Next Article in Journal
Resveratrol Promotes Proliferation, Antioxidant Properties, and Progesterone Production in Yak (Bos grunniens) Granulosa Cells
Next Article in Special Issue
Effects of Photoperiod on Survival, Growth, Physiological, and Biochemical Indices of Redclaw Crayfish (Cherax quadricarinatus) Juveniles
Previous Article in Journal
The Impact of a Proprietary Blend of Yeast Cell Wall, Short-Chain Fatty Acids, and Zinc Proteinate on Growth, Nutrient Utilisation, and Endocrine Hormone Secretion in Intestinal Cell Models
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Animals
Volume 14
Issue 2
10.3390/ani14020239
animals-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Influence of Pediveliger Larvae Stocking Density on Settlement Efficiency and Seed Production in Captivity of Mytilus galloprovincialis in Amsa Bay, Tetouan

by
Hafsa Janah
1,2,
Adil Aghzar
1,
Pablo Presa
3 and
Yassine Ouagajjou
2,*
1
Research Team of Agriculture and Aquaculture Engineering (G2A), Polydisciplinary Faculty of Larache, Abdelmalek Essaadi University, Tetouan 93000, Morocco
2
Amsa Shellfish Research Station, National Institute of Fisheries Research, Tetouan 93000, Morocco
3
Laboratory of Marine Genetic Resources (ReXenMar), CIM—Universidade de Vigo, 36310 Vigo, Spain
*
Author to whom correspondence should be addressed.
Animals 2024, 14(2), 239; https://doi.org/10.3390/ani14020239
Submission received: 6 December 2023 / Revised: 7 January 2024 / Accepted: 10 January 2024 / Published: 12 January 2024
(This article belongs to the Special Issue Advancing Sustainable Aquaculture: Enhancing Production Methods, Innovating Feeds, Promoting Animal Welfare and Minimizing Environmental Impact)
Browse Figures
Versions Notes

Abstract

:

Simple Summary

The development of cost-effective protocols with optimized rearing systems and conditions is an essential requirement to assure the economic feasibility of mussel production in hatchery systems. The Mediterranean mussel Mytilus galloprovincialis plays a relevant role on the scale of bivalve aquaculture, whose expansion has deep economic and environmental consequences. The current study establishes the protocols to improve pediveliger settlement, post-larvae production and growth, through the optimization of the stocking density of larvae and post-larvae in hatchery systems. This standardized protocol for mussel rearing can enhance the production of quality seed to supply the fattening industry as well as for environmental restoration.

Abstract

In mussel hatchery systems, the settlement process is a crucial element influencing seed yield. The current study assayed the influence of five densities of competent pediveliger larvae on settlement success and post-larvae production. We showed an inverse relationship between density and settlement efficiency, e.g., an attachment success of 99.4% at the lowest density (35 larvae/cm2) but only 9% at the highest density (210 larvae/cm2). However, post-larvae production was higher at intermediate larvae densities (70 larvae/cm2). The reimplementation of treatments upon post-larvae density after 6 weeks post settlement showed that the lowest-density groups bore both the highest post-larvae growth rate (22.24 ± 4.60 µm/day) and the largest head batch (48% of the size distribution), as compared to the higher-post-larvae-density groups. These results highlight the importance of optimizing both pediveliger larvae density and post-larvae density, to maximize high-quality seed yield in local hatcheries. Current rearing technologies would assure a timely commercial seed production to protect natural sea rocky beds in Alboran Sea coasts.

1. Introduction

In recent times, bivalve hatchery production has attracted high interest due to its potential to mitigate environmental hazards caused by human activities [1,2,3]. Also, the rapid human growth demands more shellfish supply [4], which has begun to exert pressure on the bivalve industry due to its limited availability [5,6]. Traditionally, mussel seed supply for aquaculture production was supported by the hand scrapping of natural seed from rocky shores [7]. However, the unsustainable exploitation of intertidal habitats has led to a serious decrease in natural populations and a reduction in ecosystem diversity [7,8]. Thanks to the advancement in bivalve husbandry, multiple opportunities exist nowadays for the development of reliable and certified mussel seed, to cope with both the industrial demands and the rehabilitation of overexploited habitats [5].
The Mediterranean mussel Mytilus galloprovincialis is native to the Mediterranean Sea [9,10,11,12,13] and is the most distributed of its genus worldwide [9]. Studies have suggested that this mussel expanded on the Atlantic coast of Morocco [14,15,16], where its commercial production has recently been launched [17]. For instance, several ongoing research initiatives are being developed in Morocco because of biological, environmental and commercial concerns to assess the viability and pertinence of the promotion of the mussel industry. From the genetic insight, the dispersal pattern and connectivity of its Atlantic metapopulation have been described [16,18], as well as its coexistence with the green mussel Perna Perna in northern Africa [19]. From an ecological perspective, the suitability of this mussel as a bio-monitor of environmental fluctuations has been shown [20,21,22,23]. Also, investigations of its growth performance in different culture systems suggest that the offshore mussel technology (longline systems) can assure higher yield and is deemed to a convenient solution for seed supply [17,24,25].
The reproductive cycle of this species has been studied on a world scale for a better understanding of its ecophysiology [26,27], biology [28,29] and genetics [30]. For instance, broodstock conditioning in hatchery systems is key to improve the quality and quantity of commercial stocks [31]. Those studies have revealed a high potential innovation in mussel aquaculture development [32], especially to produce juveniles and reduce pressure on natural populations [33]. It is well known that mussel larval rearing is directly influenced by temperature [34], feeding regimes [35,36,37], stocking density [38,39], bacterial loads [40] and CO2 levels [41], all of which improve or worsen the final larvae performance. Irreversible physiological processes in the pediveliger larvae of bivalves lead to the transition from planktonic to benthic life throughout metamorphosis and settlement [42]. Much research has been devoted to those latter processes because of their importance for the viability of populations. For instance, the exposure of M. edulis and M. galloprovincialis pediveliger larvae to microbial films resulted in the induction of metamorphosis and further settlement [43]. Those processes are also influenced by chemical cues such as in the silver-lip pearl oyster Pinctada maxima [44], by diet composition such as in M. galloprovincialis larvae [45,46] and by stocking density such as in clam juveniles and many other invertebrates [38,47]. Namely, larvae stocking density adds to the intra-specific competition for space and food in marine organisms [48,49] and, therefore, it is a key factor influencing settlement success, as shown in pediveliger larvae of the pearl oyster [50], as well as in the clam Meretrix meretrix [51].
Stocking density in Mytilus galloprovincialis has been shown to be a primary factor influencing not only the performance of post-larvae cultured in captivity but also that of mussel spat cultured in longline systems. For instance, increasing the stock density decreased the production efficiency and increased the fattening time to reach the commercial size in M. galloprovincialis cultured in the Black Sea [52]. A negative effect of a high post-larvae density on bivalve seed performance has also been reported in other species where the highest post-larvae growth rate was achieved under their lowest initial density [53]. In addition, pilot experiments on mussel broodstock from La Atunara (southwestern Spain) using static hatchery systems have shown that a medium larvae density (25 larvae/mL) assured good growth and high seed yield [54]. Given the multifactorial influence on larvae production and performance such as organic matter accumulation, it is also recommended to evaluate the influence of post-larvae density on the posterior mussel performance in flow-through systems [54]. Therefore, in this work, we assess how the stocking density of competent pediveliger larvae affects their settlement rate and post-settlement production. We also address the effect of post-larvae density on the growth rate and size distribution over a 6-week culturing experiment. This knowledge would help to optimize the large-scale production of mussel seed to cope with incipient industrial needs as well as for the restoration and enhancement of natural habitats in the Mediterranean region in conjunction with genetic background knowledge.

2. Materials and Methods

2.1. Spawning, Fertilization and Larval Rearing

Mytilus galloprovincialis broodstock originated from longline rearing systems in Amsa Bay, 35°31′59.5″ N 5°13′29.0″ W, Tetouan (Morocco). On 19 May 2022, fifty adult specimens from each sex were placed alternatively in hot (26 ± 1 °C) and cold (14 ± 1.5 °C) sea water baths for several cycles until they begun spawning and were placed in individual beakers with filtered and UV-treated seawater. Oocytes and spermatozoids were filtered separately through a 20 μm sieve, and oocyte quality (roundness) and spermatozoid motility were assessed under a binocular microscope. The density of gametes was assessed in a Malassez counting chamber (BR719005-1EA). The fertilization was performed in a 200 L polycarbonate tank using a ratio of 120 spermatozoids per oocyte. Total number of fertilized oocytes was defined as the average no. of fertilized oocytes from three sub-samples (25 µL) using a binocular microscope. Eggs were placed in a 1000 L tank containing filtered (0.2 µm) and UV-treated seawater at an initial density of 15 eggs/mL at 20 °C. Larvae feeding began from 48 hpf (hours post fertilization) to 17th dpf (days post fertilization). Food consisted of a microalgae mixture of increasing density with rearing age, i.e., 500–3000 cell/larvae of Isochrysis galbana, 500–1500 cell/larvae of Chaetoceros calcitrans and 500–1000 cell/larvae of Tetraselmis suecica.

2.2. Experimental Design

A panoramic view of the experimental design is given in Figure 1.
After the 17th dpf of larval rearing (pediveliger stage), larvae became competent for settlement and were distributed at a predesigned experimental density (Figure 2a). The settlement process was monitored in a series of cylinders (300 mm of diameter) bottomed with a 150 µm nylon mesh and placed into a 1200 L rectangular fiberglass tank. The rearing system (recirculating aquaculture system: RAS) provided filtered seawater, continuous aeration and controlled temperature (21.7 ± 0.69 °C) until the end of the experiment (Figure 2b). Post-larvae were fed a mixture of three strains with the total number of cells per milliliter varying progressively from 16,000 to 20,000 cell/mL of I. galbana, 4000 to 35,000 cell/mL of C. calcitrans and 4000 to 25,000 cell/mL of T. suecica. The physicochemical parameters recorded during the settlement process were pH (8.73 ± 0.08), salinity (36.75 ± 0.51 psu) and dissolved oxygen (7.08 ± 0.12 mg/L). Feeding, regular water renewal and cleaning of the culture system were performed every 48 h during the six weeks of the experiment (from 4 June to 16 July 2022). Settled post-larvae from each container were washed out, gently collected and counted using a profile projector (Nikon V-12B).
After 17 days of larvae rearing, the 1 m3 tank was sieved through a 150 µm mesh. The larvae were washed with microfiltered water and then resuspended in 1 L filtered-UV water. The no. of pediveliger was counted with a binocular on 3 samples of 25 µL each to obtain its concentration per mL. That density (A, No. of pediveliger/mL) was used to estimate the total no. of pediveliger (Nx) in the 1 L recuperation volume (Vr), as follows:
Nx = A × Vr
About 4 million pediveliger larvae were produced and the corresponding aliquot was taken for each density treatment (Dx). Several larvae stocking densities were enforced in five treatments with replicates (rep), i.e., low density: DL = 35 larvae/cm2 (4 rep), medium density: DM1 = 70 larvae/cm2 (5 rep) and DM2 = 100 larvae/cm2 (8 rep); and high density: DH1 = 140 larvae/cm2 (5 rep) and DH2 = 210 larvae/cm2 (5 rep).
Each density treatment was defined using the following formula:
Dx = Nx/S
where Dx is the density treatment, Nx is the number of pediveliger larvae needed to reach density x, and S is the surface of cylinder bottom (700 cm2).
The volume distributed to reach the required density treatment was calculated as
Vx = (N1 × V1)/Nx
where Vx is the required volume to reach the density Dx; N1 is the initial number of pediveliger larvae; V1 is the total volume of pediveliger larvae; Nx is the number of pediveliger larvae needed to reach Dx.
The settlement rate (Sr), defined as the percentage of post-larvae settled on the nylon mesh cylinder (Figure 3a), was estimated on each replicate using the following formula:
Sr (%) = (Nf/Ni) × 100
where Sr is the settlement rate; Nf is the final number of produced post-larvae; Ni is the initial number of seeded larvae.
At the end of the 6-week experiment, the settled post-larvae from each nylon-mesh cylinder (Figure 3b) were collected and weighted and the final number of post-larvae (Nf) was calculated using the following formula:
Nf = (Wt/Wi)
where Nf is the final number of produced post-larvae; Wt is the total weight of settled post-larvae per replicate; Wi is the individual average weight per post-larvae as estimated using a random subsample and the following expression:
Wi = (Wsub/Nsub)
where Wsub is the total weight of the subsample per replicate; Nsub is the total number of post-larvae per subsample and replicate. The post-larvae production per cm2 was calculated as
Pr (post-larvae/cm2) = (Nf/S)
where Pr is the post-larvae production (post-larvae/cm2); Nf is the final number of produced post-larvae; S is the surface area of cylinder mesh (cm2).
To assess growth in shell length, 45 pediveliger larvae and 75 post-larvae were measured per treatment using a profile projector (Nikon V-12B). The growth rate was calculated using the following formula:
Gr = (M1 − M0)/t
where Gr is the growth rate; M0 is the initial average length of pediveliger larvae; M1 is the final average length of post-larvae; t is the number of days between the beginning and end of the experiment (42 days).
The five post-larvae production densities (Pr) obtained, i.e., 35, 50, 38, 26 and 20 post-larvae/cm2, were regrouped in three major density groups: low (DF1 = 20–26 post-larvae/cm2, 10 containers, five from each DH1 and DH1), moderate (DF2 = 35–38 post-larvae/cm2, 12 containers, four from DL and eight from DM2) and high (DF3 = 50 post-larvae/cm2, 5 containers from DM1). All post-larvae within those three density groups were length-classified using two sieves of different meshes (1.0 mm and 1.3 mm) to produce a head batch >1.3 mm, a modal batch = 1.0–1.3 mm and a tail batch < 1 mm. The growth rate was estimated on 75 randomly sampled individuals from each batch.

2.3. Statistical Analyses

One-way ANOVA analyses (Fisher test, p ≤ 0.05) as corrected with the Welch test [55] were applied to determine the effect of stocking density on post-larvae performance, settlement rate and percentage of size classes (the head, modal and tail batches). Boxplots were built to compare settlement rate, post-larvae production and size classes from each post-larval density. All the statistical analyses were conducted using R software, version 2021.05.29, Rcmdr package, version 2.7.2 and RStudio version 2022.12.0+353 [56].

3. Results

3.1. Effect of Density on Post-Larvae Settlement (Sr)

The settlement rate decreased with larvae density in a negative exponential trend (r2 = 0.98; y = 1.6452 × 10−0.014x) (Figure 4a). The highest settlement (99.4%) was observed at the lowest larvae density (DL = 35 larvae/cm2), whereas the lowest settlement (9%) was observed at the highest density (DH2 = 210 larvae/cm2).
The ANOVA showed that the initial larval density significantly influenced larvae settlement (F = 93.45; df_num = 4; df_den = 8.9; p < 0.001) (Table 1). The settlement rate differed significantly between treatments except between the two highest ones, DH1 and DH2.

3.2. Effect of Density on Post-Larvae Production (Pr)

Density DM1 exhibited the highest post-larvae production (Pr, DM1 = 50 post-larvae/cm2), whereas DH1 and DH2 showed the lowest productions, i.e., 26 and 20 post-larvae/cm2, respectively (Figure 4b).
The initial larval density significantly influenced post-larvae production (F = 25.73; df_num = 4; df_den = 9.08; p < 0.001) (Table 2). Among the densities, a major difference in post-larvae production was observed between DM1 and the rest of the densities, whereas no difference was detected between (DL–DM2) and (DH1–DH2).

3.3. Effect of Stocking Density on Post-Larvae Shell Length Classes

Three post-larvae shell length batches were obtained from each post-larvae density distribution (DF1, DF2, DF3) (Figure 5). The lower the post-larvae density, the higher the share of the head batch, e.g., DF1, head batch = 48.19%. By contrast, the higher the post-larvae density, the higher the share of the tail batch, e.g., DF3, tail batch = 26.70%.

3.4. Effect of Stocking Density on Post-Larvae Growth (Gr)

Significant differences in growth rate were observed between spat groups from different post-larvae densities (F = 16.683; df_num = 2; df_den = 250.97; p < 0.001). The highest growth rate (Gr) (22.24 ± 4.60 µm/day) was observed in the lowest-density treatment (DF1) (Table 3).
One-factor ANOVA on post-larvae growth rate (µm/day ± SD) within the shell length batch (H, M, T) was significantly influenced by post-larvae density (DF1, DF2, DF3) in the head and tail batches (F= 31.66; df_num = 2; df_den = 89.42; p < 0.001 and F = 6.72; df_num = 2; df_den = 20.06; p < 0.01, respectively). No effect of post-larvae density was observed on the growth rate of the modal batch (F = 0.20; df_num = 2; df_den = 105.85; p = 0.83) (Table 4). The highest growth across treatments was observed in the head batches of the lowest post-larvae densities, i.e., DF1 and DF2 (Table 4).

4. Discussion

In mussels as in other bivalves, the transition from a planktonic to a sedentary life requires a sudden increase in food and space [57]. These needs make mussel nursery particularly challenging biologically as well as economically [33]. Several studies have been conducted to optimize early life cycle phases in both experimental pilot assays and commercial nursery systems [58,59,60,61,62]. The success of the settlement process is critical for seed production in quality and quantity and depends on multiple biological and physical factors. Particularly, stocking density is one of the key manageable factors able to enhance productivity [63]. The current study shows the correlation of pediveliger larval density on post-larvae settlement, production and growth in a hatchery RAS in the Alboran Sea coasts of northern Morocco (Amsa Bay).
The negative correlation observed between larvae stocking density and settlement rate suggests that seeding M. galloprovincialis culture substrates with more than ~35 larvae/cm2 (Sr = 99%) would not improve the settlement rate under local conditions. Moreover, increasing the density above 70 larvae/cm2 makes the settlement rate decay below 50%. Such a negative correlation between density and post-larvae settlement has been reported in numerous studies on bivalves. For instance, in the Asiatic hard clam Meretrix meretrix, more than 50% larvae were successfully settled [51], but settlement was delayed at higher densities (40–60 larvae/mL). The number of metamorphosed and settled larvae decreased at high stocking densities so that fewer larvae were able to release chemical cues to induce metamorphosis in the rest of the floating larvae [64]. The low post-larvae settlement under high larvae densities can result from space limitation as an essential element to optimize bivalve production in hatchery systems. For instance, space limitation in silver-lip pearl oyster juveniles (Pinctada maxima) provoked an aggregate settlement phenomenon which significantly reduced settlement rates [50]. In the zebra mussel (Dreissena polymorpha), it is believed that the aggregation phenomenon at high densities inhibits larvae site-selection as a natural behavior of mussel larvae prior to settlement [65]. Such behavior is time-limited, and consequently, unsettled larvae die soon afterwards [66].
The high settlement rates scored in the current RAS flow-through system (36–99.4%) are unexpectedly high as compared to previous observations, e.g., 6.3% settlement in Black-Lip Pearl Oyster (Pinctada margaritifera) [67], 50% in the clam Asiatic hard clam (Meretrix meretrix) [51] and >39% in the clam (Venerupis pullastra) in response to chemical induction [68]. It is worth noting that the flow generated in downwelling systems such as the current one, which provides velocity and turbulence, has been shown to be correlated to settlement success [69]. Previous studies were conducted on mussel settlement in response to different rearing factors. For instance, the highest settlement (49.3%) in the green mussel Perna viridis was achieved at a rearing temperature of 29 °C [70]. Also, velon screen was reported as the most suitable substratum for the settlement of green mussel Perna viridis post-larvae [71]. However, a hatchery study carried out on Australian blue mussels (Mytilus spp.) reported that optimal conditions for settlement may be achieved if other manipulations (food availability, airflow and type of substratum) are associated with stocking density [72].
Post-larvae production (Pr) has been reported as correlated to hatchery productivity in the Pacific oyster (Crassostrea gigas) [73]. In the present study, the significant relationship between larvae stocking density and post-larvae production indicates that tuning the former should not only consider settlement success but also yield. It is worth noting that the lowest pediveliger density (DL = 35 larvae/cm2) which showed the best settlement rate (>99%) had a lower post-larvae production (Pr = 35 post-larvae/cm2) than the medium pediveliger density, i.e., DM1 = 70 larvae/cm2 and 50 post-larvae/cm2. Therefore, a stocking density that assures the highest larvae settlement does not assure the highest post-larvae production. Stocking at moderate densities between 70 and 100 larvae/cm2 is, in this case, the threshold beyond which enhanced post-larvae production was not attainable.
A negative correlation has been reported between density and size, growth rate and weight gain in cultured Cortez oyster (Crassostrea corteziensis) [48], in silver-lip pearl oyster juveniles (Pinctada maxima) [50] and in fishes, such as the dover sole (Solea solea) [74] or the seabream (Sparus aurata) [75]. It is worth noting that the same result appears masked in the current experiment, i.e., the positive correlation observed between the increase in initial pediveliger density (DH1 + DH2 = DF1) and the gain in shell length (Gr, µm/day) is counterintuitive. It indeed suggests that the higher the initial pediveliger density is, the lower both the settlement and production rate are, but the higher the growth rate of the surviving post-larvae becomes. The high growth rates achieved by post-larvae in the current study (e.g., DF1 = 20–26 post-larvae/cm2; Gr = 22.24 ± 4.60 µm/day) may also be caused by the downwelling system enforced. The flow velocity implemented can enhance the filtration activity of post-larvae and therefore their phytoplankton intake as previously reported in green-lipped mussel (Perna canaliculus) [61,76]. Despite the flow velocity also benefiting other density groups, the significantly slowed growth at high post-larvae density (DF3 = 50 post-larvae/cm2; Gr = 19.23 ± 3.75 µm/day) may be due to the digestive interference of metabolic waste, as shown in post-larval growth of the taquilla clam (Mulinia edulis) [77]. Nevertheless, this waste accumulation hypothesis is unlikely herein due to the water quality maintenance applied throughout this study, e.g., see the efficiency of recirculating aquaculture systems (RASs) to achieve optimized survival and biomass on the “tambaqui” (Colossoma macropomum) [78]. Therefore, we refer to overcrowding and subsequent aggregation as the potential factors slowing down mussel post-larvae growth as reported in the pearl oyster (Pinctada fucata) [79].
In this experiment, we have seen that post-larvae density is negatively correlated with growth rate. This relationship produced a growth variability between treatments but also a post-larvae growth heterogeneity between batches (head, modal, tail) within density treatments (DF1, DF2, DF3). It is worth noting that the lower the post-larvae density, the lower the inter-batch heterogeneity in shell length, e.g., DF1 = (48:46:6)% for the (head/modal/tail) batches, respectively. Heterogeneity is a byproduct of differential growth among individuals, which increases the size variance, and it is known as growth depensation [80]. This phenomenon is currently sorted out in aquaculture practice by size grading, implying higher labor costs and culture slots [81]. Therefore, the lower the heterogeneity in favor of shell length, the better the productivity would be in terms of reducing fattening time [24], e.g., the highest-density treatments (DH1 + DH2 = DF1) produced a 48% share of post-larvae reaching the fastest development time but only a 6% share of post-larvae with delayed growth. By contrast, the higher the heterogeneity within density treatment, the more dispersed was the shell length between size batches DF2 and DF3, requiring higher management costs and fattening time. As also reported in cupped oysters (Crassostrea gigas), a low stocking density assures a more homogeneous batch growth [82].
The usual mussel seed transfer to nursery systems around 1 mm in shell length increases the risk of seed mortality [83]. A head start with pediveliger larvae (average shell length of 260 µm), to spat with a shell length >1.3 mm in micro-nursery systems as obtained here in 6 weeks can produce an outstanding mussel culture performance. Specifically, the low post-larvae density treatment (DF1) resulted in a seed stock >1.4 mm in shell length average prior to their transfer to nursery systems when juveniles are stronger and more resilient to manipulation.

5. Conclusions

The development of cost-effective protocols to produce high-quality mussel seed in hatchery systems requires the optimization of early rearing systems to assure viability. This work stresses the relevance of enforcing a pediveliger stocking density upon the production goal pursued. We show that an initial low density of larvae significantly enhanced the settlement rate (Sr = 70–90)% and post-larvae production (Pr = 35–50 post-larvae/cm2). However, high initial larvae densities (140–210 larvae/cm2) resulted in low Sr and Pr. Nevertheless, those low-density post-larvae cultures showed the highest growth rate among treatments (Gr = 22.24 ± 4.60 µm/day) and a size heterogeneity, i.e., 48% of post-larvae exhibiting the fastest development (head batch) and only 6% with a delayed growth (tail batch). The profitability of hatchery mussel production for enhancement and commercial production in the Alboran Sea of northern Morocco can be improved with the optimized stocking density protocols used in this study. Noteworthy, the resilience and consistency of our results are limited to the set of parameters established during this study (RAS, physicochemical environment, etc.). Therefore, future avenues related to the rearing system, hydrodynamics, type of settlement substratum, genetic improvement and probiotics are firm candidates to advance large-scale mussel production in captivity.

Author Contributions

Conceptualization, experimentation, data analyses and first draft, H.J. and Y.O.; data curation, A.A.; interpretation of results and editing, H.J. and P.P.; funding acquisition and project administration, P.P. All authors have read and agreed to the published version of the manuscript.

Funding

This study has been developed at the Amsa Shellfish Research Station, National Institute of Fisheries Research. The Moroccan Ministry of Higher Education, Scientific Research, and Innovation supported H.J. with a CNRST (Centre National pour la Recherche Scientifique et Technique) Excellence Research Scholarship grant (#19UAE 2022 (2022–2024). P.P. was supported by EU-Erasmus+ action KA131—Call 2021—Learning Mobility of Individuals—UVIGO Staff mobility for teaching STA to Université Abdelmalek Essaadi and training activities STT to AMSA Aquaculture Station (Tetouan, Morocco). This article APC was sponsored by project SEASOS as funded by program FORCYT (OEI–UE 2021–2023).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

All data generated are contained either in the article or extendable upon request to the Corresponding Author.

Acknowledgments

The authors are indebted to the Polydisciplinary Faculty of Larache (Abdelmalek Essaadi University—Tetouan) and the University of Vigo (ORI and OPI) for the academic support, to the Amsa Shellfish Research Station (National Institute of Fisheries Research INRH, Tetouan) for the technical facilities offered to carry on the experiments, and to their institutional staff for technical assistance.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Smaal, A.C.; Ferreira, J.G.; Grant, J.; Petersen, J.K.; Strand, Ø. Provisioning of Mussel Seed and Its Efficient Use in Culture. In Goods and Services of Marine Bivalves, 1st ed.; Smaal, A.C., Ferreira, J.G., Grant, J., Petersen, J.K., Strand, Ø., Eds.; Springer: Cham, Switzerland, 2019; pp. 27–50. [Google Scholar] [CrossRef]
  2. Maynard, D.J.; Trial, J.G. The use of hatchery technology for the conservation of Pacific and Atlantic salmon. Rev. Fish Biol. Fish. 2014, 24, 803–817. [Google Scholar] [CrossRef]
  3. Naish, K.A.; Taylor, J.E.; Levin, P.S.; Quinn, T.P.; Winton, J.R.; Huppert, D.; Hilborn, R. An Evaluation of the Effects of Conservation and Fishery Enhancement Hatcheries on Wild Populations of Salmon. Adv. Mar. Biol. 2007, 53, 61–194. [Google Scholar] [CrossRef] [PubMed]
  4. Valin, H.; Sands, R.D.; Van der Mensbrugghe, D.; Nelson, G.C.; Ahammad, H.; Blanc, E.; Bodirsky, B.; Fujimori, S.; Hasegawa, T.; Havlik, P.; et al. The future of food demand: Understanding differences in global economic models. J. Agric. Econ. 2014, 45, 51–67. [Google Scholar] [CrossRef]
  5. Nascimento-Schulze, J.C.; Bean, T.P.; Houston, R.D.; Santos, E.M.; Sanders, M.B.; Lewis, C.; Ellis, R.P. Optimizing hatchery practices for genetic improvement of marine bivalves. Rev. Aquac. 2021, 13, 2289–2304. [Google Scholar] [CrossRef]
  6. Galley, T.H.; Batista, F.M.; Braithwaite, R.; King, J.; Beaumont, A.R. Optimisation of larval culture of the mussel Mytilus edulis (L.). Aquac. Int. 2010, 18, 315–325. [Google Scholar] [CrossRef]
  7. Figueras, A. Biología y Cultivo del Mejillón (Mytilus galloprovincialis) en Galicia; CSIC: Madrid, Spain, 2007; p. 282. ISBN 9788400085261. [Google Scholar]
  8. Rothschild, B.J.; Ault, J.S.; Goulletquer, P.; Héral, M. Decline of the Chesapeake Bay oyster population: A century of habitat destruction and overfishing. Mar. Ecol. Prog. Ser. 2023, 111, 29–39. [Google Scholar] [CrossRef]
  9. Ouagajjou, Y.; Aghzar, A.; Presa, P. Population Genetic Divergence among Worldwide Gene Pools of the Mediterranean Mussel Mytilus galloprovincialis. Animals 2023, 13, 3754. [Google Scholar] [CrossRef]
  10. Gosling, E. Genetics of Mytilus; Elsevier: Amsterdam, The Netherlands, 1992; pp. 309–382. [Google Scholar]
  11. Hilbish, T.J.; Mullinax, A.; Dolven, S.I.; Meyer, A.; Koehn, R.K.; Rawson, P.D. Origin of the antitropical distribution pattern in marine mussels (Mytilus spp.): Routes and timing of transequatorial migration. Mar. Biol. 2000, 136, 69–77. [Google Scholar] [CrossRef]
  12. Lichtfouse, E.; Schwarzbauer, J.; Robert, D. Environmental Chemistry for a Sustainable World: Remediation of Air and Water Pollution, 1st ed.; Springer: Dordrecht, The Netherlands, 2011; p. 541. [Google Scholar]
  13. Wonham, M.J. Mini-review: Distribution of the Mediterranean mussel, Mytilus galloprovincialis (Bivalvia: Mytilidae), and hybrids in the northeast Pacific. J. Shellfish Res. 2023, 23, 535–544. [Google Scholar] [CrossRef]
  14. Comesaña, A.S.; Posada, D.; Sanjuan, A. Mytilus galloprovincialis Lmk. in northern Africa. J. Exp. Mar. Biol. Ecol. 1998, 223, 271–283. [Google Scholar] [CrossRef]
  15. Jaziri, H.; Benazzou, T. Différenciation allozymique multilocus des populations de moule Mytilus galloprovincialis Lmk. des côtes marocaines. Comptes Rendus Biol. 2002, 325, 1175–1183. [Google Scholar] [CrossRef] [PubMed]
  16. Ouagajjou, Y.; Presa, P. The connectivity of Mytilus galloprovincialis in northern Morocco: A gene flow crossroads between continents. Estuar. Coast Shelf Sci. 2015, 152, 1–10. [Google Scholar] [CrossRef]
  17. Idhalla, M.; Nhhala, H.; Kassila, J.; Ait Chattou, E.M.; Orbi, A.; Moukrim, A. Comparative production of two mussel species (Perna perna and Mytilus galloprovincialis) reared on an offshore submerged longline system in Agadir, Morocco. Int. J. Sci. Eng. Res. 2017, 8, 1203–1213. [Google Scholar] [CrossRef]
  18. Ouagajjou, Y.; Aghzar, A.; Miñambres, M.; Presa, P.; Pérez, M. Differential gene flow between populations of Mytilus galloprovincialis distributed along Iberian and north African coasts. Thalassas 2010, 26, 75–78. [Google Scholar]
  19. Abada-Boudjema, Y.M.; Dauvin, J.C. Recruitment and life span of two natural mussel populations Perna perna (Linnaeus) and Mytilus galloprovincialis (Lamarck) from the Algerian coast. J. Molluscan Stud. 1995, 61, 467–481. [Google Scholar] [CrossRef]
  20. Maanan, M. Biomonitoring of heavy metals using Mytilus galloprovincialis in Safi coastal waters, Morocco. Environ. Toxicol. 2007, 22, 525–531. [Google Scholar] [CrossRef]
  21. Azizi, G.; Layachi, M.; Akodad, M.; Yáñez-Ruiz, D.R.; Martín-García, A.I.; Baghour, M.; Mesfioui, A.; Skalli, A.; Moumen, A. Seasonal variations of heavy metals content in mussels (Mytilus galloprovincialis) from Cala Iris offshore (Northern Morocco). Mar. Pollut. Bull. 2018, 137, 88–694. [Google Scholar] [CrossRef]
  22. Ben Haddouch, A.; Amanhi, R.; Amzil, Z.; Taleb, H.; Rovillon, G.A.; Adly, F.; Loutfi, M. Lipophilic Toxin Profile in Mytilus galloprovincialis from the North Atlantic Coast of Morocco: LC-MS/MS and Mouse Bioassay Analyses. Int. J. Sci. Res. 2017, 6, 186. [Google Scholar] [CrossRef]
  23. Kouali, H.; Chaouti, A.; Achtak, H.; Elkalay, K.; Dahbi, A. Trace metal contents in the mussel Mytilus galloprovincialis from Atlantic coastal areas in northwestern Morocco: Levels of contamination and assessment of potential risks to human health. Mar. Pollut. Bull. 2022, 179, 113680. [Google Scholar] [CrossRef]
  24. Aghzar, A.; Talbaoui, M.; Benajiba, M.H.; Presa, P. Influence of depth and diameter of rope collectors on settlement density of Mytilus galloprovincialis spat in Baie de M’diq (Alboran Sea). Mar Freshw. Behav. Physiol. 2012, 45, 51–61. [Google Scholar] [CrossRef]
  25. Ait Chattou, E.M.; Abounahel, N.; Kassila, J.; Ouagajjou, Y.; Abouhala, A.; Idhalla, M.; Moukrim, A. Differential growth of the brown mussel, Perna perna (Linnaeus, 1758), in longline and pole cultures in Dakhla Bay (SW Morocco, Atlantic Ocean). Aquac. Res. 2019, 50, 736–747. [Google Scholar] [CrossRef]
  26. Fearman, J.A.; Bolch, C.J.; Moltschaniwskyj, N.A. Energy Storage and Reproduction in Mussels, Mytilus galloprovincialis: The Influence of Diet Quality. J. Shellfish Res. 2009, 28, 305–312. [Google Scholar] [CrossRef]
  27. Drissou, H.; Ouagajjou, Y.; Aghzar, A. Ecophysiology of the Mediterranean mussel Mytilus galloprovincialis L; effect of different microalgae diets and ration on broodstock conditioning. In Proceedings of the 1st International Congress on Coastal Research, Al Hoceima, Morocco, 6–9 July 2021. [Google Scholar] [CrossRef]
  28. Aghzar, A.; Talbaoui, M.; Benajiba, M.H.; Presa, P. Small-fast growers of Mytilus galloprovincialis do not catch up: An experimental test with size-graded mussels cultured in longline suspended bags. Mar. Freshw. Behav. Physiol. 2012, 45, 223–234. [Google Scholar] [CrossRef]
  29. Martínez-Pita, I.; Sánchez-Lazo, C.; Herrera, M. A non-lethal method for establishing sexual maturation in mussels (Mytilus galloprovincialis (Lamarck, 1819)) during broodstock conditioning in hatcheries. Aquac. Int. 2016, 24, 1247–1254. [Google Scholar] [CrossRef]
  30. Díaz-Puente, B.; Miñambres, M.; Rosón, G.; Aghzar, A.; Presa, P. Genetic decoupling of spat origin from hatchery to harvest of Mytilus galloprovincialis cultured in suspension. Aquaculture 2016, 460, 124–135. [Google Scholar] [CrossRef]
  31. Pronker, A.E.; Nevejan, N.M.; Peene, F.; Geijsen, P.; Sorgeloos, P. Hatchery broodstock conditioning of the blue mussel Mytilus edulis (Linnaeus 1758). Part I. Impact of different micro-algae mixtures on broodstock performance. Aquacult. Int. 2008, 16, 297–307. [Google Scholar] [CrossRef]
  32. Kamermans, P.; Galley, T.; Boudry, P.; Fuentes, J.; McCombie, H.; Batista, F.M.; Blanco, A.; Dominguez, L.; Cornette, F.; Pincot, L.; et al. Blue mussel hatchery technology in Europe. In Advances in Aquaculture Hatchery Technology, 1st ed.; Allan, G., Burnell, G., Eds.; Woodhead Publishing: Sawston, UK, 2013; pp. 339–373. [Google Scholar] [CrossRef]
  33. Aghzar, A.; Miñambres, M.; Alvarez, P.; Presa, P. A cost-benefit assessment of two multi-species algae diets for juveniles of Mytilus galloprovincialis. Int. J. Mar. Sci. 2013, 29, 9–16. [Google Scholar]
  34. Martínez-Pita, I.; Sánchez-Lazo, C.; Ruíz-Jarabo, I.; Herrera, M.; Mancera, J.M. Biochemical composition, lipid classes, fatty acids, and sexual hormones in the mussel Mytilus galloprovincialis from cultivated populations in south Spain. Aquaculture 2012, 358–359, 274–283. [Google Scholar] [CrossRef]
  35. El Moussaoui, M.A.; Ouagajjou, Y.; Aghzar, A.; Saoud, Y.; Nhhala, H. Feeding and growth coupling during different development larvae phases of the Mediterranean mussel Mytilus galloprovincialis L. from Amsa Bay. In Proceedings of the 1st International Congress on Coastal Research, Al Hoceima, Morocco, 6–9 July 2021. [Google Scholar] [CrossRef]
  36. Sánchez-Lazo, C.; Martínez-Pita, I. Effects of different mono, bi and trispecific microalgal diets on survival, growth, development, settlement, and fatty acid composition of mussel Mytilus galloprovincialis (Lamarck, 1819) larvae. Aquaculture 2014, 426–427, 138–147. [Google Scholar] [CrossRef]
  37. Pettersen, A.K.; Turchini, G.M.; Jahangard, S.; Ingram, B.A.; Sherman, C.D. Effects of different dietary microalgae on survival, growth, settlement, and fatty acid composition of blue mussel (Mytilus galloprovincialis) larvae. Aquaculture 2010, 309, 115–124. [Google Scholar] [CrossRef]
  38. Bordignon, F.; Trocino, A.; Rossetti, E.; Zomeno, C.; Pascual, A.; Birolo, M.; Martinez-Llorens, S.; Xiccato, G. Effect of stocking density on growth and survival of juvenile Manila clams (Ruditapes philippinarum) farmed in suspended lanterns in a North Italian lagoon. Aquac. Rep. 2021, 20, 100719. [Google Scholar] [CrossRef]
  39. Liu, Y.; Catalano, S.R.; Qin, J.; Han, J.; Zhan, X.; Li, X. Effects of cryopreservation on redox status and gene expression of trochophore larvae in Mytilus galloprovincialis. J. World. Aquac. Soc. 2022, 53, 516–526. [Google Scholar] [CrossRef]
  40. Anguiano-Beltrán, C.; Lizárraga-Partida, M.L.; Searcy-Bernal, R. Effect of Vibrio alginolyticus on larval survival of the blue mussel Mytilus galloprovincialis. Dis. Aquat. Organ. 2004, 59, 119–123. [Google Scholar] [CrossRef] [PubMed]
  41. Kurihara, H.; Asai, T.; Kato, S.; Ishimatsu, A. Effects of elevated pCO2 on early development in the mussel Mytilus galloprovincialis. Aquat. Biol. 2008, 4, 225–233. [Google Scholar] [CrossRef]
  42. Joyce, A.; Vogeler, S. Molluscan bivalve settlement and metamorphosis: Neuroendocrine inducers and morphogenetic responses. Aquaculture 2018, 487, 64–82. [Google Scholar] [CrossRef]
  43. Satuito, C.G.; Natoyama, K.; Yamazaki, M.; Fusetani, N. Inductin of Attachment and Metamorphosis of Laboratory Cultures Mussel Mytilus edulis galloprovincialis Larvae by Microbial Film. Fish. Sci. Res. 1995, 61, 223–227. [Google Scholar] [CrossRef]
  44. Tamburri, M.N.; Zimmer-Faust, R.K.; Tamplin, M.L. Natural Sources and Properties of Chemical Inducers Mediating Settlement of Oyster Larvae: A Re-examination. Biol. Bull. 1992, 183, 327–338. [Google Scholar] [CrossRef]
  45. Nevejan, N.; Davis, J.; Little, K.; Kiliona, A. Use of a formulated diet for mussel spat Mytilus galloprovincialis (Lamarck 1819) in a commercial hatchery. J. Shellfish Res. 2007, 26, 357–363. [Google Scholar] [CrossRef]
  46. Yang, J.L.; Satuito, C.G.; Bao, W.Y.; Kitamura, H. Larval settlement, and metamorphosis of the mussel Mytilus galloprovincialis on different macroalgae. Mar. Biol. 2007, 152, 1121–1132. [Google Scholar] [CrossRef]
  47. Crisp, D.J. Factors influencing the settlement of marine invertebrate larvae. In Chemoreception in Marine Organisms; Academic Press: London, UK, 1974. [Google Scholar]
  48. Mazón-Suástegui, J.M.; Ruíz-Ruíz, K.M.; Parres-Haro, A.; Saucedo, P.E. Combined effects of diet and stocking density on growth and biochemical composition of spat of the Cortez oyster Crassostrea corteziensis at the hatchery. Aquaculture 2008, 284, 98–105. [Google Scholar] [CrossRef]
  49. Roland, W.G.; Albrecht, K.J. Production of Pacific oysters, Crassostrea gigas Thunberg, from wild-caught and hatchery-produced seed grown at several densities on oyster shells. Aquac. Res. 1990, 21, 31–38. [Google Scholar] [CrossRef]
  50. Taylor, J.J.; Rose, R.A.; Southgate, P.C.; Taylor, C.E. Effects of stocking density on growth and survival of early juvenile silver-lip pearl oysters, Pinctada maxima (Jameson), held in suspended nursery culture. Aquaculture 1997, 153, 41–49. [Google Scholar] [CrossRef]
  51. Liu, B.; Dong, B.; Tang, B.; Zhang, T.; Xiang, J. Effect of stocking density on growth, settlement, and survival of clam larvae, Meretrix meretrix. Aquaculture 2006, 258, 344–349. [Google Scholar] [CrossRef]
  52. Karayücel, S.; Çelik, M.Y.; Karayücel, İ.; Öztürk, R.; Eyüboğlu, B. Effects of stocking density on survival, growth, and biochemical composition of cultured mussels (Mytilus galloprovincialis, Lamarck 1819) from an offshore submerged longline system. Aquac. Res. 2015, 46, 1369–1383. [Google Scholar] [CrossRef]
  53. Bacalhau de Oliveira, I.; Lavander, H.D.; Lima, P.; Barbosa de Oliveira, C.Y.; Macedo Dantas, D.M.; Olivera-Gálvez, A. Effect of stocking density on the growth and survival of Anomalocardia brasiliana (Gmelin, 1791) (Bivalvia: Veneridae) post-larvae. Ciência Rural 2019, 49, e20190420. [Google Scholar] [CrossRef]
  54. Lagos, L.; Herrera, M.; Sánchez-Lazo, C.; Martínez-Pita, I. Effect of larval stocking density on growth, survival, and whole body cortisol of the Mediterranean mussel Mytilus galloprovincialis (Lamarck, 1819) larvae reared under laboratory conditions. Aquac. Res. 2015, 46, 1648–1656. [Google Scholar] [CrossRef]
  55. Welch, B.L. The generalization of ‘STUDENT’S’ problem when several different population variances are involved. Biometrika 1947, 34, 28–35. [Google Scholar] [CrossRef] [PubMed]
  56. RStudio. Integrated Development for R. RStudio, Inc., Boston, MA, USA (Computer Software v0.98.1074). Available online: http://www.rstudio.com/ (accessed on 14 February 2022).
  57. Kamermans, P.; Blanco, A.; Joaquim, S.; Matias, D.; Magnesen, T.; Nicolas, J.L.; Petten, B.; Robert, R. Recirculation nursery systems for bivalves. Aquac. Int. 2016, 24, 827–842. [Google Scholar] [CrossRef]
  58. De Pauw, N. Use and production of microalgae as food for nursery bivalves. In Nursery Culturing of Bivalve Molluscs: Proceedings of the International Workshop on Nursery Culturing of Bivalve Molluscs Ghent, Belgium, 24–26 February 1981; Special Publication European Mariculture Society: Bredene, Belgium, 1981; Volume 7, pp. 35–69. [Google Scholar]
  59. Rodhouse, P.G.; O’kelly, M. Flow requirements of the oysters Ostrea edulis L. and Crassostrea gigas Thunb. in an upwelling column system of culture. Aquaculture 1981, 22, 1–10. [Google Scholar] [CrossRef]
  60. Laing, I.; Millican, P.F. Indoor nursery cultivation of juvenile bivalve molluscs using diets of dried algae. Aquaculture 1992, 102, 231–243. [Google Scholar] [CrossRef]
  61. Sanjayasari, D.; Jeffs, A. Optimising environmental conditions for nursery culture of juvenile GreenshellTM mussels (Perna canaliculus). Aquaculture 2019, 512, 734338. [Google Scholar] [CrossRef]
  62. Coleman, S.; Morse, D.; Brayden, W.C.; Brady, D.C. Developing a bioeconomic framework for scallop culture optimization and product development. Aquac. Econ. Manag. 2023, 27, 25–49. [Google Scholar] [CrossRef]
  63. Turini, C.S.; Sühnel, S.; Lagreze-Squella, F.J.; Ferreira, J.F.; Melo, C.M.R.D. Effects of stocking-density in flow-through system on the mussel Perna perna larval survival. Acta Sci. 2014, 36, 247–252. [Google Scholar] [CrossRef]
  64. Zhang, M.; Gao, X.; Lyu, M.; Lin, S.; Luo, X.; You, W.; Ke, C. Effect of stocking density on the growth and settlement of Haliotis discus hannai larvae in a recirculating aquaculture system. Aquac. Res. 2022, 53, 1468–1480. [Google Scholar] [CrossRef]
  65. Kobak, J. Light, gravity, and conspecifics as cues to site selection and attachment behaviour of juvenile and adult Dreissena polymorpha Pallas, 1771. J. Molluscan Stud. 2001, 67, 183–189. [Google Scholar] [CrossRef]
  66. Toonen, R.J.; Pawlik, J.R. Foundations of gregariousness. Nature 1994, 370, 511–512. [Google Scholar] [CrossRef]
  67. Alagarswami, K.; Dharmaraj, S.; Chellam, A.; Velayudhan, T.S. Larval and juvenile rearing of black-lip pearl oyster, Pinctada margaritifera (Linnaeus). Aquaculture 1989, 76, 43–56. [Google Scholar] [CrossRef]
  68. García-Lavandeira, M.; Silva, A.; Abad, M.; Pazos, A.J.; Sánchez, J.L.; Pérez-Parallé, M.L. Effects of GABA and epinephrine on the settlement and metamorphosis of the larvae of four species of bivalve molluscs. J. Exp. Mar. Biol. Ecol. 2005, 316, 149–156. [Google Scholar] [CrossRef]
  69. Crimaldi, J.P.; Thompson, J.K.; Rosman, J.H.; Lowe, R.J.; Koseff, J.R. Hydrodynamics of larval settlement: The influence of turbulent stress events at potential recruitment sites. Limnol. Oceanogr. 2002, 47, 1137–1151. [Google Scholar] [CrossRef]
  70. Manoj Nair, R.; Appukuttan, K.K. Effect of temperature on the development, growth, survival, and settlement of green mussel Perna viridis (Linnaeus, 1758). Aquac. Res. 2003, 34, 1037–1045. [Google Scholar] [CrossRef]
  71. Laxmilatha, P.; Rao, G.S.; Patnaik, P.; Rao, T.N.; Rao, M.P.; Dash, B. Potential for the hatchery production of spat of the green mussel Perna viridis Linnaeus (1758). Aquaculture 2011, 312, 88–94. [Google Scholar] [CrossRef]
  72. Weston, K.; Jahangard, S.; Ingram, B.A.; Miller, A.D.; Jennings, G.; Sherman, C.D. Factors affecting settlement, growth, and metamorphosis of hatchery-produced Australian blue mussel larvae. Aquacult. Int. 2021, 29, 1963–1977. [Google Scholar] [CrossRef]
  73. Dégremont, L.; Bédier, E.; Soletchnik, P.; Ropert, M.; Huvet, A.; Moal, J.; Samain, J.F.; Boudry, P. Relative importance of family, site, and field placement timing on survival, growth, and yield of hatchery-produced Pacific oyster spat (Crassostrea gigas). Aquaculture 2005, 249, 213–229. [Google Scholar] [CrossRef]
  74. Schram, E.; Van der Heul, J.W.; Kamstra, A.; Verdegem, M.C.J. Stocking density-dependent growth of Dover sole (Solea solea). Aquaculture 2006, 252, 339–347. [Google Scholar] [CrossRef]
  75. Canario, A.V.; Condeca, J.; Power, D.M.; Ingleton, P.M. The effect of stocking density on growth in the gilthead seabream, Sparus aurata (L.). Aquac. Res. 1998, 29, 177–181. [Google Scholar] [CrossRef]
  76. Alfaro, A.C. Effect of water flow and oxygen concentration on early settlement of the New Zealand green-lipped mussel, Perna canaliculus. Aquaculture 2005, 246, 285–294. [Google Scholar] [CrossRef]
  77. Oliva, D.; Abarca, A.; Gutiérrez, R.; Celis, Á.; Herrera, L.; Pizarro, V. Effect of stocking density and diet on growth and survival of post-larvae of the taquilla clam Mulinia edulis cultivated in sand in a hatchery. Rev. Biol. Mar. Oceanogr. 2013, 48, 37–44. [Google Scholar] [CrossRef]
  78. Santos, F.A.; Boaventura, T.P.; da Costa Julio, G.S.; Cortezzi, P.P.; Figueiredo, L.G.; Favero, G.; Palheta, G.; Melo, N.; Luz, R.K. Growth performance and physiological parameters of Colossoma macropomum in a recirculating aquaculture system (RAS): Importance of stocking density and classification. Aquaculture 2021, 534, 736274. [Google Scholar] [CrossRef]
  79. Gervis, M.H.; Sims, N.A. The Biology and Culture of Pearl Oysters (Bivalvia pteriidae); Overseas Development Administration and International Center for Living Aquatic Resources Management: London, UK; Manila, Philippines, 1992; p. 49. [Google Scholar]
  80. Magnuson, J.J. An analysis of aggressive behavior, growth, and competition for food and space in medaka (Oryzias latipes (Pisces, Cyprinodontidae)). Can. J. Zool. 2011, 40, 313–363. [Google Scholar] [CrossRef]
  81. Wilson, J. Hatchery rearing of Ostrea edulis and Crassostrea gigas. Aquac. Tech. Bull. 1981, 4, 1–34. [Google Scholar]
  82. Roncarati, A.; Felici, A.; Magi, G.E.; Bilandžić, N.; Melotti, P. Growth and survival of cupped oysters (Crassostrea gigas) during nursery and pregrowing stages in open sea facilities using different stocking densities. Aquac. Int. 2017, 25, 1777–1785. [Google Scholar] [CrossRef]
  83. Gui, Y.; Kaspar, H.F.; Zamora, L.N.; Dunphy, B.J.; Jeffs, A.G. Capture efficiency of artificial food particles of post-settlement juveniles of the Greenshell™ mussel, Perna canaliculus. Aquaculture 2016, 464, 1–7. [Google Scholar] [CrossRef]
Animals 14 00239 g001
Figure 1. Experimental design and row results on larvae and post-larvae density testing in Mytilus galloprovincialis. DL, DM1, DM2, DH1, DH2, initial pediveliger stocking densities; reps, no. of replicates per density treatment; DF1, DF2, DF3, final post-larvae densities.
Figure 1. Experimental design and row results on larvae and post-larvae density testing in Mytilus galloprovincialis. DL, DM1, DM2, DH1, DH2, initial pediveliger stocking densities; reps, no. of replicates per density treatment; DF1, DF2, DF3, final post-larvae densities.
Animals 14 00239 g001
Animals 14 00239 g002
Figure 2. (a) Specific replicated structure implemented to monitor metamorphosis and settlement of mussel post-larvae; (b) detail of the downwelling system designed for rearing and tracking the larvae–post-larvae transition.
Figure 2. (a) Specific replicated structure implemented to monitor metamorphosis and settlement of mussel post-larvae; (b) detail of the downwelling system designed for rearing and tracking the larvae–post-larvae transition.
Animals 14 00239 g002
Animals 14 00239 g003
Figure 3. (a) Water flow through recirculating pipes into a nylon-mesh cylinder; (b) post-larvae settled in a screened nylon-mesh cylinder.
Figure 3. (a) Water flow through recirculating pipes into a nylon-mesh cylinder; (b) post-larvae settled in a screened nylon-mesh cylinder.
Animals 14 00239 g003
Animals 14 00239 g004
Figure 4. (a) Settlement rate (Sr, %); (b) post-larvae production (Pr, post-larvae/cm2) at different larvae stocking densities: DL = 35 larvae/cm2; DM1 = 70 larvae/cm2; DM2 = 100 larvae/cm2; DH1 = 140 larvae/cm2; and DH2 = 210 larvae/cm2. A significant difference exists between treatments with different superscripts (a–d).
Figure 4. (a) Settlement rate (Sr, %); (b) post-larvae production (Pr, post-larvae/cm2) at different larvae stocking densities: DL = 35 larvae/cm2; DM1 = 70 larvae/cm2; DM2 = 100 larvae/cm2; DH1 = 140 larvae/cm2; and DH2 = 210 larvae/cm2. A significant difference exists between treatments with different superscripts (a–d).
Animals 14 00239 g004
Animals 14 00239 g005
Figure 5. Percentage of post-larvae shell length batches (head, blue; modal, yellow; tail, grey) from three final post-larvae densities (DF1 = 20–26 post-larvae/cm2, DF2 = 35–38 post-larvae/cm2 and DF3 = 50 post-larvae/cm2).
Figure 5. Percentage of post-larvae shell length batches (head, blue; modal, yellow; tail, grey) from three final post-larvae densities (DF1 = 20–26 post-larvae/cm2, DF2 = 35–38 post-larvae/cm2 and DF3 = 50 post-larvae/cm2).
Animals 14 00239 g005
Table 1. Results of ANOVA (one factor) for the settlement rate at different stocking densities (DL = 35 larvae/cm2; DM1 = 70 larvae/cm2; DM2 = 100 larvae/cm2; DH1 = 140 larvae/cm2; and DH2 = 210 larvae/cm2). Sr, settlement rate; F = Fisher test; df_num = numerator degrees of freedom, df_den = denominator degrees of freedom, *** p < 0.001.
Table 1. Results of ANOVA (one factor) for the settlement rate at different stocking densities (DL = 35 larvae/cm2; DM1 = 70 larvae/cm2; DM2 = 100 larvae/cm2; DH1 = 140 larvae/cm2; and DH2 = 210 larvae/cm2). Sr, settlement rate; F = Fisher test; df_num = numerator degrees of freedom, df_den = denominator degrees of freedom, *** p < 0.001.
Initial Density (Larvae/cm2)Mean Sr (±SD) 1df_numdf_denFp-Value
DL0.99 ± 0.16 d4.008.9093.452.859 × 10−7 ***
DM10.71 ± 0.08 c
DM20.36 ± 0.05 b
DH10.18 ± 0.01 a
DH20.09 ± 0.02 a
1 A significant difference exists between treatments with different superscripts (a–d).
Table 2. ANOVA (one factor) for post-larvae production under different stocking densities (DL = 35 larvae/cm2; DM1 = 70 larvae/cm2; DM2 = 100 larvae/cm2; DH1 = 140 larvae/cm2; and DH2 = 210 larvae/cm2), F = Fisher test; df_num = numerator degrees of freedom, df_den = denominator degrees of freedom, *** p < 0.001.
Table 2. ANOVA (one factor) for post-larvae production under different stocking densities (DL = 35 larvae/cm2; DM1 = 70 larvae/cm2; DM2 = 100 larvae/cm2; DH1 = 140 larvae/cm2; and DH2 = 210 larvae/cm2), F = Fisher test; df_num = numerator degrees of freedom, df_den = denominator degrees of freedom, *** p < 0.001.
Initial Density (Larvae/cm2)Post-Larvae Production (Mean Pr ± SD) 1df_numdf_denFp-Value
DL35.25 ± 5.85 b49.0825.735.718 × 10−5 ***
DM150.20 ± 5.80 a
DM238.35 ± 5.93 b
26.00 ± 1.58 c
DH126.00 ± 1.58 c
DH219.20 ± 5.21 c
1 A significant difference exists between yields with different superscripts (a–c).
Table 3. Results of ANOVA (one factor) for post-larvae growth (shell length in µm/day) at different densities (DF1 = 20–26 post-larvae/cm2, DF2 = 35–38 post-larvae/cm2 and DF3 = 50 post-larvae/cm2). F = Fisher test; df_num = numerator degree of freedom, df_den = denominator degree of freedom *** p < 0.001.
Table 3. Results of ANOVA (one factor) for post-larvae growth (shell length in µm/day) at different densities (DF1 = 20–26 post-larvae/cm2, DF2 = 35–38 post-larvae/cm2 and DF3 = 50 post-larvae/cm2). F = Fisher test; df_num = numerator degree of freedom, df_den = denominator degree of freedom *** p < 0.001.
Density (Post-Larvae/cm2)Growth Rate (Mean Gr ± SD) 1df_numdf_denFp-Value
DF122.24 ± 4.60 a2 16.681.576 × 10−7 ***
DF220.73 ± 4.65 b250.97
DF319.23 ± 3.75 c
1 A significant difference exists between densities with different superscripts (a–c).
Table 4. Results of ANOVA (one factor) of the effect of post-larvae density on growth rate (Gr, µm/day) in the different batches (head, modal, tail). F = Fisher test; df_num = numerator degrees of freedom; df_den = denominator degrees of freedom; ns = not significant; ** p < 0.01, *** p < 0.001.
Table 4. Results of ANOVA (one factor) of the effect of post-larvae density on growth rate (Gr, µm/day) in the different batches (head, modal, tail). F = Fisher test; df_num = numerator degrees of freedom; df_den = denominator degrees of freedom; ns = not significant; ** p < 0.01, *** p < 0.001.
BatchDensity GroupGrowth (Gr ± SD) 1df_numdf_denFp-Value
TailDF112.76 ± 3.14 a220.066.722.25 × 10−3 **
DF214.68 ± 0.84 b
DF314.58 ± 1.14 b
ModalDF119.27 ± 1.66 a2105.850.200.82 ns
DF219.45 ± 1.86 a
DF319.38 ± 2.08 a
HeadDF126.40 ± 2.55 b289.4231.664.02 × 10−11 ***
DF226.24 ± 2.19 b
DF323.99 ± 0.97 a
1 Superscripts (a, b) indicate a significant difference between densities within batches.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Janah, H.; Aghzar, A.; Presa, P.; Ouagajjou, Y. Influence of Pediveliger Larvae Stocking Density on Settlement Efficiency and Seed Production in Captivity of Mytilus galloprovincialis in Amsa Bay, Tetouan. Animals 2024, 14, 239. https://doi.org/10.3390/ani14020239

AMA Style

Janah H, Aghzar A, Presa P, Ouagajjou Y. Influence of Pediveliger Larvae Stocking Density on Settlement Efficiency and Seed Production in Captivity of Mytilus galloprovincialis in Amsa Bay, Tetouan. Animals. 2024; 14(2):239. https://doi.org/10.3390/ani14020239

Chicago/Turabian Style

Janah, Hafsa, Adil Aghzar, Pablo Presa, and Yassine Ouagajjou. 2024. "Influence of Pediveliger Larvae Stocking Density on Settlement Efficiency and Seed Production in Captivity of Mytilus galloprovincialis in Amsa Bay, Tetouan" Animals 14, no. 2: 239. https://doi.org/10.3390/ani14020239

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop