50 and 10³ CFU/g, indicating an intensification of microbial activity during the warm season. Salmonella spp. was absent from all samples throughout the entire study period.

3.5. Condition Index and Proximate Composition

The Condition Index (CI) of M. gigas investigated ranged between 5.33 and 9.46%, with a mean CI (±standard deviation) of 7.02 ± 0.94% (Figure 7).

The proximate composition of oysters grown under experimental conditions is detailed below. The moisture content of M. gigas meat ranged between 78.83 and 84.95%, with a mean of 82.88 ± 1.32%. The dry weight (DW of oyster meat was 15.05–21.17%, with a mean of 17.12 ± 1.32%, while the ash content in meat ranged from 1.38 to 2.76%, with a mean of 1.73 ± 0.29%. The organic substance (OS) content in meat varied within 13.32–19.79%, recording a mean of 15.39 ± 1.32%.

4. Discussion

The experimental long-line installation showed good structural resilience under the energetic hydrodynamic conditions of the north-western Black Sea. The system remained stable during storm events with waves exceeding 6 m (Răzvan Mateescu, Department of Physical Oceanography and Coastal Engineering—NIMRD Constanța, personal observation, 2026), indicating that appropriately designed suspended culture structures, with adequate anchoring and buoyancy control, can operate in exposed coastal environments [39]. Biofouling rapidly developed on the submerged structures, confirming that aquaculture installations provide suitable hard substrate for colonization in the predominantly soft-bottom habitats of the region [40]. Barnacles and bryozoans were among the first colonizers, followed by seasonal settlement of mussel spat (M. galloprovincialis) and hydrozoans [41]. These taxa are typical components of fouling communities reported on artificial substrates along the Romanian and Bulgarian Black Sea coasts [42,43]. The seasonal appearance of mussel spat corresponds with the known reproductive cycle of M. galloprovincialis in the region, where spawning generally occurs from late spring to early summer [44]. Occasional occurrence of polychaetes such as Polydora spp. and H. diversicolor on oyster shells indicates the potential for interactions between cultured shellfish and benthic invertebrates [45]. Shell-boring polychaetes of the genus Polydora are known pests in oyster culture and have previously been reported in Black Sea mollusk populations [34,35]. Although their abundance in this study was low, continued monitoring is recommended in long-term farming operations.

The aquaculture structure also functioned as habitat for mobile fauna, including the protected long-snouted seahorse (H. guttulatus). In soft-bottom coastal environments such as the north-western Black Sea, aquaculture installations may increase local habitat complexity and provide refuge for associated species [46]. However, the strong epibiosis event observed in late summer highlights that fouling remains the main operational constraint in productive Black Sea coastal waters [47,48]. High nutrient availability and strong larval supply favor rapid biomass accumulation on farming equipment, making regular maintenance essential for sustainable shellfish culture [49,50]. Epibiosis likely contributes indirectly to stress by increasing metabolic demand and reducing feeding efficiency, particularly under already unfavorable summer conditions. While not a direct cause of mortality, fouling may exacerbate physiological stress and should be considered an important secondary factor in farm management, with quantitative assessments (coverage/biomass) included in future monitoring.

Regarding the growth pattern observed, cultivation depth significantly influenced the growth performance of M. gigas, with oysters deployed at 6 m consistently exhibiting higher biomass accumulation and shell growth compared to those maintained at 3 m. This depth-related pattern was observed in both the HG and J cohorts, suggesting that the deeper cultivation horizon provided more favorable environmental conditions for oyster growth [51]. Similar depth-dependent growth responses have been reported in suspended oyster culture systems, where moderate depths often provide improved feeding conditions and reduced environmental stress compared to near-surface layers [52,53,54,55,56].

The enhanced growth at the 6 m depth may be related to several environmental factors typical of stratified coastal systems [57]. In suspended bivalve culture, deeper layers frequently exhibit more stable temperature regimes, reduced hydrodynamic disturbance, and higher concentrations of suspended particulate matter, which can enhance feeding efficiency and energy allocation to growth [58]. In the Black Sea, productive phytoplankton blooms occurring during spring provide abundant food resources for suspension-feeding bivalves [59], which likely explains the seasonal peak in specific growth rate observed during this period [60,61,62].

The higher specific growth rates recorded in the J cohort are consistent with the well-established allometric growth pattern of bivalves, whereby smaller individuals exhibit faster relative growth rates than larger oysters. This trend has been widely reported in oyster aquaculture studies and reflects the physiological scaling of metabolic processes during early developmental stages [63]. Similar seasonal growth patterns have been documented in M. gigas populations cultivated in other temperate environments, where growth accelerates during periods of high primary productivity and favorable temperature conditions [64,65,66,67].

Meat yield values recorded in this study indicate good physiological condition and commercial quality of the oysters cultured under the tested conditions. Much lower meat-yield ranges have been reported for market-sized M. gigas cultivated in suspended culture systems worldwide [68,69,70,71], thus underlining the importance of the Black Sea’s high trophicity. The peak recorded in late spring (27.7%) likely corresponds to the accumulation of tissue reserves prior to the reproductive season, a common pattern in oyster populations [20].

The analysis of growth data strongly suggests that suspended culture at approximately 6 m depth provides favorable conditions for oyster growth in the studied Black Sea environment, highlighting the importance of optimizing vertical positioning within the water column to maximize aquaculture productivity. Our results indicate that although both factors (oyster age and water depth) influence growth, cultivation depth exerted a consistently stronger and more uniform effect across both cohorts (juveniles and half-grown oysters). This is evidenced by the parallel growth trends observed at 6 m compared to 3 m, regardless of initial size. The effect of cultivation depth on growth performance was markedly stronger than that of initial developmental stage, indicating that vertical positioning in the water column is the primary driver of biomass accumulation under Black Sea conditions. This conclusion is supported by the consistently higher wet weight and shell length values at 6 m for both cohorts.

The environmental conditions recorded during the monitoring period reflected the strong seasonal variability typical of the north-western Black Sea and played a major role in shaping oyster survival patterns. Temperatures ranged from 3.5 °C in winter to 26 °C in summer, while salinity remained relatively stable within the brackish range characteristic of the basin environmental parameters [72]. Such environmental variability is typical for coastal areas influenced by the Danube River, where seasonal freshwater input and atmospheric forcing strongly affect water-column properties [73,74]. The strong positive correlation between temperature and oyster mortality indicates that summer thermal stress was a major driver of mortality events. Water temperatures approaching 25–26 °C are known to represent the upper tolerance range for M. gigas, particularly when combined with other stress factors such as low oxygen availability [75,76]. Similar temperature-related mortality episodes have been documented in oyster farming areas worldwide and are often associated with metabolic stress and increased susceptibility to disease [77,78,79].

Dissolved oxygen levels also showed a strong relationship with mortality, reaching a minimum of about 25% saturation during the summer period, when oyster mortality peaked. Hypoxic events are relatively common in eutrophic coastal environments and may occur during periods of high temperature and elevated biological oxygen demand [80,81]. In the Black Sea, seasonal stratification and high productivity frequently lead to oxygen depletion in nearshore waters during summer months, potentially creating stressful conditions for benthic and cultured organisms [82]. The strong negative correlation between oxygen saturation and mortality observed in this study therefore suggests that hypoxia, in combination with elevated temperature, likely contributed to the observed mortality peak.

In contrast, chlorophyll a concentration, used as an indicator of phytoplankton biomass and food availability [83], showed only weak correlation with mortality. This suggests that food limitation was not a primary driver of oyster mortality under the studied conditions. Instead, seasonal phytoplankton dynamics, including the winter bloom observed in February, likely supported oyster growth during cooler periods when metabolic stress was reduced [84].

The environmental monitoring performed suggested a visible ranking among stressors. Temperature showed the strongest relationship with mortality, identifying it as the dominant stressor. Dissolved oxygen had a strong negative correlation, indicating hypoxia as a secondary but critical factor. Salinity showed a moderate effect, while chlorophyll a had only a weak contribution (r ≈ −0.15). Understanding these seasonal stressors is essential for optimizing farming practices, including site selection, stocking density, and harvesting schedules in regional oyster aquaculture [85].

The experimental oyster farming facility is located in a microbiologically suitable area, as Mamaia Bay is classified as class A according to EU Regulation 629/2019 [13,14,15], indicating that shellfish harvested from this zone are fit for direct human consumption under current regulatory standards. The microbiological analysis of M. gigas revealed a clear seasonal pattern, with higher levels of total bacteria, total coliforms, fecal coliforms, and E. coli recorded during summer and lower levels in autumn. This trend is consistent with previous findings [86] and can be explained by the combined effect of elevated seawater temperature and enhanced oyster filtration rates, which promote bacterial proliferation and accumulation in shellfish tissues during warmer periods [87,88,89]. Conversely, the reduction observed in autumn likely reflects both decreased bacterial survival in cooler conditions and the natural depuration capacity of oysters [90]. Similar seasonal dynamics have been widely reported in shellfish farming areas, confirming the strong influence of environmental conditions on microbiological parameters [13,89,91,92,93,94].

Bacteria of the genus Vibrio, commonly associated with oysters, are autochthonous components of marine ecosystems and should be regarded as part of the natural microbiota of seawater and marine organisms. Although often discussed in relation to human pathogenicity, their presence is not inherently indicative of contamination, as these bacteria are endemic to marine environments and are expected to occur in healthy oysters [95,96,97]. In the present study, Vibrio spp. were detected between June and October, with concentrations ranging from 50 to 10⁴ CFU/g and peak values in August. This seasonal increase coincides with the highest recorded seawater temperatures and salinity values at the study site and is consistent with previous studies demonstrating that Vibrio abundance is strongly driven by environmental factors, particularly temperature and salinity [98,99,100]. Comparable seasonal trends have been reported in European coastal waters, further supporting the role of environmental conditions in regulating Vibrio dynamics [101,102,103]. Importantly, the results refer to total Vibrio spp., and no differentiation of pathogenic species was performed, which should be considered when evaluating potential public health implications.

Pseudomonas spp. were identified in M. gigas between June and September, with concentrations ranging from 50 to 10³ CFU/g, showing a similar seasonal pattern with higher values during warmer months. This observation is in agreement with previous studies indicating that Pseudomonas spp. are common components of oyster-associated microbial communities and tend to increase in abundance under favorable environmental conditions, particularly higher temperatures and nutrient availability [88,104,105,106]. Their presence reflects the dynamics of heterotrophic bacterial populations in the surrounding marine environment rather than a specific contamination source.

Overall, the results indicate that oyster farming under the studied conditions maintains good microbiological quality, with seasonal increases in bacterial abundance primarily driven by natural environmental variability. The observed values are consistent with those reported in comparable marine aquaculture systems and should be interpreted within the ecological context of coastal environments, where temperature, salinity, and biological activity govern microbial dynamics.

The CI is a term for evaluating the edible parts of shellfish and is known to be an important factor in aquaculture production and marketability [107]. This index is correlated with the meat yield, which declines in bivalves under stressful environmental conditions which require significant energetic consumption [108]. The fluctuation of the CI is associated with the reproductive condition or environmental parameters (temperature, salinity and nutrition) of bivalves [109]. The CI varies according to the season and the region where shellfish are cultivated [110]. An index between 6.5 and 9 corresponds to “fine” oysters, while a higher one rates them as “special” oysters [111]. While a CI < 5.5 was reported in Northern Ireland [112] and a CI < 7.5 in Gadeok-do Korea [113], the oysters from the Romanian Black Sea pilot long-line system recorded a CI which classifies them in general as “fine” and, in some cases, “special” oysters.

The proximate composition, including moisture, ash, protein, lipids and minerals, can be used as a measure of nutritional quality of food [114], and seafood contains high levels of these important components compared to other meats [115]. Oysters are known as a natural source of these nutritive compounds [116]. For the same season (summer), M. gigas from Gadeok-do, Korea, recorded mean values of 70.67% moisture, 29.33% dry weight, 7.61% ash and 21.72% organic substances [113], which differ slightly from Romanian samples due to different habitat conditions.

The cycle of energy storage and use varies depending on the species and habitat, and the biochemical composition varies according to the season [117]. Water constitutes the major part of oysters [118]. This component is linked to juiciness, which is an important sensory trait of oysters [119]. The proximate composition of M. gigas reared under Black Sea conditions indicates a good moisture content, which is linked to juiciness and oyster marketability, making them an attractive product for the local seafood market.

5. Conclusions

This study provides the first integrated assessment of Pacific oyster cultivation under Romanian Black Sea conditions, demonstrating that offshore suspended culture is both technically feasible and biologically effective in this environment. The experimental long-line system showed good structural resilience, confirming its suitability for exposed coastal conditions.

The use of triploid oysters, which are functionally sterile and therefore allocate more energy to growth rather than reproduction, proved advantageous for aquaculture purposes. This results in faster growth rates, improved meat quality, and more stable commercial value throughout the year, making them particularly suitable for emerging farming sectors such as the Black Sea.

Among the tested conditions, cultivation at 6 m depth is clearly recommended, as it consistently produced superior growth performance compared to shallower deployment. Growth dynamics followed a strong seasonal pattern, with rapid biomass accumulation during spring, driven by favorable trophic conditions. In contrast, summer represents a critical period, where elevated temperatures, reduced dissolved oxygen, and intense biofouling require targeted management measures. These should include strategies to mitigate mortality (e.g., optimized stocking density or harvest timing) and regular control of epibiosis to maintain culture efficiency.

From a production strategy perspective, both juvenile and half-grown oysters proved suitable for cultivation, but with different operational implications. Juveniles exhibited higher relative growth rates, while half-grown oysters reached market size more rapidly. This suggests that starting with half-grown oysters may be more cost-effective in reducing production time and exposure to environmental risks, particularly during the first critical months. However, juvenile stocking may offer advantages in terms of lower initial investment and greater flexibility, especially for scaling production. The choice between the two should therefore be aligned with farm management strategy, available resources, and risk tolerance.

From an applied perspective, the results indicate that oyster farming in the Romanian Black Sea can achieve good biological performance and product quality, supporting its potential for commercial development. However, operational challenges such as biofouling and seasonal environmental stress must be carefully managed. Microbiological analyses showed seasonal increases in bacterial loads during warm months, but overall contamination levels remained within ranges commonly reported for shellfish farming environments. Importantly, the observed patterns support a clear environmental control mechanism, in which high summer temperatures combined with hypoxia drive increased mortality, subsequently associated with elevated microbial loads, rather than microbial contamination acting as a primary cause.

Although the present study focuses primarily on biological and environmental aspects, some key elements of technical and economic feasibility are evident, including system robustness, maintenance requirements, and production potential. A comprehensive economic analysis (covering investment costs, operational expenditure, and profitability scenarios) is beyond the scope of this study but represents an essential next step. This is particularly relevant in the context of the recently established aquaculture licensing framework in Romania, which creates new opportunities for scaling up marine farming activities.

Overall, the findings highlight that with appropriate site selection, depth optimization, and seasonal management, the Romanian Black Sea offers favorable conditions for the successful development of oyster aquaculture, while also emphasizing the need for adaptive management strategies to address predictable seasonal stressors.