# In vitro dynamics of oyster hemocytes in plasma-based primary cultures

Danielle Mello, Rafael Trevisan, Yasmine Even, Valentin Foulon, Christophe Lambert, Heloísa Gabe, Manon Le Goff, Claudie Quéré, Fernando Ramos Queiroga, Christophe Brigaudeau, et al.

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## To cite this version:

Danielle Mello, Rafael Trevisan, Yasmine Even, Valentin Foulon, Christophe Lambert, et al. In vitro dynamics of oyster hemocytes in plasma-based primary cultures. Experimental Cell Research, 2025, 450 (2), pp.114662. 10.1016/j.yexcr.2025.114662. hal-05465524

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Submitted on 20 Jan 2026

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Research article

*In vitro* dynamics of oyster hemocytes in plasma-based primary cultures

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Danielle F. Mello<sup>a, *, D</sup>, Rafael Trevisan<sup>a, 1</sup>, Yasmine Even<sup>a, 1</sup> <sup>D</sup>, Valentin Foulon<sup>b</sup>,

Christophe Lambert<sup>a</sup>, Heloísa B. Gabe<sup>a, c</sup>, Manon Le Goff<sup>a</sup>, Claudie Quéré<sup>a</sup>,

Fernando R. Queiroga<sup>a</sup>, Christophe Brigaudeau<sup>d, D</sup>, Hélène Talarmin<sup>e</sup>, Caroline Montagnani<sup>f</sup>,

Guillaume M. Charrière<sup>f</sup>, Delphine Destoumieux-Garzon<sup>f</sup>, Stéphanie Madec<sup>a, 2</sup>,

Charlotte Corporeau<sup>a, 2, D</sup>

ARTICLE INFO

Keywords:

Marine invertebrates
Marine cell biology
Immune cells
Immunometabolism
Primary cell culture
Live-cell imaging

A B S T R A C T

Hemocytes are specialized cells found in the circulatory system of invertebrates, such as marine bivalves, and play crucial roles in physiology and health. However, in bivalves, in-depth studies on their *in vitro* biology beyond one or two days remain scarce, particularly in *Crassostrea (Magallana) gigas*. In this study, we characterized the hemocytes isolated from triploid *C. gigas* oysters, maintained *in vitro* in their natural medium, hemolymph plasma, to assess their morphological, behavioral, and metabolic status over 21 days. A diverse population of hemocyte morphotypes, including granulocytes, hyalinocytes, and blast-like cells, was preserved for up to seven days, displaying various shapes and spreading patterns. By days 14 and 21, blast-like cells and hyalinocytes were less represented, showcasing granulocytes as the most resilient subpopulation *in vitro*. Despite this decline in subpopulation diversity, we confirmed through cell viability and metabolic assays, combined with live-cell imaging, that the remaining cells remained alive and active throughout the culture period. Notably, key indicators of cellular homeostasis such as membrane integrity, ATP levels, mitochondrial content, and mitochondrial reactive oxygen species (ROS) remained stable in these surviving cells. Phagocytic activity persisted over the full 21 days, with a marked increase after two weeks in culture. Certain time-dependent changes were observed, including a decrease in total basal ROS production from day 7 and a significant increase in neutral lipid storage by day 21. Live-cell imaging revealed two modes of cell motility: pseudopod-dependent and lamellipodium-dependent membrane extensions. Fibroblast-like hyalinocytes remained non-motile. Interestingly, not all motile granulocytes display phagocytic behavior and non-motile fibroblast-like hyalinocytes lack phagocytic activity entirely, highlighting the functional diversity within the different morphotypes identified in our cultures. By presenting an innovative method for sustaining primary marine invertebrate cell cultures using hemolymph plasma and leveraging different cell-based assays and live-cell imaging, this study provides a platform for investigating morphology, behavior, metabolism, and immune function of these cells under controlled *in vitro* conditions. Additionally, by highlighting the selective decline of certain hemocyte subpopulations, this study offers an *in vitro* model well-suited for future research into hemocyte lifespan and cell death mechanisms. Importantly, our framework not only advances research on bivalve hemocyte biology but also contributes to the broader field of experimental and comparative cell biology, with potential with applications in immunology, ecotoxicology, ecophysiology, molecular screening, and health innovation.

<footnote><sup>a</sup> Univ Brest, Ifremer, CNRS, IRD, LEMAR, IUEM, F-29280, Plouzane, France</footnote>

<footnote><sup>b</sup> ENIB, UMR CNRS 6285 LabSTICC, F-29238, Brest, France</footnote>

<footnote><sup>c</sup> Department of Biochemistry, Federal University of Santa Catarina, 88040-900 Florianópolis, Brazil</footnote>

<footnote><sup>d</sup> Univ Brest, INSERM, EFS, UMR 1078, GGB, F-29200, Brest, France</footnote>

<footnote><sup>e</sup> Univ Brest, ORPHY EA 4324, F-29200, Brest, France</footnote>

<footnote><sup>f</sup> IHPE, Univ Montpellier, CNRS, IFREMER, Univ Perpignan Via Domitia, Montpellier, France</footnote>

<footnote>* Corresponding author.
E-mail address: danielle.mello@univ-brest.fr (D.F. Mello).</footnote>

<footnote n="1">¹ These authors contributed equally to this work.</footnote>

<footnote n="2">² Co-last authors.</footnote>
# 1. Introduction

Hemocytes are specialized cells found in the circulatory system of invertebrates, such as marine bivalves, and play crucial roles in physiology and health. Hemocytes are immune cells acting as the first line of defense against pathogens, parasites, and foreign particles. The most important functions of hemocytes are the detection and engulfment of invading microorganisms through phagocytosis and the production of proteolytic and antimicrobial molecules, effectively neutralizing these threats and preventing infections. Hemocytes are also involved in encapsulation, a mechanism where they surround larger foreign bodies or parasites that cannot be readily phagocytosed. By encapsulating these threats, hemocytes prevent their spread and limit the damage caused to the tissues. Hemocytes also participate in wound healing and tissue repair when a bivalve sustains an injury: they migrate to the site of damage and assist in sealing the wound, facilitating the regeneration of damaged tissues. Beyond classical immune roles, hemocytes exhibit multifunctional characteristics akin to those of primitive cell systems. These cells play key roles in nutrient transport, osmotic regulation, shell mineralization, and the removal of metabolic waste from the hemolymph [1-3]. In many bivalve species, including oysters, despite ongoing debates regarding their classification, three main types of hemocytes are generally recognized: blast-like cells, hyalinocytes, and granulocytes [2]. Blast-like cells are thought to be hemocyte precursors based on their morphometric features and low organelle content, while granulocytes are typically considered the most immunocompetent cell type, generally presenting enhanced phagocytic activity [2]. Hyalinocytes, on the other hand, are believed to play a key role in hemocyte aggregation and wound healing processes [2], and a recent study has highlighted their involvement in the expression of antimicrobial peptides [4].

The study of hemocyte biology has emerged as a fundamental aspect of ecotoxicology and ecophysiology research in understanding the resilience and adaptability of marine bivalves in their natural environments and in the context of environmental changes. Measuring the functions of hemocytes (e.g., phagocytosis and viability) and their abundance (e.g., number of cells per volume of hemolymph) have long been used as key indicators of the overall health and physiological condition of bivalves [5]. Indeed, changes in hemocyte numbers, morphology, or function may reflect the exposure to pollutants, toxins, or environmental stressors, making them valuable biomarkers for assessing environmental quality and the impact of human activities on marine ecosystems [6-8]. Thus, research focusing on hemocyte biology is crucial for predicting and mitigating the impacts of a diversity of stressors like ocean acidification, pollution, and climate change on bivalve populations. This can support the development of strategies aiming at the conservation and management of bivalve populations and their coastal environments under current and future environmental conditions.

The Pacific oyster *Crassostrea (Magallana) gigas* (Thunberg, 1793) is a marine invertebrate that holds a major economic and ecological significance at the global scale. It is one of the most cultivated shellfish around the world and an abundant mollusk species in an increasing number of coastal areas. *C. gigas* is widely used as a sentinel bivalve species for biomonitoring studies in coastal marine environments [9] and to understand the impacts of environmental stressors [10]. In such studies, the effects on oyster health are often evaluated by assessing hemocyte parameters *ex vivo* [11–17]. However, typically, this approach requires large facilities, expensive experimental conditions, co-maintenance of microalgae cultures for animal feeding, and intensive hemolymph sampling on the days of experimentation.

*In vitro* studies provide numerous advantages for biological research. They are potent and valuable tools in marine bivalve research, offering crucial insights into the physiology, health, and responses of these organisms to various environmental stressors [18–22]. *In vitro* cell studies can provide the investigation of precise mechanisms (gene expression,

enzyme activities, biomarkers) linked to changes in cell function, morphology, and metabolism, shedding light on how individuals adapt their cellular responses to environmental challenges. This is particularly important for bivalves, who are constantly exposed to a variety of environmental stressors such as temperature, food, and oxygen fluctuations within the intertidal zone, as well as pathogens and pollutants [23–26]. Additionally, *in vitro* studies reduce ethical concerns associated with *in vivo* experiments, as they minimize the number of animals needed for large-scale experimentations. *In vivo* studies not only require a greater number of animals to be sacrificed but also, in the case of waterborne pollutant exposures, they generate significantly larger amounts of chemical waste.

Nonetheless, maintaining bivalve hemocytes viable and functional under *in vitro* conditions has been a major challenge [27]. Most studies in immunology and ecotoxicology research utilizing bivalve hemocytes have traditionally been restricted to investigating these cells *ex vivo* for a limited duration, typically ranging from a few hours to 2–3 days [2,28, 29]. Several attempts have been made to optimize cell culture media and extend the duration of hemocyte viability. The longest reported maintenance of viable bivalve hemocytes in culture was with scallop (*Chlamys farreri*), where at least 50% of the initial cells remained viable for up to 48 days [30]. A study on *Mytilus galloprovincialis* established primary hemocyte cultures with stable viability for up to 20 days [31], while another study reported the maintenance of primary hemocyte cultures from *Ruditapes decussatus* for up to 7 days [32]. More recent studies by Potts et al. [33] and Balakrishnan et al. [34] also described primary cultures of hemocytes from *C. gigas* and *C. madrasensis*, respectively; however, neither study specifies the duration for which the hemocyte cultures were maintained. In fact, while the studies cited above suggest that bivalve hemocytes can survive in culture for extended periods, none have provided a comprehensive characterization of cell types and their morphologies, visual documentation of culture dynamics over time, or detailed analyses of the cells' functional and metabolic properties throughout the culture period.

In this study, we hypothesized that *C. gigas* hemocytes maintained in their native plasma could retain cellular viability and functionality for extended periods. To test this, we conducted a comprehensive 21-day *in vitro* characterization, assessing hemocyte morphology, behavior, metabolism, and phagocytic activity. This work establishes a robust framework for long-term *in vitro* monitoring of hemocyte physiology and behavior under conditions that more closely mimic the *in vivo* environment, offering a valuable tool for advancing research in cellular biology, immunology, ecophysiology, and ecotoxicology of marine bivalves.

# 2. Methods

## 2.1. Animals

Hatchery-produced adult *C. gigas* (triploid, size n°2-86-110 g of whole-body weight) were obtained from an oyster farm (Beg Ar Vil) located at the Aber Benoit (France), an area considered to have a low anthropogenic impact [35]. The main reasons why choosing triploid oysters over the naturally occurring diploid ones are their easy access through the aquaculture industry, the possibility of obtaining hemolymph without contamination by gametes all year long, and the absence of major metabolic changes induced by reproductive maturation. Oysters were conditioned from 48 h up to 4 weeks before cell sampling. They were maintained either in natural seawater with natural food source at Ifremer's experimental site (Ste Anne, Plouzané, France) or under controlled conditions with seawater temperature at 16 ± 1.5 °C, constant aeration, water flow, and daily feeding by a commercial phytoplankton diet (0.5 mL per animal; SA Live Marine Phytoplankton®, Premium Blend) in the laboratory (Plouzané, France). Oysters maintained with natural seawater were used for the optimization experiments and the November 2022 sampling, whereas oysters
maintained under laboratory settings were used for the April and May 2023 samplings (see details in Section 2.2).

## 2.2. Hemolymph collection

Hemolymph was collected by performing a small notch in the shell immediately before sampling. A cold 1 mL syringe coupled with a 23G x 1" needle was injected into the adductor muscle, and hemolymph was gently withdrawn, avoiding bubbles and seawater contamination. The use of a cold syringe is essential to prevent hemocyte aggregation during hemolymph collection, effectively eliminating the need for an anticoagulant solution. A drop of each hemolymph sample collected was used to inspect hemolymph quality under a light microscope using both 4 × and 10 × objectives. Only samples containing zero visualization of contaminants (e.g., tissue and cellular debris and visible microorganisms: bacteria, protozoa, and microalgae) and no excessive hemocyte aggregation were used. Hemolymph samples that passed this rigorous quality check were pooled in 15 mL centrifuge tubes maintained on ice. Hemolymph collection usually lasted about 1 h but depending on the volume needed and easiness of collection, collection time varied between 1 and 3 h. Both the rigorous quality check of each hemolymph extraction and the limited time on ice (up to 3 h) were critical factors for the success of the primary cultures. Samplings for the three independent 21-day experiments were conducted in November 2022 (autumn), April 2023 (spring) and May 2023 (spring). In the first experiment (November 2022), two independent pools of hemolymph were analyzed (N = 2), while in the second and third experiments (April and May 2023), five independent pools of hemolymph were analyzed (N = 5 for each). Each hemolymph pool consisted of samples from 2 to 6 oysters.

## 2.3. Primary hemocyte cultures

The first step in preparing the primary hemocyte cultures was to determine the concentration of hemocytes in hemolymph pools. For this, 50 µL of each hemolymph pool was mixed with 50 µL of filtered sterile seawater (FSSW; 0.22 µm polyethersulfone -PES- syringe filter) containing 3.9 % formaldehyde to avoid cell clumping and adherence to the counting chamber, which are detrimental factors for accurate counting. Then 10 µL of the fixed hemolymph was added to the counting slides adapted to the EVE™ Automatic Cell Counter. Once the cell concentration was determined, whole hemolymph pool samples containing 500.000 or 80.000 cells were seeded into each well of 24 or 96-well plates (~250,000 cells/cm²), respectively. Plates were pre-coated with poly-D-lysine either from the manufacturer or at the laboratory following the manufacturer's protocol (Gibco™, ThermoFisher Scientific, A3890401; for microplate references, see Supp. Table 1). The microplates containing the freshly seeded cells were then incubated for at least 1 h (up to 3 h) at 16 ± 1 °C for adherence using a standard cooling incubator (Memmert IPP55; which uses ambient air, i.e., no O₂ or CO₂ control). The supernatant was then gently completely removed and replaced with the hemocyte culture media. The cell culture medium was composed of filtered-sterile oyster plasma (FSOP) containing penicillin (+P) 100 U/ml and streptomycin (+S) 100 µg/ml (Gibco™, ThermoFisher Scientific, 15140148). The FSOP medium predominantly consisted of two pools of whole hemolymph samples from about 100 oysters (each collected at two different periods, September 2022 and March 2023), which were filtered (0.22 µm PES syringe filter), split into aliquots, and stored without antibiotics at -80 °C until use at the start of the cultures or for complete medium renewal every 7 days. A final volume of 500 µL or 100 µL of the cell culture medium FSOP + P/S was used for the 24 or 96-well plates, respectively.

## 2.4. Characterization of cell morphotypes and behavior

In this study, we define "cell morphotype" as a recurring and distinguishable morphological pattern observed among the oyster

primary hemocyte cultures, reflecting either distinct subpopulations (granulocytes, hyalinocytes, blast-like cells) or stable shape variations within subpopulations under our *in vitro* conditions. We combined both May-Grünwald Giemsa staining with live-cell imaging to characterize the morphotypes present in the cultures over time. All assays described in this section were optimized to be conducted in cells cultured in 96-well microplates.

For the May-Grünwald Giemsa (MGG; RAL Diagnostics, 320070) staining, microplates were centrifuged (1000×g, 5 min, 16 °C), the supernatant was carefully removed, replaced with 100 µL of May-Grünwald and incubated for 3 min at room temperature. Then, 200 µL of distilled H₂O was added and samples were incubated for an extra 1 min. The supernatant was then removed, cells were rinsed once with 200 µL of distilled H₂O, and incubated with 100 µL of Giemsa R solution for 10 min. Finally, the supernatant was removed, and cells were rinsed with 200 µL of distilled H₂O. Plates were left to dry, and just before the microscope visualization, a drop of microscopy immersion oil was added to each well to improve cell visualization. Images were recorded using an inverted microscope (Zeiss AXIO Observer Z1) coupled with a color camera system (Canon EOS 700D).

Live unstained cells were imaged through phase-contrast microscopy (EVOS XL Core Imaging System, ThermoFisher Scientific) or Confocal Laser Scanning Microscopy (Zeiss LSM780, Carl Zeiss) using the transmitted-light channel equipped with a photomultiplier (TPMT). Phase-contrast and transmitted-light TPMT images were taken every 10s or 3s, using × 10 or × 40 air objectives, respectively.

The phagocytic behavior of hemocytes in primary cultures was performed by time-lapse imaging of cells incubated with Fluoresbrite® YG Microspheres beads (2 µm, Polysciences Inc.). The beads were added to the primary cultures at a final concentration of 0.005 %, and cultures were incubated for 4 h at 16 ± 1 °C before imaging. Image acquisition was performed every 10 s for 3 min at room temperature (20 ± 1 °C) using the transmitted-light channel of the Zeiss AXIO Observer Z1 coupled with the Canon EOS 700D color camera system.

Time-lapse videos were created with the MakeAVI software using 5 frames per second (each second of the video corresponds to 50 s in real-time), except for the videos using confocal transmitted-light TPMT images, which were created with the ZEN 2012 software (Carl Zeiss) using 10 frames per second (each second of the video corresponds to 30 s in real-time).

## 2.5. Cellular physiological parameters

Cellular parameters involving size, complexity, viability, immune function, and metabolism were analyzed on hemocytes maintained in primary cultures over time. Assays were performed through flow cytometry (cell viability, ROS production, phagocytosis, mitochondrial and neutral lipid contents) or by multimode plate readers (ATP content and NADH production) as described below. For these assays, the start of each primary cell culture was staggered, meaning that cells were seeded 21, 14, and 7 days before the day of the analysis and compared to fresh hemolymph samples (Day 0). The assays were thus performed on the same day so that all cell culture time points could be more readily comparable. Therefore, it is important to highlight that each time-point of the cell-based assays corresponded to cultures obtained from different hemolymph pools (thus different oysters).

Flow cytometry analyses were performed from primary hemocyte cultures maintained in 24-well plates (Supp. Table 1). Cells were recovered from the culture wells at Days 7, 14, and 21 by gently and repeatedly pipetting the medium within each well, then transferring the suspension to centrifuge microtubes kept on ice. In contrast, the Day 0 hemocytes (fresh hemolymph, used as control) were directly transferred to microtubes on ice without being plated, to avoid early cell adhesion and potential loss. This precaution was necessary because most hemocyte types tend to be more adherent to the bottom of the well within the first hours of culture and cannot be easily detached. Flow
cytometry assays were performed by transferring 50 µL of cells into flow cytometer tubes containing 150 µL of FSSW containing different fluorophores. Cell viability was measured by incubation with 10 µg/mL of propidium iodide (Sigma, France) and 1X SYBR Green I (10,000X stock; Sigma, France) for 30 min, at 18 °C, in the dark [36]. Cell size and complexity were quantified using the same samples stained for cell viability through measurements obtained by the side scatter (SSC), representing the granularity or internal complexity of particles, and the forward scatter (FSC), estimating the size of cells. SYBR Green I-stained hemocytes could be distinguished from other hemolymph particles by gating and differentiating them using a density plot visualization of SSC vs. FL1, displayed on a log scale [36]. For the quantification of ROS production, incubation was performed with 10 µM dichlorodihydrofluorescein diacetate (DCFH-DA; Sigma, France) for 60 min, at 18 °C, in the dark [37]. Hemocyte phagocytosis was measured using the final concentration of Fluoresbrite® YG Microspheres beads at 0.005 % (2 µm, Polysciences Inc., France) and incubation was performed for 120 min, at 18 °C, in the dark. The percentages of active phagocytes were defined as the percentage of cells that ingested three or more beads [38]. Mitochondrial content, mitochondrial ROS production, and neutral lipid content were measured according to the manufacturer's protocols. Mitochondrial content was assessed by incubating cells with a 1/1000 dilution of MITO-ID® Green (Enzo Life Sciences) for 1 h at 18 °C, in the dark. Mitochondrial ROS production was quantified using 5 µM MitoSOX Red (Thermo Fisher Scientific) and a 30 min incubation period at 18 °C, in the dark. Neutral lipids content was measured by incubating cells with 10 µM of BODIPY™ 493/503 (Thermo Fisher Scientific) for 15 min at 18 °C, in the dark. Cell viability, ROS production, and phagocytosis analyses were performed on a BD FACSVerse™ flow cytometer (BD Biosciences, San Jose, CA, USA). Mitochondrial content and ROS production and neutral lipid content analyses were performed on a Guava® easyCyte HT System (Luminex, Merck Millipore).

ATP production was measured in primary hemocyte cultures maintained in 96-well plates (Supp. Table 1) using the kit CellTiter-Glo™ 3D (Promega, G9681), following the manufacturer's protocol with some minor modifications. On the day of the assay, plates were centrifuged (1000×g, 5 min, 16 °C), and the supernatant was carefully removed and replaced with 50 µL of FSSW plus 50 µL of the ATP kit reagent. Plates were submitted to linear shaking for 2 min, then samples were transferred to opaque white 96-well plates (Supp. Table 1), incubated for 10 min at room temperature, in the dark, and luminescence was recorded using the EnSpire® Multimode Plate Reader (PerkinElmer®). ATP absolute values were calculated based on ATP standard curves (0, 1, 10, 100, 625, 1250, 2500 nM) performed on the same day and plate with the samples.

## 2.6. Statistical analyses

Statistical analyses of the cellular parameter results were conducted using one-way analysis of variance (ANOVA) followed by Tukey's post hoc test. Statistical significance was set at $p < 0.05$. All experiments were performed in three independent runs (N = 12; comprising 2 pools from the November 2022 experiment and 5 pools each from the April and May 2023 experiments), except for the neutral lipid content and mitochondrial ROS production assays, which were conducted in two independent runs (N = 10; 5 pools from the April and May 2023 experiments each).

# 3. Results

## 3.1. A subpopulation of hemocytes remains viable and metabolically active in plasma-based culture for at least 21 days

C. gigas hemocytes cultured in a plasma-based medium (FSOP) at 16 ± 1 °C, using poly-D-lysine-coated vessels and antibiotics (PS) to prevent bacterial contamination, sustained a subpopulation of viable cells for at

least 21 days (Fig. 1). Viability assessments showed that despite a significant decrease from day 0 to day 7, approximately 70 % of hemocytes remained viable up to 14 days in culture, declining to ~50 % by day 21, as measured by the Alamar Blue assay (Fig. 1B). Whole-well ATP measurements indicated stable ATP production up to day 7, followed by a gradual decline to ~50 % by days 14 and 21 (Fig. 1C). However, when normalized to the percentage of viable cells (based on the Alamar Blue assay), ATP levels per cell remained constant (Fig. D). Furthermore, flow cytometry analysis of propidium iodide-stained cells confirmed intact membrane integrity in the remaining hemocytes (Fig. 1E). These findings indicate that despite the initial partial cell loss, the remaining hemocyte subpopulation maintained both membrane integrity and metabolic activity. Notably, while our primary focus was to characterize hemocyte dynamics over this period, some cultures remained viable beyond 21 days, with a few persisting for over two months.

## 3.2. Hemocyte diversity and morphology are sustained during early culture stages, followed by morphology shifts for the remaining cells

The diversity of morphologically distinct cell types (morphotypes; see section 2.4 for a more complete definition of this term) remained similar between Day 0 and Day 7 of culture (Figs. 1 and 2). During this first week, seven major morphotypes were identified, corresponding to the three commonly described hemocyte subpopulations: granulocytes, hyalinocytes, and blast-like cells (Fig. 2). Among granulocytes, three distinct morphotypes were observed: one with low spreading, typically round or dendritic-like (Fig. 2A), and two with high spreading, exhibiting either a semi-circular "half-moon" shape (Fig. 2B) or a macrophage-like "fried-egg" appearance (Fig. 2C). Similarly, hyalinocytes displayed three morphotypes: one with low spreading, also round or dendritic-like (Fig. 2D), and two with high spreading, appearing either as small irregular or half-moon-shaped cells (Fig. 2E) or as large, elongated fibroblast-like cells (Fig. 2F). Blast-like cells, characterized by a high nucleus-to-cytoplasm ratio and small size, were also present (Fig. 2G). It is important to note that the terms 'small' and 'large' are used qualitatively based on visual comparison (see scale bars in Fig. 2), and not from direct cell size measurements.

Between days 14 and 21, all three major hemocyte subpopulations remained represented, though blast-like cells and large highly spread hyalinocytes became less frequent (Fig. 1). When present, these fibroblast-like hyalinocytes appeared larger and more irregularly shaped (Fig. 2K and P). Over this period, some granulocytes and hyalinocytes remained with a similar morphology as in the early culture stages (Fig. 2H, I, 2L, and 2N), while others exhibited a round or oval morphology (Fig. 2H, J, 2M, and 2O). By day 21, granulocytes were the predominant cell type, with fewer hyalinocytes and only a small fraction of fibroblast-like cells remaining (Figs. 1 and 2).

This decline in morphological diversity correlated with the appearance of cells displaying hallmarks of apoptosis, such as nuclear condensation, cell shrinkage, and nuclear fragmentation, as evidenced by MGG staining. Cells presenting these features were first observed at day 7 and persisted, albeit at lower frequencies, on days 14 and 21. Additional nuclear abnormalities, including micronuclei and nuclear blebbing, were primarily observed on day 7, particularly among large, highly spread hyalinocytes (Fig. 2R). Highly pigmented granules appearing in cells with intact nuclei were also observed, suggesting the engulfment of apoptotic corpses by the remaining live cells (Fig. 2S, U, 2X, and 2Y). These granules were predominantly observed in hyalinocytes, especially from day 7 onward (Fig. 2S, U, and 2X), but by day 21, granulocytes also contained these granules (Fig. 2Y).

Flow cytometry further supported these morphological trends, revealing a time-dependent increase in mean cellular complexity (Fig. 3A). Additionally, a marginal but significant increase in cell size was observed at day 21. These differences were clearly reflected in forward scatter (cell size) vs. side scatter (cell complexity) dot plots, which illustrated shifts in cell distribution over time (Fig. 3B).
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** (A) Primary cultures of hemocytes from triploid *Crassostrea gigas* oysters maintained for 21 days in 24-well plates (500,000 cells/well) coated with poly-D-lysine, using filtered, sterile oyster plasma supplemented with antibiotics (FSOP + PS) as the culture medium. Scale bar = 50 µm. (B) Cell viability assessed using Alamar Blue. (C) Total ATP levels quantified per well. (D) ATP levels normalized to the percentage of viable cells as determined by the Alamar Blue assay. (E) Cell viability assessed using propidium iodide. Data are presented as means ± standard deviation. Groups labeled with different letters indicate significant differences (p < 0.05).</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Morphotypes (top panel) and deformities (bottom panel) of triploid *Crassostrea gigas* hemocytes in primary culture over time. Cells were cultivated with filtered-sterile oyster's blood plasma with antibiotics (FSOP + PS) in 96-well plates (µClear bottom) coated with poly-D-lysine and colored with May-Grünwald Giemsa. **Morphotypes at days 0 and 7:** (A) small granulocytes with round or dendritic-like shape; (B) large granulocytes with a semi-circular or half-moon shape; (C) large granulocytes with a macrophage-like or fried-egg shape; (D) small hyalinocytes with round shape; (E) small hyalinocytes with irregular or half-moon shape; (F) large hyalinocytes with fibroblast-like or irregular shape; (G) blast-like cells with high nucleus/cytoplasm ratio and small size. **Morphotypes at day 14:** (H) granulocytes with mostly round or oval shape; (I) small hyalinocytes with mostly round shape; (J) hyalinocytes with oval or dendritic-like shape; (K) large hyalinocytes with fibroblast-like or irregular shape. **Morphotypes at day 21:** granulocytes with mostly round or oval shape presenting two different size categories overall; small (L) and large (M); (N) small hyalinocytes with mostly round shape; (O) larger hyalinocytes with oval shape; (P) large hyalinocytes with mostly irregular shape. **Nuclei and cellular deformities at days 7, 14, and 21:** (Q, T, and V) cells presenting the typical signs of apoptosis such as nuclear condensation, loss of cell volume, and/or nuclear fragmentation; (R) nuclear abnormalities on day 7 cultures such as micronuclei and blebbing or amorphous nuclei; (S, U, X, and Y) highly pigmented granules within cells presenting normal nucleus morphology, suggesting phagocytosis of apoptotic corpses. Bar = 10 µm for all images.</figcaption></figure>

Together, these results indicate that while hemocyte morphotype diversity remains stable during the early stages of culture (up to Day 7), a progressive shift toward a more complex population occurs over time, suggesting that granulocytes represent a more resilient subpopulation in our plasma-based cultures.

### 3.3. Resilient hemocyte morphotypes maintain motility and phagocytosis behaviors

Despite the gradual loss of cells over time, live-cell imaging revealed that the remaining hemocyte subpopulations retained their motility
<figure><img src="image_7.png" /><figcaption>**Fig. 3.** Flow cytometry analyses of the size and complexity of triploid oyster *Crassostrea gigas* hemocytes in primary culture over time. Cells (500,000/well) were cultivated with filtered-sterile oyster's blood plasma containing antibiotics (FSOP + PS) in 24-well plates coated with poly-D-lysine. (A) Geometric means of cell size and complexity; and (B) representative forward scatter (FSC-A; cell size) vs side scatter (SSC-A; cell complexity) dot plots, showing the cell distribution within the primary cultures throughout time as compared to control (Day 0, cells freshly collected). Bars represent means ± standard deviation. Groups labeled with different letters indicate significant differences (p < 0.05).</figcaption></figure>

behavior (see Supplementary Video Files 1–12). Throughout all culture days, cells primarily exhibited two movement strategies: pseudopod-driven motility (Fig. 4A; Supplementary Video File 13) and lamellipodium-driven motility, resembling mesenchymal and amoeboid migration modes, respectively (Fig. 4B; Supplementary Video File 14). While some cells maintained a single motility type throughout the recorded timeframe, others alternated between both modes (see

Supplementary Video 2 for clear examples).

The time-lapse videos further revealed that cells displaying lamellipodium-driven motility often acquired a transient half-moon shape (Figs. 4B and 120s; Supplementary Video File 14). This suggests that the half-moon morphotype, previously identified through MGG staining, does not correspond to a distinct hemocyte subpopulation but rather represents a dynamic movement state. Consequently, while MGG

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Cell movement behavior of triploid *Crassostrea gigas* hemocytes in primary cultures. Cells were cultivated with filtered-sterile oyster's blood plasma with antibiotics (FSOP + PS) in 96-well plates (μClear bottom) coated with poly-D-lysine. Live-cell imaging revealed two main cell movement strategies employed by the hemocytes in culture: movement through pseudopod or lobopodian formation (A), and movement through lamellipodium formation (B). (C) Cell membrane movement in non-motile highly spread cells. Bar = 10 µm.</figcaption></figure>
staining initially indicated the presence of seven distinct morphotypes, incorporating live-cell imaging findings supports the existence of only five distinguishable hemocyte subpopulations in our plasma-based primary cultures.

In contrast, large, highly spread hyalinocytes (fibroblast-like cells; Fig. 2F) displayed only membrane expansion and retraction (Fig. 4C), without active migration within the analyzed timeframe (up to 5 min).

In addition to maintaining cell motility, the resilient hemocyte sub-populations also preserved their phagocytic capacity, as evidenced by their ability to engulf polystyrene microbeads throughout the entire culture period (Fig. 5). The percentage of phagocytic cells, relative to the total viable cell population, remained stable over the 21-day period (Fig. 5A). Notably, the number of microbeads engulfed per cell significantly increased at days 14 and 21 (Fig. 5B). These observations were further validated through microscopy (Fig. 5C), which revealed that day 21 cultures contained cells capable of engulfing more microbeads (Fig. 5C-e*) than those observed at earlier time points (Fig. 5C-e).

### 3.4. Live-cell imaging reveals that mobility and phagocytosis are not systematically correlated in vitro

Additional insights into the phagocytic behavior of distinct hemocyte morphotypes were gained through live-cell imaging. Highly spread fibroblast-like cells, for example, did not exhibit phagocytic activity (Fig. 5C-a). Moreover, not all highly mobile cells were phagocytic; some motile cells, both with (Fig. 5C-c) and without (Fig. 5C-d) granules, did not engulf microbeads (see Supplementary Video Files 15 and 16, respectively). Interestingly, phagocytosis was also observed in non-motile or low-motility cells, such as highly spread macrophage-like (fried-egg-shaped) cells (Fig. 5C-b; Supplementary Video File 17). These findings, observed throughout all stages of culture, suggest that key functional traits such as motility and phagocytosis do not necessarily overlap in vitro.

### 3.5. Plasma-based cultures maintain hemocytes with stable mitochondrial activity and low oxidative stress but promote lipid accumulation

Additional assays were conducted to evaluate the overall metabolic and physiological state of hemocytes in primary plasma-based cultures.

<figure><img src="image_9.png" /></figure>

**Fig. 5.** Phagocytic behavior of triploid *Crassostrea gigas* hemocytes in primary culture over time. Cells were cultivated with filtered-sterile oyster's blood plasma with antibiotics (FSOP + PS) in either 24-well plates (A and B) or 96-well plates (µClear bottom) (C) coated with poly-D-lysine and incubated with microsphere beads (2 µm) for 2 (A and B) or 4 (C) hours at 16 ± 1 °C before imaging. (A) Percentage of cells capable of engulfing polystyrene beads, and (B) the average number of polystyrene beads engulfed per cell. Groups labeled with different letters indicate significant differences (p < 0.05). (C) Detailed observation of images and time-lapse videos (supplementary video files 15–17) revealed distinct phagocytic behaviors of the different cell morphotypes: highly spread fibroblast-like cells without phagocytic behavior (a); (ii) highly spread macrophage-like (fried-egg) cells which did not display mobility with phagocytosis behavior (b); highly mobile cells with (c) or without (d) granules without phagocytosis behavior; highly mobile cells capable of concentrating a great number of beads (e), and possibly an even greater amount within the day 21 cultures (e*). Bar = 10 µm.
Single-cell flow cytometry analysis revealed no significant changes in mitochondrial ROS production (Fig. 6A) or mitochondrial content over time (Fig. 6B). However, hemocytes in culture exhibited lower intracellular ROS levels at all time points analyzed (Fig. 6C). Finally, analysis of neutral lipid reserves showed a time-dependent accumulation of lipids, with statistically significant increases observed in day 21 cultures (Fig. 6D).

## 4. Discussion

At present, primary culture is the only approach for culturing mollusk cells as not a single marine mollusk cell line is available, [39]. Although immortalized cell lines offer the advantage of ease of maintenance and long-term reproducibility, primary cell cultures also present notable advantages, as they more closely approximate the *in vivo* environment and retain key characteristics of the original tissue. Consequently, compared to cell lines, they provide a more relevant platform for investigating the physiology and biochemistry of the whole organism [40,41]. Researchers have been developing primary culture systems for bivalves since the mid-1900s, which led to the initial establishment of oyster heart and mantle tissue cell cultures [42]. Even though such cell types can be cultured for several weeks and even months, hemocytes are still bivalves most used primary cells [39] and the subject of great interest in biology and environmental sciences due to their crucial role in immune defense and maintenance of bivalve health and as key indicators of organism physiological status [2,43,44]. Given the significance of marine bivalve aquaculture to the global economy and

prevailing disease threats, the need for a marine bivalve immune cell culture tool is critical [45]. However, until now, researchers have relied mostly on short-term cultures of bivalve hemocytes, limiting their applicability in research.

As mentioned in the Introduction section, maintaining bivalve hemocytes viable and functional under *in vitro* conditions has been a major challenge [27]. A common feature of the studies mentioned above and many others is the use of commercial culture media, particularly Eagle's Basal Medium or Leibovitz L-15, and supplementation with fetal bovine serum or glucose [27,33,39] or 0.22 µm filtered seawater [20,46]. It is important to note that Eagle's Basal Medium and L-15 medium do not contain certain amino acids like proline and taurine found in high concentrations in oyster hemolymph [47]. In addition, traditional cell culture media are developed for mammalian cells to saturate cell culture with the necessary nutrients to maintain stable and maximal proliferation of cancer cells [48]. However, these nutrients are not necessarily suitable for hemocyte primary cultures. Such balance between culture medium components is critical to ensure hemocyte health and immunological function during primary culture conditions.

To better mimic the hemocyte *in vivo* microenvironment, we used *C. gigas* oyster hemolymph plasma as the culture medium. By using a plasma-based medium, our approach likely offers a culture environment that more closely mimics *in vivo* conditions than standard commercial media. Indeed, the use of a plasma-like cell culture medium has also recently been adopted for mammalian cell culture, based on increasing evidence that artificial cell media can interfere with cellular processes and metabolism [48]. Hemolymph plasma also contains key immune

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /><figcaption>**Fig. 6.** Metabolism-related parameters of triploid *Crassostrea gigas* hemocytes in primary culture over time. Cells were cultivated with filtered-sterile oyster's blood plasma with antibiotics (FSOP + PS) in 24-well and harvested at the specified days of culture. Day 0 corresponds to cells freshly collected from oysters and not placed in culture. Flow cytometry analyses were made to quantify (A) mitochondrial ROS production, (B) mitochondrial content, (C) total basal ROS levels, and (D) lipid reserves per cell. Bars represent means ± standard deviation. Groups labeled with different letters indicate significant differences (p < 0.05).</figcaption></figure>
components such as neurotransmitters, opsonins, hormones, and LPS-binding proteins [49,50]. For example, it is well-established that the presence of hemolymph plasma enhances the *in vitro* binding, internalization, and killing of bacteria [51], making it an ideal cellular medium for studies on immune responses, inflammation, and host-microbe interactions.

Nonetheless, one major disadvantage of using a plasma-based medium is the lack of standardized formulation. Indeed, oyster plasma composition can vary among individuals and environmental factors [52, 53], thus possibly introducing differences in cellular responses to abiotic and biotic factors. To minimize this drawback, we adopted the strategy of creating a C. gigas plasma bank to be used throughout a same set of experiments. Nevertheless, it is important to note that the development of a standardized plasma-like cell culture medium will be crucial for long-term applications. Further optimization is needed to create a medium that can perhaps support extended *in vitro* hemocyte survival and cell proliferation. Our protocol, therefore, provides a valuable foundation for testing additional molecules that could enhance culture conditions and contribute to the formulation of a standardized medium in the future.

Our cell culture protocol maintained the fresh hemolymph diversity of C. gigas morphotypes for up to seven days, while a subset of more resilient morphotypes persisted for at least 21 days, with some cultures extending to two months. By day 21, viability assessments indicated that approximately half of the initial cell quantity remained viable. In this study, we used two complementary methods to assess hemocyte viability: Alamar Blue assay and propidium iodide staining. Alamar Blue measures overall metabolic activity (NADH production rate) at the well level, with reductions indicating a decrease in the number of metabolically active cells. In contrast, propidium iodide staining, analyzed through flow cytometry, assesses membrane integrity on a per-cell basis among all cells that remain in the wells. While dead or dying cells can still be detected by propidium iodide, the phagocytic activity of hemocytes may lead to the clearance of dead cells, potentially underestimating their presence, which is supported by our images showing the presence of apoptotic bodies within the remaining cells. Together, these complementary approaches capture both the loss of viable cells and the integrity of those that persist.

Morphological analyses based on phase contrast and MGG images revealed that the decrease in the percentage of viable cells was accompanied by a decline in cell diversity after 14 and 21 days, marked by a reduction in blast-like and fibroblast-like cells. This decline, along with the presence of apoptotic bodies, suggests that certain hemocyte subtypes may be more prone to physiological decline and senescence under our *in vitro* conditions. The lifespan of bivalve hemocytes *in vivo* remains poorly understood, but in mammals, some immune cells such as neutrophils are short-lived typically persisting for only about 5 days [54]. Thus, rather than a random loss of cells, this cell-specific decline over time could reflect inherent differences in hemocyte longevity and *in vitro* stability. Future studies investigating hemocyte turnover in our plasma-based primary cultures could offer valuable insights into oyster hemocyte lifespan and senescence, which remain poorly understood in bivalves.

On the other hand, our protocol revealed a resilient subpopulation of cells that maintained membrane integrity, ATP production, and phagocytic activity, with no apparent decline in these parameters over time. These cells also exhibited increased morphological complexity, consistent with an enrichment of granulocytes. It is important to note that some of the highly adherent hyalinocytes (mainly fibroblast-like cells and some macrophage-like cells) could not be recovered from wells for the flow cytometry analyses, as we could not apply any trypsin treatment. This was a choice to avoid cell damage and dysfunction, as we used such samples for further physiological analyses such as immunocompetence and metabolism. Thus, even though this approach might have introduced a bias toward the quantification of overall population complexity in the culture over time through flow cytometry, it is evident

through whole primary culture images and videos (supplementary video files 1–12) that highly adherent hyalinocytes were much less present and granulocytes more enriched in cultures after 14 days. A lower proportion of highly adherent hyalinocytes in older cultures was also evident through images taken after cells were harvested for the flow cytometry analyses (Supp. Fig. 1).

Live-cell imaging has become an invaluable technique in cell biology, allowing researchers to observe dynamic processes in living cells in real-time. In this study, we innovatively adopted this non-invasive, label-free approach to monitor hemocytes over an extended period *in vitro*. Our study extends previous research on hemocyte motility by tracking cell behavior over a longer period *in vitro*, yet our findings remain consistent with reports that bivalve hemocytes are typically active and mobile [54–56]. The combination of live-cell imaging with MGG staining was crucial to determine that some morphotypes corresponded to different states of the same hemocyte subtype (see Fig. 4 and supplementary video files 1–17). Therefore, as mentioned in Section 3.3, while MGG staining initially indicated the presence of seven distinct morphotypes, incorporating these live-cell imaging findings supports the existence of only five distinguishable hemocyte subpopulations in our plasma-based primary cultures: (i) blast-like cells, (ii) small granulocytes (dendritic-like cells), (iii) small hyalinocytes, (iv) large granulocytes (macrophage-like cells), and (v) large hyalinocytes (fibroblast-like cells). All subtypes exhibited motility, except for the fibroblast-like cells. These findings slightly expand the current understanding of C. gigas hemocyte subpopulations, which typically identify three main types: blast-like cells, hyalinocytes, and granulocytes [2]. This broader morphological diversity may be attributed to our culture setup, which allowed cells to attach and spread freely on poly-D-lysine-coated plastic. In contrast, many previous studies analyzed fixed hemocytes, which can restrict cells from displaying their full range of plasticity and dynamic morphological transitions. Nonetheless, De La Forest Divonne et al. [4] identified seven distinct hemocyte types in C. gigas: two subpopulations of blast-like cells (one basophilic and another acidophilic), one subpopulation of hyalinocytes, three subpopulations of granulocytes (macrophage-like cells, small granule cell, and big granule cell), and vesicular cells. Their description is overall in line with our observations, except that their study did not describe small hyalinocytes and we did not detect the big granule cells and vesicular cells. These last two cell types were described in low frequencies (4 % and 2 %, respectively) in their study, which may explain why we did not observe them in our cultures.

Live-cell imaging uncovered real-time dynamic phenotypes of hemocyte subpopulations, facilitating morphotype-function mapping and broadening our understanding of cellular immunophysiology. Notably, while some highly motile granulocytes exhibited phagocytic activity, others lacked this capacity. In contrast, large hyalinocytes (fibroblast-like cells) were spread out, adhered to the culture plate, and displayed neither motility nor phagocytic activity. These observations suggest that granulocyte subpopulations can be categorized into phagocytic and non-phagocytic groups, each likely serving distinct physiological roles. The lack of motility and phagocytosis in large hyalinocytes suggests their involvement in aggregation and wound healing, as seen in other bivalve species [2]. Supporting our findings, De La Forest Divonne et al. [4] identified two of three granulocyte subtypes as professional phagocytes: the macrophage-like cells (likely corresponding to the macrophage-like cells described in our study) and the small granule cells (likely corresponding to our small dendritic-like granulocytes). Additionally, their observation that large hyalinocytes do not exhibit phagocytic activity further aligns with our results. However, other studies have reported that hyalinocytes from oysters, including C. gigas, exhibit phagocytic activity [2,57], highlighting a discrepancy that warrants further investigation into the factors influencing these functional differences. Altogether, these results support functional specialization within hemocyte lineages, reminiscent of cellular compartmentalization observed in higher organisms, with potential implications for comparative
immunology and cell-type evolution.

Phagocytosis is a key indicator of immune function in hemocytes [58–60]. Our protocol demonstrated that hemocyte phagocytic activity and capacity were maintained over 21 days of culture. Interestingly, we observed an increase in phagocytic capacity (i.e., the number of beads engulfed per cell) within a resistant cell subpopulation by day 21. Hemocyte phagocytosis is known to fluctuate seasonally [61], in response to infection and disease susceptibility [62], and under varying environmental conditions [63]. However, the reason for the enhanced phagocytic capacity in these granulocytes at later culture stages remains unclear. Previous studies have shown that certain granulocyte subtypes exhibit significantly higher phagocytic activity [4,57], potentially influenced by the presence of β-glucan receptors [57]. This raises the question of whether the remaining granulocytes at day 21 represent an enrichment of a more phagocytosis-competent subtype and/or an upregulation of these receptors. Additionally, whether this functional enhancement extends to other immune parameters remains an open question and could provide valuable insights into the immune plasticity of hemocytes *in vitro*.

Together with recent efforts, our study underscores the need for further research integrating morphological, functional, molecular, and live-cell imaging analyses to achieve a more comprehensive and unified characterization of hemocyte types and their roles in oyster health and physiology.

To further understand the physiology of the hemocyte in our plasma-
based cultures, we analyzed key metabolic parameters to investigate
possible metabolic alterations and reprogramming in response to the *in vitro* settings. ROS production is closely related to cellular metabolic
activity. Indeed, mitochondrial ROS is the major contributor to ROS
levels in *C. gigas* hemocytes [64]. In this study, we have shown that total
basal ROS production is higher in fresh hemocytes on day 0, then be-
comes lower and stable until day 21 cultures. Recently, we have found
that, in *C. gigas*, the average physiological oxygen level within oyster
hemolymph is ~5 kPa, but it can vary between 0 kPa and 12 kPa
depending on valve opening behavior and filtering activities [23].
However, ambient air is composed of 21 kPa O₂. Thus, when hemocytes
are transferred from the *in vivo* to *in vitro* conditions, they are exposed to
much higher levels of oxygen than they would naturally experience
(hyperoxia). Indeed, ROS production in cells is known to increase at
hyperoxia conditions [65,66], which could explain the high ROS levels
in the fresh hemocytes (day 0). The subsequent decrease and stabiliza-
tion of ROS production at the following periods of culture (Days 7, 14,
and 21) can be due to hemocyte acclimation mechanisms. We hypoth-
esize that such mechanisms may be mediated by the activation of the
nuclear factor erythroid 2-related factor 2 (Nrf2) transcription factor
which is a master regulator of antioxidant cellular responses [23]. This
hypothesis is supported by the fact that the conserved role of Nrf2 in
mediating redox changes was already confirmed in *C. gigas* [23,67,68],
and that Nrf2 activation in cells submitted to hyperoxia condition
(ambient air) is a well-described phenomenon within mammalian cell
culture models [69]. Future studies performing gene expression analyses
of Nrf2-target genes over the course of the hemocyte primary cultures
should be performed to confirm this hypothesis. Despite this decrease in
total ROS levels, we found stable levels of mitochondrial ROS
throughout the 21 days of culture. The differences reported between the
total basal ROS and mitochondrial ROS can be related to differences in
probe specificities. DCF is a broader marker of ROS, reacting with
several different ROS types, whereas MitoSOX is a marker specific for
superoxide anion related to mitochondrial activity, membrane potential,
and size [70,71]. Therefore, these results suggest that the lower total
basal ROS levels of hemocytes in culture are not linked to a decrease in
mitochondrial activity. In line with this hypothesis, the analysis of total
mitochondrial content remained constant in hemocytes throughout the
cell culture period. Cellular stress and mitochondrial dysfunction usu-
ally alter mitochondrial dynamics, often leading to changes in mito-
chondrial content through mitophagy or biogenesis mechanisms, for

example [72]. Since this was not the case in our study, our primary
hemocyte culture model offers stable conditions to study mitochondrial
dynamics under different stress and disease scenarios. Collectively,
these findings indicate that this *in vitro* model provides a controlled
experimental system for investigating how immune cells from marine
invertebrates adapt to oxidative stress at the molecular level and both
conserved and species-specific aspects of immunometabolic regulation.

Lipid droplets act as dynamic organelles for storing neutral lipids and play a crucial role in energy homeostasis and lipid metabolism. They are involved in various cellular processes, including the regulation of lipid synthesis and degradation, signaling pathways, and the maintenance of cellular lipid balance [73]. In our study, we have identified changes in intracellular dynamics of neutral lipid storage in hemocytes, leading to higher lipid accumulation at 21 days of cell culture. Lipid accumulation in cultured cells can be a sign of changes in lipid metabolism, or the ability to take up lipids from the media, or as a result of autophagic processes [74]. Indeed, to interpret our result, there are at least three critical points to consider. Firstly, in bivalves, hemocytes are known to transport nutrients, including lipids, by absorbing them from the alimentary tract and transferring them to other tissues or developing oocytes [75,76]. Secondly, granulocytes, which were enriched at the 21-day cultures, have a higher neutral lipid content than other cell types in marine bivalves [77]. Thirdly, reserves of neutral lipids are related to the balance between synthesis and degradation in response to metabolic needs and shifts. Thus, it is likely that the absence of interaction between hemocytes and other tissues in the cultures blocked their ability to transport lipids to other cells, promoting a higher accumulation of lipids absorbed through the cell culture medium (FSOP). This higher neutral lipid accumulation could be related to the larger cell size observed after 21 days of culture. Taken together, the increased levels of neutral lipids in hemocytes during the second and third weeks of culture may be associated with changes in cell size, composition, and energy allocation. These findings highlight the potential of this *in vitro* model to investigate the largely understudied mechanisms of lipid storage in bivalve hemocytes and their metabolic plasticity, particularly in the context of lipid utilization as a potential energy source under stress conditions.

Altogether, our results reveal a few dynamic changes of hemocytes in
plasma-based cultures over 21 days. Although some of these changes
may represent a limitation when studying the biology of hemocytes in
extended culture periods, it adds important information on the impli-
cations of cell microenvironment conditions to the physiology of bivalve
hemocytes. The framework presented here opens new avenues to
explore biotic and abiotic conditions that can further improve hemocyte
primary cultures and potentially even develop a cell line if conditions
favoring proliferation are found. Moreover, it also provides the possi-
bility to investigate the behavior of cell cultures under different stress
conditions. For example, it is possible now to explore whether patho-
gens, contaminants, temperature, or plasma composition alterations will
affect hemocyte viability and function over extended periods. Thus, our
oyster hemocyte primary culture protocol, allied with the cell-based
assays optimized in this study, allows the investigation of factors that
can induce stress responses affecting cellular morphology, behavior,
viability, immunocompetence, mitochondrial metabolism, and lipid
storage. This could lead to new insights and discoveries in hemocyte
biology and ecotoxicology research, thus expanding our understanding
of bivalves’ immunological and metabolic responses to the current and
future challenges of our changing ocean.

In this study, we opted to use triploid oysters, which are functionally
sterile and produce significantly fewer gametes [78]. Our choice was
motivated by the need for a consistent, year-round supply of oysters and
high-quality hemolymph, without the major complication of gamete
contamination. It is well established that the reproductive stage can
significantly alter hemocyte parameters in bivalves [79] including
*C. gigas* [80,81], likely due to energy reallocation during different stages
of reproduction. Consequently, the use of triploid oysters offers the
advantage of greater stability in hemocyte parameters throughout the
year compared to diploid oysters [80,81], making our model more reliable and reproducible across seasons. It is also worth noting that triploid oysters account for 30–50 % of global oyster production [78] and improving their survival and immune resilience remains a major focus in aquaculture, for which our model is directly applicable. Nevertheless, a limitation of this approach is the lack of direct representation of wild diploid oyster populations, whose hemocyte dynamics and *in vitro* responses may differ. Addressing these differences, including potential variability in hemocyte parameters linked to reproductive stage and sex, would be an important direction for future research.

Despite the strengths of our approach, this study has several limitations that warrant consideration. Some assays required cell detachment, which may introduce bias by favoring less adherent subpopulations. Additionally, while we used flow cytometry to assess general cell complexity and viability, it did not allow for clear discrimination between hemocyte subtypes. In parallel, our classification into morphotypes was based solely on morphological criteria under *in vitro* conditions and not supported by molecular or functional markers. Additionally, while we provided qualitative insights into hemocyte behavior and morphology, more robust quantification, ideally supported by automated cell imaging systems, would strengthen our conclusions and allow for more objective comparisons across time points. Future work should aim to integrate higher-resolution approaches, such as single-cell transcriptomics or immunophenotyping, to refine hemocyte classification and subtype-specific analyses. Apoptotic features were identified morphologically but not confirmed with specific assays, and the use of poly-D-lysine, while beneficial for cell culture stability, may influence cell behavior. Lastly, as only triploid oysters were used, our findings may not fully represent the diversity of hemocyte responses in diploid or wild populations. These limitations highlight key areas for technical refinement and future research. Nonetheless, our study provides a reliable, reproducible platform for long-term *in vitro* culture of *C. gigas* hemocytes, offering a foundation for improved and standardized bivalve immune cell models.

## 5. Concluding remarks and perspectives

This study presents a comprehensive framework for investigating oyster hemocyte biology *in vitro*, offering valuable insights into immune competence, metabolic plasticity, and cellular behavior over an extended 21-day period. By maintaining hemocytes in their natural medium, hemolymph plasma, our protocol successfully preserved a diverse population of hemocyte morphotypes for up to 7 days, with granulocytes emerging as the most resilient subpopulation during longer culture durations. Importantly, our findings highlight the persistence of key immune and cellular functions throughout the culture period. Additionally, metabolic assessments revealed that surviving hemocytes not only maintained certain metabolic parameters but also demonstrated physiological plasticity. These adaptations underscore the ability of hemocytes to modulate their physiological state in response to prolonged *in vitro* conditions and establish a foundation for optimizing culture parameters to enhance *in vitro* resilience and potentially achieve hemocyte de-differentiation and proliferation.

Altogether, by integrating real-time imaging, metabolic assays, and immunological readouts, this platform enables in-depth analyses of bivalve hemocyte behavior, function, and fate over time. This approach opens new opportunities to explore fundamental cellular processes such as senescence, immunometabolic reprogramming, mitochondrial plasticity, and redox regulation, which are essential for understanding how bivalve cells adapt to the fluctuating stresses of their marine environments.

Beyond its relevance to bivalve biology, this system provides a valuable comparative model for studying conserved and divergent pathways of cellular stress adaptation across invertebrates and vertebrates. Additionally, it serves as a versatile tool for assessing cellular responses to environmental stressors, pathogens, and chemical

exposure, with potential applications in ecophysiology, immunology, ecotoxicology, molecular screening, and health innovation.

## CRediT authorship contribution statement

**Danielle F. Mello:** Writing – original draft, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization. **Rafael Trevisan:** Writing – review & editing, Methodology, Investigation, Formal analysis. **Yasmine Even:** Writing – review & editing, Methodology, Investigation, Data curation. **Valentin Foulon:** Writing – review & editing, Methodology. **Christophe Lambert:** Writing – review & editing, Methodology. **Heloïsa B. Gabe:** Writing – review & editing, Methodology. **Manon Le Goff:** Writing – review & editing, Methodology. **Claudie Quéré:** Writing – review & editing, Methodology. **Fernando R. Queiroga:** Writing – review & editing, Methodology. **Christophe Brigaudeau:** Writing – review & editing, Conceptualization. **Hélène Talarmin:** Writing – review & editing, Conceptualization. **Caroline Montagnani:** Writing – review & editing, Supervision, Conceptualization. **Guillaume M. Charrière:** Writing – review & editing, Supervision, Methodology, Conceptualization. **Delphine Destoumieux-Garzon:** Writing – review & editing, Supervision. **Stéphanie Madec:** Writing – review & editing, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Conceptualization. **Charlotte Corporeau:** Writing – review & editing, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Conceptualization.

## Declaration of generative AI and AI-assisted technologies in the writing process

During the preparation of this work, the authors used ChatGPT-4 and Grammarly version 14.1229.0 in order to improve English fluency and enhance clarity. After using these tools, the authors reviewed and edited the content as needed and take full responsibility for the content of the publication.

## Funding

The majority of this work was conducted thanks to a Postdoctoral fellowship from the French government (Finistère Appel APRE and Région Bretagne Appel SAD n°2050). Currently, DF Mello as well as R Trevisan, and FR Queiroga are recipients of postdoctoral fellowships through the Bienvenue Program, co-funded by the European Union's Horizon 2020 research and innovation programme under the Marie Skłodowska-Curie grant agreement number 899546, the Region Bretagne, and the Université de Bretagne Occidentale. HB Gabe was recipient of a sandwich doctorate scholarship from the Government of Brazil funded by the “Conselho Nacional de Desenvolvimento Científico e Tecnológico – CNPq” (Processo 200247/2022-0). This work was supported by ISblue project, Interdisciplinary graduate school for the blue planet (ANR-17-EURE-0015) and co-funded by a grant from the French government under the program “Investissements d’Avenir”; embedded in France 2030. This study is set within the framework of the « Laboratoire d’Excellence (LabEx) » TULIP (ANR-10-LABX-41). It was supported by the École Universitaire de Recherche TULIP-GS (ANR-18-EURE-0019).

## Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

The authors would like to thank Christine Dubreuil, Olivier Mouchel,
Moussa Diagne, Hugo Koechlin, Luc Lebrun, Nelly Le Goic, and Emma Denmat for their support during the experiments; Philippe Elies for his support to acquire the confocal microscopy images; Claire Hellio for manuscript revision; and Sébastien Herve for the graphical abstract.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.yexcr.2025.114662.

### Data availability

Data will be made available on request.

## References

[1] T.P. Yoshino, U. Bickham, C.J. Bayne, Molluscan cells in culture: primary cell cultures and cell lines, Can. J. Zool. 91 (2013), https://doi.org/10.1139/cjz-2012-0258.

[2] N.R. de la Ballina, F. Maresca, A. Cao, A. Villalba, Bivalve haemocyte subpopulations: a review, Front. Immunol. 13 (2022) 826255, https://doi.org/10.3389/FIMMU.2022.826255/BIBTEX.

[3] L. Song, L. Wang, L. Qiu, Z. Huan, Bivalve immunity, in: K. Söderhäll (Ed.), Invertebrate Immunity, Landes Bioscience and Springer Science+Business Media, New York, 2010, pp. 44–65.

[4] S. de L.F. Divonne, J. Pouzadoux, O. Romatif, C. Montagnani, G. Mitta, D. Destoumieux-Garzon, B. Gourbal, G.M. Charrière, E. Vignal, Diversity and functional specialization of oyster immune cells uncovered by integrative single cell level investigations, eLife 13 (2024), https://doi.org/10.7554/ELIFE.102622.1.

[5] B.L. Bayne, M.N. Moore, J. Widdows, D.R. Livingstone, P. Salkeld, Measurement of the responses of individuals to environmental stress and pollution: studies with bivalve molluscs, Philos. Trans. R. Soc. Lond. B Biol. Sci. 286 (1979) 563–581, https://doi.org/10.1098/RSTB.1979.0046.

[6] R.K. Pipe, J.A. Coles, Environmental contaminants influencing immunefunction in marine bivalve molluscs, Fish Shellfish Immunol. 5 (1995) 581–595, https://doi.org/10.1016/S1050-4648(95)0043-3.

[7] T. Renault, Immunotoxicological effects of environmental contaminants on marine bivalves, Fish Shellfish Immunol. 46 (2015) 88–93, https://doi.org/10.1016/J.FSI.2015.04.011.

[8] M. Auffret, Bivalves as models for marine immunotoxicology, in: Investigative Immunotoxicology, CRC Press, 2005, pp. 49–68, https://doi.org/10.1201/9781420036817-8.

[9] M. Abdou, L. Dutruch, J. Schäfer, B. Zaldibar, R. Medrano, U. Izagirre, T. Gil-Díaz, C. Bossy, C. Catrouillet, R. Hu, A. Coynel, A. Lerat, A. Cobelo-García, G. Blanc, M. Soto, Tracing platinum accumulation kinetics in oyster *Crassostrea gigas*, a sentinel species in coastal marine environments, Sci. Total Environ. 615 (2018) 652–663, https://doi.org/10.1016/J.SCITOTENV.2017.09.078.

[10] E. David, A. Tanguy, R. Riso, L. Quiniou, J. Laroche, D. Moraga, Responses of Pacific oyster *Crassostrea gigas* populations to abiotic stress in environmentally contrasted estuaries along the Atlantic coast of France, Aquat. Toxicol. 109 (2012) 70–79, https://doi.org/10.1016/J.AQUATOX.2011.11.014.

[11] D.F. Mello, R. Trevisan, N.M. Danielli, A.L. Dafre, Vulnerability of glutathione-depleted *Crassostrea gigas* oysters to Vibrio species, Mar. Environ. Res. 154 (2020) 104870, https://doi.org/10.1016/j.marenvres.2019.104870.

[12] L. Nogueira, D.F. Mello, R. Trevisan, D. Garcia, D. da Silva Acosta, A.L. Dafre, E. A. de Almeida, Hypoxia effects on oxidative stress and immunocompetence biomarkers in the mussel *Perna perna* (Mytilidae, Bivalvia), Mar. Environ. Res. 126 (2017) 109–115, https://doi.org/10.1016/j.marenvres.2017.02.009.

[13] R. Trevisan, D.F. Mello, M. Uliano-Silva, G. Delapedra, M. Arl, A.L. Dafre, The biological importance of glutathione peroxidase and peroxiredoxin backup systems in bivalves during peroxide exposure, Mar. Environ. Res. 101 (2014) 81–90, https://doi.org/10.1016/j.marenvres.2014.09.004.

[14] R. Trevisan, M. Arl, C.L. Sacchet, C.S. Engel, N.M. Danielli, D.F. Mello, C. Brocardo, A.F. Maris, A.L. Dafre, Antioxidant deficit in gills of Pacific oyster (*Crassostrea gigas*) exposed to chlorodinitrobenzene increases menadione toxicity, Aquat. Toxicol. 108 (2012), https://doi.org/10.1016/j.aquatox.2011.09.023.

[15] R. Trevisan, G. Delapedra, D.F. Mello, M. Arl, E.C. Schmidt, F. Meder, M. Monopoli, E. Cargnin-Ferreira, Z.L. Bouzon, A.S. Fisher, D. Sheehan, A.L. Dafre, Gills are an initial target of zinc oxide nanoparticles in oysters *Crassostrea gigas*, leading to mitochondrial disruption and oxidative stress, Aquat. Toxicol. 153 (2014) 27–38, https://doi.org/10.1016/j.aquatox.2014.03.018.

[16] D.F. Mello, L.A. de O. Proença, M.A. Barracco, Comparative study of various immune parameters in three bivalve species during a natural bloom of *Dinophysis acuminata* in Santa Catarina Island, Brazil, Toxins 2 (2010) 1166–1178, https://doi.org/10.3390/toxins2051166.

[17] M. Auffret, N. Mujdzic, C. Corporeau, D. Moraga, Xenobiotic-induced immunomodulation in the European flat oyster, *Ostrea edulis*, Mar. Environ. Res. 54 (2002) 585–589, https://doi.org/10.1016/S0141-1136(02)00120-4.

[18] N. Weng, J. Meng, S. Huo, F. Wu, W.X. Wang, Hemocytes of bivalve mollusks as cellular models in toxicological studies of metals and metal-based nanomaterials, Environ. Pollut. 312 (2022) 120082, https://doi.org/10.1016/J.ENVPOL.2022.120082.

[19] A. Katsumiti, A.J. Thorley, I. Arostegui, P. Reip, E. Valsami-Jones, T.D. Tetley, M. P. Cajaraville, Cytotoxicity and cellular mechanisms of toxicity of CuO NPs in mussel cells *in vitro* and comparative sensitivity with human cells, Toxicol. Vitro 48 (2018) 146–158, https://doi.org/10.1016/J.TIV.2018.01.013.

[20] D.F. Mello, E.S. de Oliveira, R.C. Vieira, E. Simoes, R. Trevisan, A.L. Dafre, M. A. Barracco, Cellular and transcriptional responses of *Crassostrea gigas* hemocytes exposed in vitro to Brevetoxin (PbTx-2), Marine Drugs 2012 10 (2012) 583–597, https://doi.org/10.3390/MD10030583. Pages 583-597 10.

[21] D.F. Mello, M. Arl, R. Trevisan, A.L. Dafre, How important are glutathione and thiol reductases to oyster hemocyte function? Fish Shellfish Immunol. 46 (2015) 566–572, https://doi.org/10.1016/j.fsi.2015.07.017.

[22] J.R. Meadows, J.C. DeWitt, A.A. Rooney, Ecoimmunotoxicology-An overview, in: Comprehensive Toxicology, third ed., Elsevier, 2018, pp. 886–891, https://doi.org/10.1016/B978-0-12-801238-3.64244-7.

[23] R. Trevisan, D.F. Mello, Redox control of antioxidants, metabolism, immunity, and development at the core of stress adaptation of the oyster *Crassostrea gigas* to the dynamic interstitial environment, Free Radic. Biol. Med. 210 (2024) 85–106, https://doi.org/10.1016/J.FREERADRIAMED.2023.11.003.

[24] L. Donaghy, A.K. Volety, Functional and metabolic characterization of hemocytes of the green mussel, *Perna viridis*: in vitro impacts of temperature, Fish Shellfish Immunol. 31 (2011) 808–814, https://doi.org/10.1016/J.FSI.2011.07.018.

[25] P. Moreau, T. Burgeot, T. Renault, Pacific oyster (*Crassostrea gigas*) hemocyte are not affected by a mixture of pesticides in short-2m in vitro assays, Environ. Sci. Pollut. Control Ser. 21 (2014) 4940–4949, https://doi.org/10.1007/S11356-013-1931-3/METRICS.

[26] C. Roman, P. Mahé, O. Latchere, C. Catrouillet, J. Gigault, I. Métais, A. Châtel, Effect of size continuum from nanoplastics to microplastics on marine mussel *Mytilus edulis*: Comparison in vitro/in vivo exposure scenarios, Comp. Biochem. Physiol. C Toxicol. Pharmacol. 264 (2023) 109512, https://doi.org/10.1016/J.CBCP.2022.109512.

[27] A. Barrick, C. Guillet, C. Mouneyrac, A. Châtel, Investigating the establishment of primary cultures of hemocytes from *Mytilus edulis*, Cytotechnology 70 (2018) 1205–1220, https://doi.org/10.1007/S10616-018-0212-X/METRICS.

[28] R. Ladhar-Chaabouni, A. Hamza-Chaffai, The cell cultures and the use of hemocytes from marine molluscs for ecotoxicology assessment, Cytotechnology 68 (2015) 1669, https://doi.org/10.1007/S10616-015-9932-3.

[29] L. Canesi, C. Ciacci, L.C. Lorusso, M. Betti, T. Guarnieri, S. Tavolari, G. Gallo, Immunomodulation by 17beta-estradiol in bivalve hemocytes, Am. J. Physiol. Regul. Integr. Comp. Physiol. 291 (2006) R664–R673, https://doi.org/10.1152/ajpregu.0019506.2006.

[30] A. Ji, X. Li, S. Fang, Z. Qin, C. Bai, C. Wang, Z. Zhang, Primary culture of Zhikong scallop *Chlamys farreri* hemocytes as an *in vitro* model for studying host-pathogen interactions, Dis. Aquat. Org. 125 (2017) 217–226, https://doi.org/10.3354/DAO03145.

[31] R. Ladhar-Chaabouni, W. Ayadi, E. Sahli, R. Mokdad-Gargouri, Establishment of primary cell culture of *Ruditapes decussatus* haemocytes for metal toxicity assessment, In Vitro Cell. Dev. Biol. Anim. 57 (2021) 477–484, https://doi.org/10.1007/S11626-021-00561-X.

[32] A. Cao, L. Mercado, J.I. Ramos-Martinez, R. Barcia, Primary cultures of hemocytes from *Mytilus galloprovincialis* Lmk.: expression of IL-2R subunit, Aquaculture 216 (2003) 1–8, https://doi.org/10.1016/S0044-8486(02)00140-0.

[33] R.W.A. Potts, A.P. Gutierrez, Y. Cortés-Araya, R.D. Houston, T.P. Bean, Developments in marine invertebrate primary culture reveal novel cell morphologies in the model bivalve *Crassostrea gigas*, PeerJ 2020 (2020) e9180, https://doi.org/10.7717/PEERJ.9180/SUPP15.

[34] S. Balakrishnan, A.K.M. Sajeevan, S.C. Parvathi, I.S. Bright Singh, J. Puthumana, An optimized protocol for routine development of cell culture from adult oyster, *Crassostrea madrasensis*, Cell Biol. Int. 48 (2024) 923–938, https://doi.org/10.1002/CBIN.12159.

[35] B. Petton, D. Destoumieux-Garzón, F. Pernet, E. Toulza, J. de Lorgeril, L. Degremont, G. Mitta, The Pacific oyster mortality syndrome, a polymicrobial and multifactorial disease: state of knowledge and future directions, Front. Immunol. 0 (2021) 52, https://doi.org/10.3389/FIMMU.2021.630343.

[36] M. Delaporte, P. Soudant, J. Moal, E. Giudicelli, C. Lambert, C. Séguineau, J. F. Samain, Impact of 20:4n-6 supplementation on the fatty acid composition and hemocyte parameters of the Pacific oyster *crassostrea gigas*, Lipids 41 (2006) 567–576, https://doi.org/10.1007/S11745-006-5006-9.

[37] C. Lambert, P. Soudant, G. Choquet, C. Paillard, Measurement of *Crassostrea gigas* hemocyte oxidative metabolism by flow cytometry and the inhibiting capacity of pathogenic vibrios, Fish Shellfish Immunol. 15 (2003) 225–240, https://doi.org/10.1016/S1050-4648(02)00160-2.

[38] M. Delaporte, P. Soudant, J. Moal, C. Lambert, C. Quéré, P. Miner, G. Choquet, C. Paillard, J.F. Samain, Effect of a mono-specific algal diet on immune functions in two bivalve species - *crassostrea gigas* and *Ruditapes philippinarum*, J. Exp. Biol. 206 (2003) 3053–3064, https://doi.org/10.1242/JEB.00518.

[39] S. Balakrishnan, I.S.B. Singh, J. Puthumana, Status in molluscan cell line development in last one decade (2010-2020): impediments and way forward, Cytotechnology 74 (2022) 433, https://doi.org/10.1007/S10616-022-00539-X.

[40] C. Pan, C. Kumar, S. Bohl, U. Klingmüeller, M. Mann, Comparative proteomic phenotyping of cell lines and primary cells to assess preservation of cell type-specific functions, Mol. Cell. Proteomics 8 (2009) 443–450, https://doi.org/10.1074/MCP.M800258-MCP200.
[41] S.E. Burdall, A.M. Hanby, M.R.J. Lansdown, V. Speirs, Breast cancer cell lines: friend or foe? Breast Cancer Res. 5 (2003) 89-95, https://doi.org/10.1186/BCR577/FIGURES/2.

[42] E. Mialhe, V. Boulo, H. Grizel, Bivalve Mollusc Cell Culture, vol. 18, American Fisheries Society Special Publication, 1988, pp. 311-315.

[43] R. Ladhar-Chaabouni, A. Hamza-Chaffai, The cell cultures and the use of haemocytes from marine molluscs for ecotoxicology assessment, Cytotechnology 68 (2016) 1669, https://doi.org/10.1007/S10616-015-9932-3.

[44] P. Brousseau, J. Pellerin, Y. Morin, D. Cyr, B. Blakley, H. Boermans, M. Fournier, Flow cytometry as a tool to monitor the disturbance of phagocytosis in the clam *Mya arenaria* hemocytes following in vitro exposure to heavy metals, Toxicology 142 (1999) 145-156, https://doi.org/10.1016/S0300-483X(99)00165-1.

[45] B. Morga, N. Faury, S. Guesdon, B. Chollet, T. Renault, Haemocytes from *Crassostrea gigas* and OsHV-1: a promising *in vitro* system to study host/virus interactions, J. Invertebr. Pathol. 150 (2017) 45-53, https://doi.org/10.1016/J.JIP.2017.09.007.

[46] B. Gagnaire, H. Thomas-Guyon, T. Burgeot, T. Renault, Pollutant effects on Pacific oyster, *Crassostrea gigas* (Thunberg), hemocytes: screening of 23 molecules using flow cytometry, Cell Biol. Toxicol. 22 (2006) 1-14, https://doi.org/10.1007/S10565-006-0011-6/METRICS.

[47] S.N. Chen, C.M. Wen, Establishment of cell lines derived from oyster, *Crassostrea gigas* Thunberg and hard clam, *Metereut lusoria* roding, Methods Cell Sci. 21 (1999) 183-192, https://doi.org/10.1023/A:1009829807954/METRICS.

[48] M.V. Golikov, V.T. Valuev-Elliston, O.A. Smirnova, A.V. Ivanov, Physiological media in studies of cell metabolism, Mol. Biol. 56 (2022) 629-637, https://doi.org/10.1134/S0026893322050077/METRICS.

[49] P. Schmitt, R.D. Rosa, M. Duperthuy, J. de Lorgeril, E. Bachère, D. Destoumieux-Garzón, The antimicrobial defense of the Pacific oyster, *Crassostrea gigas*. How diversity may compensate for scarcity in the regulation of resident/pathogenic microflora, Front. Microbiol. 3 (2012) 23326, https://doi.org/10.3389/FMICB.2012.00160/BIBTEX.

[50] Z. Liu, L. Wang, Z. Zhou, Y. Sun, M. Wang, H. Wang, Z. Hou, D. Gao, Q. Gao, L. Song, The simple neuroendocrine-immune regulatory network in oyster *Crassostrea gigas* mediates complex functions, Scientific Reports 2016 6 (1 6) (2016) 1-13, https://doi.org/10.1038/srep26396.

[51] L. Canesi, G. Gallo, M. Gavioli, C. Pruzzo, Bacteria-hemocyte interactions and phagocytosis in marine bivalves, Microsc. Res. Tech. 57 (2002) 469-476, https://doi.org/10.1002/JEMT.10100.

[52] N. Itoh, G.Q. Xue, K.L. Schey, Y. Li, R.K. Cooper, J.F. La Peyre, Characterization of the major plasma protein of the eastern oyster, *Crassostrea virginica*, and a proposed role in host defense, Comp. Biochem. Physiol. B Biochem. Mol. Biol. 158 (2011) 9-22, https://doi.org/10.1016/j.CBPB.2010.06.006.

[53] H.C. Song, C.Y. Xie, Q. Kong, L. Wei, X.T. Wang, Daylight ultraviolet B radiation ruptured the cell membrane, promoted nucleotide metabolism and inhibited energy metabolism in the plasma of Pacific oyster, Sci. Total Environ. 862 (2023), https://doi.org/10.1016/j.SCITOTENV.2022.160729.

[54] G. Udayan, M.E. Giordano, P. Pagliara, M.G. Lionetto, Motility of *Mytilus galloprovincialis* hemocytes: sensitivity to paracetamol in vitro exposure, Aquat. Toxicol. 265 (2023) 106779, https://doi.org/10.1016/j.AQUATOX.2023.106779.

[55] H. Gendre, M. Palos Ladeiro, A. Geffard, A. Poret, F. Le Foll, Y. Ben Cheikh, Modulation of haemocyte motility by chemical and biological stresses in *Mytilus edulis* and *Dreissena polymorpha*, Fish Shellfish Immunol. 139 (2023) 108919, https://doi.org/10.1016/j.FSI.2023.108919.

[56] M.G. McCormick-Ray, T. Howard, Morphology and mobility of oyster hemocytes: evidence for seasonal variations, J. Invertebr. Pathol. 58 (1991) 219-230, https://doi.org/10.1016/0022-2011(91)90066-Y.

[57] K.G. Takahashi, N. Izumi-Nakajima, K. Mori, Unique phagocytic properties of hemocytes of Pacific oyster *Crassostrea gigas* against yeast and yeast cell-wall derivatives, Fish Shellfish Immunol. 70 (2017) 575-582, https://doi.org/10.1016/J.FSI.2017.09.027.

[58] L. Canesi, G. Gallo, M. Gavioli, C. Pruzzo, Bacteria-hemocyte interactions and phagocytosis in marine bivalves, Microsc. Res. Tech. 57 (2002) 469-476, https://doi.org/10.1002/jemt.10100.

[59] S. Jiang, Z. Jia, T. Zhang, L. Wang, L. Qiu, J. Sun, L. Song, Functional characterisation of phagocytes in the Pacific oyster *Crassostrea gigas*, PeerJ 2016 (2016) e2590, https://doi.org/10.7717/PEERJ.2590/SUPP-1.

[60] L. Wang, X. Song, L. Song, The oyster immunity, Dev. Comp. Immunol. 80 (2018) 99-118, https://doi.org/10.1016/j.DCI.2017.05.025.

[61] L. Donaghy, C. Lambert, K.S. Choi, P. Soudant, Hemocytes of the carpet shell clam (*Ruditapes decussatus*) and the Manila clam (*Ruditapes philippinarum*): current knowledge and future prospects, Aquaculture 297 (2009) 10-24, https://doi.org/10.1016/J.AQUACULTURE.2009.09.003.

[62] M. Prado-Alvarez, A. Romero, P. Balseiro, S. Dios, B. Novoa, A. Figueras, Morphological characterization and functional immune response of the carpet shell clam (*Ruditapes decussatus*) haemocytes after bacterial stimulation, Fish Shellfish Immunol. 32 (2012) 69-78, https://doi.org/10.1016/j.FSI.2011.10.019.

[63] S. Li, Y. Liu, C. Liu, J. Huang, G. Zheng, L. Xie, R. Zhang, Morphology and classification of hemocytes in *Pinctada fucata* and their responses to ocean acidification and warming, Fish Shellfish Immunol. 45 (2015) 194-202, https://doi.org/10.1016/j.FSI.2015.04.006.

[64] L. Donaghy, E. Kraffe, N. Le Goïc, C. Lambert, A.K. Volety, P. Soudant, Reactive oxygen species in unstimulated hemocytes of the pacific oyster *Crassostrea gigas*: a mitochondrial involvement, PLoS One 7 (2012) e46594, https://doi.org/10.1371/journal.pone.0046594.

[65] L. Donaghy, S. Artigaud, R. Sussarell, C. Lambert, N. Le Goïc, H. Hégaret, P. Soudant, Tolerance of bivalve mollusc hemocytes to variable oxygen availability: a mitochondrial origin? Aquat. Living Resour. 26 (2013) 257-261, https://doi.org/10.1051/ALR/2013054.

[66] D.L. Hoffman, P.S. Brookes, Oxygen sensitivity of mitochondrial reactive oxygen species generation depends on metabolic conditions, J. Biol. Chem. 284 (2009) 16236-16245, https://doi.org/10.1074/jbc.M809512200.

[67] N.M. Danielli, R. Trevisan, D.F. Mello, K. Fischer, V.S. Deconto, D. da Silva Acosta, A. Bianchini, A.C.D. Bainy, A.L. Dafre, Upregulating Nrf2-dependent antioxidant defenses in Pacific oysters *Crassostrea gigas*: investigating the Nrf2/Keap1 pathway in bivalves, Comp. Biochem. Physiol. C Toxicol. Pharmacol. 195 (2017) 16-26, https://doi.org/10.1016/j.CBPC.2017.02.004.

[68] N.M. Danielli, R. Trevisan, D.F. Mello, K. Fischer, V.S. Deconto, A. Bianchini, A.C. D. Bainy, A.L. Dafre, Contrasting effects of a classic Nrf2 activator, tert-butylhydroquinone, on the glutathione-related antioxidant defenses in Pacific oysters, *Crassostrea gigas*, Mar. Environ. Res. 130 (2017) 142-149, https://doi.org/10.1016/J.MARENVRES.2017.07.020.

[69] D.F. Mello, C. Bergemann, K. Fisher, R. Chitrakar, S. Bijwadia, Y. Wang, A. Caldwell, L.R. Baugh, J.N. Meyer, Rotenone modulates *C. elegans* immunometabolism and pathogen susceptibility, Front. Immunol. (2022), https://doi.org/10.3389/fimmu.2022.840272.

[70] M.P. Murphy, H. Bayir, V. Belousov, C.J. Chang, K.J.A. Davies, M.J. Davies, T. P. Dick, T. Finkel, H.J. Forman, Y. Janssen-Heininger, D. Gems, V.E. Kagan, B. Kalyanaraman, N.G. Larsson, G.L. Milne, T. Nyström, H.E. Poulsen, R. Radi, H. Van Remmen, P.T. Schumacker, P.J. Thornalley, S. Toyokuni, C.C. Winterbourn, H. Yin, B. Halliwell, Guidelines for measuring reactive oxygen species and oxidative damage in cells and *in vivo*, Nature Metabolism 2022 4 (6 4) (2022) 651-662, https://doi.org/10.1038/s42255-022-00591-z.

[71] B.M. Polster, D.G. Nicholls, S.X. Ge, B.A. Roelofs, Use of potentiometric fluorophores in the measurement of mitochondrial reactive oxygen species, Methods Enzymol. 547 (2014) 225-250, https://doi.org/10.1016/B978-0-12-801415-8.00013-8.

[72] D.F. Mello, L.L. Maurer, I.T. Ryde, D.H. Song, S.M. Marinakos, J.N. Meyer, In vivo effects of silver nanoparticles on development, behavior and mitochondrial function are altered by genetic defects in mitochondrial dynamics, Environ. Sci. Technol. 56 (2) (2022) 1113-1124, https://doi.org/10.1021/acs.est.1c05915.

[73] T.C. Walther, R.V. Farese, Lipid droplets and cellular lipid metabolism, Annu. Rev. Biochem. 81 (2012) 687, https://doi.org/10.1146/ANNUREV-BIOCHEM-061009-102430.

[74] G.B. Gordon, M.A. Barcza, M.E. Bush, Lipid accumulation in hypoxic tissue culture cells, Am. J. Pathol. 88 (1977) 663-678. http://europepmc.org/articles/PMC2032384. (Accessed 20 May 2024).

[75] P.G. Beninger, M. Le Pennec, Scallop structure and function, Dev. Aquacult. Fish. Sci. 40 (2016) 85-159, https://doi.org/10.1016/B978-0-444-62710-0.00003-1.

[76] R.J. Pollero, G. Huca, R.R. Brenner, Role of hemocytes and plasma on lipid transport in the freshwater mollusc *Diplonod delodontus*, Comp. Biochem. Physiol. A Physiol. 82 (1985) 339-343, https://doi.org/10.1016/0300-9629(85)90865-5.

[77] A. Rolton, N.L.C. Ragg, Green-lipped mussel (*Perna canaliculus*) hemocytes: a flow cytometric study of sampling effects, sub-populations and immune-related functions, Fish Shellfish Immunol. 103 (2020) 181-189, https://doi.org/10.1016/J.FSI.2020.05.019.

[78] J. Vignier, M. Reardon, M. Exton, L. Delisle, A. Rolton, E. Malpot, M. Scholtens, M. Welford, L. Zamora, N. Delorme, B. Dunphy, S. Adams, Comparative performance of selected triploid oysters *Crassostrea (Magallana) gigas*, produced by chemical induction and mated triploid techniques, to their diploid counterparts, Aquaculture 596 (2025) 741894, https://doi.org/10.1016/J.AQUACULTURE.2024.741894.

[79] K. Brokordt, Y. Defranchi, I. Espósito, C. Cárcamo, P. Schmitt, L. Mercado, E. De la Fuente-Ortega, G.A. Rivera-Ingraham, Reproduction immunity trade-off in a mollusk: hemocyte energy metabolism underlies cellular and molecular immune responses, Front. Physiol. 10 (2019) 423209, https://doi.org/10.3389/FPHYS.2019.00077/BIBTEX.

[80] M.B. Duchemin, M. Fournier, M. Auffret, Seasonal variations of immune parameters in diploid and triploid Pacific oysters, *Crassostrea gigas* (Thunberg), Aquaculture 264 (2007) 73-81, https://doi.org/10.1016/J.AQUACULTURE.2006.12.030.

[81] B. Gagnaire, P. Soletchnik, P. Madec, P. Geairon, O. Le Moine, T. Renault, Diploid and triploid Pacific oysters, *Crassostrea gigas* (Thunberg), reared at two heights above sediment in Marennes-Oleron Basin, France: difference in mortality, sexual maturation and hemocyte parameters, Aquaculture 254 (2006) 606-616, https://doi.org/10.1016/J.AQUACULTURE.2005.10.008.