# Experimental ingestion of fluorescent microplastics by pacific oysters, Crassostrea gigas, and their effects on the behaviour and development at early stages

Arno Bringer, Jérôme Cachot, Grégoire Prunier, Emmanuel Dubillot, Christelle Clérandeau, Hélène Thomas

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## To cite this version:

Arno Bringer, Jérôme Cachot, Grégoire Prunier, Emmanuel Dubillot, Christelle Clérandeau, et al. Experimental ingestion of fluorescent microplastics by pacific oysters, Crassostrea gigas, and their effects on the behaviour and development at early stages. Chemosphere, 2020, pp.126793. 10.1016/j.chemosphere.2020.126793. hal-02547791

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Submitted on 5 Aug 2025

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Experimental ingestion of fluorescent microplastics by Pacific oysters, *Crassostrea gigas*, and
their effects on the behaviour and development at early stages

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Arno Bringer¹, Jérôme Cachot², Grégoire Prunier¹, Emmanuel Dubillot¹,

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Christelle Clérandeau², Hélène Thomas¹

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¹ Littoral Environnement et Sociétés (LIENSs), UMR 7266, CNRS-Université de La Rochelle, 2 rue

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Olympe de Gouges, F-17042 La Rochelle Cedex 01, France.

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² Université de Bordeaux, Laboratoire Environnements et Paléoenvironnements Océaniques et
Continentaux (EPOC), UMR 5805, F-33600 Pessac, France.

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**Keywords** Microplastics; oyster; D-larvae; swimming behaviour; ingestion; malformations.

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ABSTRACT

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Plastics are persistent synthetic polymers that accumulate as waste in the marine
environment. Microplastics (MPs, <5 mm) can be found either as microbeads in body care and
some industrial products or as plastic debris through degradation. Plastic microbeads (1-5 µm,
fluorescent, Cospheric) were used to characterise the MP ingestion and determine their
potential harmful effects on both the swimming behaviour and development of oyster D-larvae
(Crassostrea gigas). For 24 hours, embryos were first exposed to MPs at a temperature of 24 °C.

In addition, 3 day-old D-larvae were exposed to the same temperature for 1, 3 and 5 hours.

Three concentrations of MPs were used: 0.1, 1 and 10 mg MP. L⁻¹. After a 24-hour period of
embryonic exposure, we noted that MP agglomerates were stuck to the D-larvae coat and locomotor eyelashes. We also observed a significant increase in severe malformations and developmental arrests for larvae exposed to MPs ranging from 1 mg MP. L⁻¹. In terms of swimming behaviour, the maximum speed recorded was lower for larvae exposed at 0.1 and 1 mg MP. L⁻¹. After an acute exposure to MPs, particles were found in the digestive tract of 3 dpf (days post fertilization) D-larvae. After one-hour exposure, the concentrations tested (0.1, 1 and 10 mg MP. L⁻¹) resulted in respectively 38%, 86% and 98%. The larvae swimming behaviour was recorded and analysed. Unlike the results observed at the embryo-larval stage, 3-dpf larvae showed significant impacts with no dose-response effect.

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ABSTRACT GRAPHIC

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# 1. INTRODUCTION

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Microplastic particles (0.1 µm - 5 mm) are the most numerically abundant form of solid waste on Earth (Ericksen et al., 2014). In 2017, the world's plastic production nearly reached 350 million tons, including more than 64 million tons in Europe (PlasticsEurope, 2018). Each year, approximately 4.8 to 12.7 million tons end in the oceans (Jambeck et al., 2015). Although there are significant regional differences in plastic use and disposal (Kershaw et al., 2011), at a worldwide level, the most used polymers are: polyethylene (PE), polypropylene (PP), polyethylene terephthalate (PET), polyvinyl chloride (PVC), polystyrene (PS) and polyamide (PA) (Andrady and Neal 2009, PlasticsEurope 2010).

Microplastics are plastic particles that are smaller than 5 mm in diameter (Arthur et al., 2009). The presence of MPs has been reported in almost all studied habitats around the globe (Ivar do Sul and Costa, 2014; Andrady et al., 2011; Law and Thompson, 2014). In 2013, the average concentration of MPs in the Northeast Atlantic was of 2.46 MP.m⁻³. 89% of these MPs were smaller than 5 mm (Lusher et al., 2014). Microplastics can either be suspended in the water column (Lattin et al., 2004), buoy at surface water (Moore et al., 2001; Morét-Ferguson et al., 2010) or accumulate in sediments (Brown et al., 2011) in coastal waters (Ng and Obbard, 2006), rivers (Sadri and Thompson, 2014), estuaries (Browne et al., 2010) and ocean gyres (Maximenko et al., 2012). In aquatic ecosystems, the presence of plastics of different sizes and forms could negatively impact all levels of the marine food web, including the zooplankton one.

For instance, marine zooplankton (copepods, decapods, bivalve larvae, etc.) can ingest MPs directly or by trophic transfer (Lee et al., 2013; Setala et al., 2014; Cole et al., 2013; Cole et al., 2013).
2015). MPs can also be transferred to higher trophic levels via mesozooplankton (Setala et al., 2014 and Beiras et al., 2018). In addition, several studies documented the MP effects on the zooplankton's feeding behaviour, growth, development, reproduction and lifespan (Cole et al., 2015; Botterell et al., 2019).

Our study supports the idea that bivalves, including the Pacific oyster, *Crassostrea gigas*, are promising sentinel species to study the impacts of small MPs (<200 µm) (Bonanno et al., 2018).

The embryo-larval stage of oysters is frequently used as a bioassay in ecotoxicology, as this life stage is easy to obtain and is very sensitive to environmental pollutants (His et al., 1999; Mai et al., 2012; Mottier et al., 2013 and Tallec et al., 2018). Oyster larvae can ingest micro and nanoplastics in the early stages (3-24 dpf) of their development (Cole et al., 2015). In the work carried out by Sussarellu et al., 2016, adult oysters *C. gigas* that were exposed to 2-6 µm polystyrene microplastics were used as oyster genitors. The authors then studied the MP impacts on larval malformations and growth and concluded to noticeable effects on larvae.

Through their work, Bringer et al. 2020, demonstrated that polyethylene MPs had detrimental impact on the *C. gigas* swimming behaviour and larval development.

In this study, the effects of 1-5 µm MPs on two early stages of oyster development were assessed: the embryo-larval stage at 24 hours and D-larvae stage at 72 hours. Two batches of larvae were used to test four concentrations (0, 0.1, 1 and 10 mg MP. L⁻¹). In the first phase, embryo larvae were exposed to MPs. Then, the MP effects on the D-larvae (24 hour-old) malformations, developmental arrests and swimming behaviour were analysed. In the second phase, D-larvae were raised for 3 days – e.g. a stage with exogenous feeding (His et al., 1988) – and were then exposed to MPs at different exposure time intervals (1, 3 and 5 hours).
There are several advantages in using fluorescent plastic microspheres. First, commercial MPs benefit from homogeneous sizes and shapes. Second, many plastic particles can be purchased with embedded fluorophores or pigments allowing easy visualisation in biodeposits and tissues (Ward et al., 2019). For both experiments, the D-larvae swimming behaviour (including the maximum speed and trajectories) was analysed. Our results provided relevant information on both the development and swimming behaviour of two different embryo-larval stages after acute exposure to MPs (1-5 µm).

## 2. MATERIAL AND METHODS

### 2.1 Animal collection

Mature oysters (*Crassostrea gigas*, Bayne *et al.*, 2017; Bayne *et al.*, 2019) came from a commercial hatchery specialised in the year-round production of mature oysters (France Naissain, France). During the September-October 2018 period, the oysters were kept at approximately 4 °C for two days (which enabled us to have better quality laying eggs for the first tests to be carried out) and acclimatised for 2 hours in filtered sea water (FSW) before being fertilised in laboratory conditions.

### 2.2 Seawater

Seawater was collected from the French island of Île de Ré (South West of France). After sampling, seawater was filtered using a membrane filter of 20 µm, 5 µm, 1 µm and 0.2 µm and a pump generating UVs to eliminate debris and microorganisms. FSW was stored at 4 °C with
continuous bubbling and was used within 7 days. A few hours before the experiment, FSW was filtered again at 0.2 µm.

## 2.3 Gonad retrieval and fertilisation

Oyster genitors were spawn by thermal stimulation (alternating immersion in 18 °C and 28 °C seawater for 30 minutes) or by stripping the gonad (Mai *et al.*, 2013 and Gamain *et al.*, 2017).

At the time of the gamete expulsion, spawning males and females were individually isolated in beakers with 0.2 µm FSW. After analysing the gamete morphology under a microscope at 10X magnification (Nikon Eclipse), both eggs and sperm were selected for a single pairing. To eliminate faeces and debris, these were separately sieved through 50 µm and 100 µm meshes (Sefar Nitex). Sperm mobility was assessed and the number of eggs counted under a microscope at 20X magnification (Nikon Eclipse). Eggs were fertilised with sperm using a ratio of 1:10 (egg:sperm) and homogenised every five minutes with an agitator to prevent polyspermy.

Fertilisation success was observed under a microscope, and embryos were then counted and transferred. The first batch was exposed to MPs for 24 hours in order to observe their effects on the embryo-larval development. Embryos were transferred to 24-well microplates (Greiner Bio-One, Cellstar free of detectable DNase, RNase, human DNA and pyrogens. Content non-cytotoxic). The second batch was held in development for 72 hours before being exposed to MPs for 1, 3 and 5 hours. Oyster larvae began to feed on exogenous phytoplankton organisms from 48-72 hours, before that they fed on their vitelline reserves (His *et al.*, 1999). Embryos were transferred to 12-well microplates (Greiner Bio-One, Cellstar free of detectable DNase, RNase, human DNA and pyrogens. Content non-cytotoxic) for further development and
exposure. For both batches, approximately 350-400 fertilised eggs were transferred in each well containing 2 mL of FSW (Figure 1). These microplates were incubated in the dark at 24 °C to reach standard development (Robinson *et al.*, 1992 and AFNOR 2009). The first batch was incubated for 24 hours and the second for 72 hours (including additional stages of 1, 3 and 5 hours for ingestion).

<figure><img src="image_4.png" /><figcaption>**Figure 1.** Experimental design of both batches schematised, (1) embryonic exposure and analysis after a 24-hour development; (2) larval exposure after 3 dpf and analysis after a 1, 3 and 5-hour exposure.</figcaption></figure>

## 2.4 Microplastic preparation and exposure conditions

The MPs used were fluorescent (510 nm excitation) and ranged between 1 and 5 µm (FMG - Green Fluorescent Microspheres 1.3g/cc 1-5um, Proprietary Polymer, Cospheric, California, USA). For the experiment, MP particles were scattered in the water column.

MP solutions were prepared in FSW 0.2 µm and stored in the dark at 2 °C to limit microbial growth. For the first batch, 2 mL of MP solution was put in the microplates with the embryos
141 for a 24-hour exposure. For the second batch, the 3-dpf D-larvae were exposed to MP
142 microbeads for 1, 3 and 5 hours. 2 mL of MP solution and 50 µL of algae mixture (Roland *et al.*,
143 1988; 50.000 cells. mL⁻¹) were injected in 12-well microplates containing 2 mL of larvae (3 dpf)
144 suspended in FSW. A mix of *Isochrysis spp.* (*T.iso* strain) and *Skeletonema costatum* (5:1 ratio;
145 France Naissain, France; Cole *et al.*, 2015) was used for feeding and growth experiments. Algal
146 size, biovolume and population density were assessed using a Nikon Eclipse microscope (x20).
147 For both batches, three different concentrations were used: 0.1 mg MP. L⁻¹; 1 mg MP. L⁻¹ and 10
148 mg MP. L⁻¹ excluding control treatment.

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150 **2.5 Chemical analyses**

151 To carry out chemical analyses of 1-5 µm fluorescent MPs at 0.1; 1 and 10 mg MP. L⁻¹, each
152 solution was assayed using a flow cytometer (Attune Acoustic Focusing Cytometer). 2 mL
153 samples of MP solution used in embryo-larval exposure were vortexed (StarLab Vortex IR,
154 12,000 rpm for 20 sec) before going into a cytometer to homogenise the solution
155 (n=4/condition). A volume of 300 µl was sampled using the cytometer, which operated at a flow
156 rate of 500 µl.min⁻¹. Control analytical samples (n=12) contained 0.2 µm FSW (MPs excluded)
157 to remove naturally-present particles. The maximum threshold to detect particles was set at
158 10,000 events. Mili-Q water samples (n=12) were also assessed to check the relevance of the
159 detected events (<200 events). Then, a calibration was performed for MPs of different sizes
160 (HDPE MPs 4-6 µm MPP-635 XF, 11-13 µm MPP-635 G and 20-25 µm MPP-1241, density 0.96,
161 Micropowders Inc. USA) to select the relevant ones to the study (1-5 µm and fluorescent).

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163 **2.6 Swimming behaviour (maximum speed recorded and trajectories)**

164 Post-exposure (meaning after 24 hours for the first batch and respectively 1, 3 and 5 hours for
165 the second batch), 2-minute videos of D-larvae were recorded using a 24 °C thermoregulated
166 box (ZEISS Axio Observer Z1, x5) to determine their swimming behaviour (maximum speed and
167 trajectories, Bringer et al., 2020). Then, oyster D-larvae were fixed (1% buffered formaldehyde)
168 for abnormal larvae analysis. Formaldehyde was purchased from Sigma-Aldrich Chemical (St.
169 Quentin Fallavier, France). A video conversion freeware (VirtualDub, Windows) set the frame
170 rate at 4 FPS and converted it to AVI format. ImageJ (1,52a software) was used to analyse the
171 videos. Videos (.avi) were opened and converted first into grayscale images, and then into
172 binary images. For each video, a plugin was installed to deduce the swimming parameters of
173 the recorded D-larvae. As a result, each larva was provided with a tracking number to easily
174 identify them in the result file. It also included the maximum speed recorded (pixel. s⁻¹). In
175 addition, swimming trajectories were assessed using an image file. Consequently, three
176 different types of larval path were identified: (1) rectilinear, (2) circular and (3) motionless
177 (Bringer et al., 2020). The use of a graphic tablet (Wacom Bamboo Pen &Touch) and image
178 processing software (Photos, Windows 10) enabled swimming trajectories to be quantified and
179 characterised. The same larva can be detected several times, entering and exiting the visual
180 field (Gamain et al., 2019 and Bringer et al., 2020).

181 The video created to assess the swimming behaviour is available upon request (additional
182 data). The results displayed are expressed in mean ± SEM for the three replicates (three
183 different couples of oyster genitors). There are seven videos by couple, resulting in n=21
184 videos/treatment.
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## 2.7 Developmental abnormalities in D-larvae

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After having recorded the D-larvae swimming behaviour, 25 µL of 1% buffered formalin was added to each well. A biometric analysis of larvae exposed for 24 hours (n = 15) was carried out on the ImageJ software, using photos obtained by microscopy (ZEISS Axio Observer, Z1, x20).

The larvae were measured, by drawing a dummy line lengthwise (Talmage *et al.*, 2009 and Helm *et al.*, 2006). Then, the percentage of abnormal oyster larvae and developmental arrest was assessed (His *et al.*, 1999 and Quiniou *et al.*, 2005). Both phenomena were observed under an inverted microscope (Nikon eclipse, x10). This test required to have less than 20% of abnormal larvae in control conditions (absence of contamination). Three different couples were used and four replicates each were performed. Screenshots of the recorded videos were taken (VLC Media Player, Windows) to observe the contiguous MPs sticking on D-larvae (after a 24-hour period). The results displayed are expressed in mean ± SEM for the three replicates (three different couples of oyster genitors). Twelve wells for each condition were analysed using approximately 200 D-larvae in each well. This resulted in n=2,400 D-larvae analysed/treatment.

## 2.8 MP ingestion by 3-dpf D-larvae

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To determine the MP presence or absence in 3-dpf D-larvae and the percentage of larvae ingesting microbeads, a digital reconstruction of the wells was created by superimposing pictures (GFC fluorescence, ZEISS Axio Observer Z1, x5 objective). By zooming well by well, a count of larvae was carried out (Figure S1). Three replicates (three different couples of oyster genitors) and four wells of each were analysed, resulting in n=12 wells analyses/treatment.
The number of microspheres ingested by each larva was assessed through fluorescence intensity measurements. To do so, the ratio between the mean fluorescence intensity of each isolated bead (n=30) and the fluorescence intensity in the D-larvae digestive tract (n=6/treatment) was calculated. Not only the fluorescence intensity of the MP beads in the larval gut indicated if the particles were ingested or not (Figure S2) but it enabled to precisely determine where these were located in the digestive tract. For this experiment, the 3-dpf larvae were individually and independently tested for MP ingestion, meaning that larvae for each exposure time interval were different. Three replicates (three different couples of oyster genitors) were analysed, resulting in n=6 for each couple and n=18/treatment.

## 2.9 Statistics

The statistical analysis was conducted using R and graphs on Microsoft Excel, except the PCA-Biplot which was created on R Studio using the "FactoMineR" package. Homogeneity of variance (Levene's test) and normality of distribution (Shapiro-Wilk) were assessed. In order to compare the various treatments, an analysis of variance (ANOVA) using Tukey's post-hoc test was performed. For data not entering in the scope of a parametric test, a statistical analysis was carried out through the Kruskal-Wallis Test. Differences in tested concentration means were then assessed using the Kruskal Nemenyi Post-hoc test with the PMCMR package (the Tukey test equivalent for non-parametric data). When p<0.05, significance was tolerated. Data are written as mean ± SEM.

# 3. RESULTS
<table><thead><tr><th>Theoretical (mg MP.L<sup>-1</sup>)</th><th>Measured (MP.mL<sup>-1</sup>)</th><th>Dilution factor</th></tr></thead><tbody><tr><td>0.1</td><td>7 211±602</td><td>X</td></tr><tr><td>1</td><td>68 733±337</td><td>9.5</td></tr><tr><td>10</td><td>54 3528±42 646</td><td>7.9</td></tr></tbody></table>

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231 **Table 1**

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length at 10 mg MP. L⁻¹ was of 55.9±6.3 µm. These results differed from other conditions, particularly from control larvae which had a length of 73.6±6.7 µm (Table S3).

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**Figure 2.** (a) Maximum speed recorded, (b) swimming trajectories, (c) percentage of abnormal D-larvae and (d) developmental arrest of oyster D-larvae exposed for 24 hours to various concentrations (0; 0.1; 1; 10 mg MP. L⁻¹) of fluorescent MPs (1-5 µm). Values are expressed as mean ± SEM for the three replicates. A variation in letters indicates significant differences between concentrations (p<0.05, Kruskal-Wallis for swimming behaviour and ANOVA for abnormal and developmental arrest quantification).
Regarding the malformations and developmental arrests of oyster D-larvae (Figure 2.c & d), a dose-response effect to exposure concentrations was noted. An abnormal larval rate was observed at the two highest concentrations of 1 and 10 mg MP. L⁻¹, with respectively 35.2±2.2 % and 37.3±2.0% (Table S3). In contrast, malformed larvae in control conditions had a rate of 14.8±1.2%. In terms of developmental arrests, the highest rate was assessed at 10 mg MP. L⁻¹, with 15.3±1.6% (Table S3). The higher the MP concentration was, the higher the rate of developmental arrests. On these screenshots (Figure S4), MPs formed agglomerates. In addition, they stuck on the mantle and the locomotor eyelashes of the 24-hour D-larvae. On this picture (Figure S4, on the right), some MP agglomerates were larger than the larvae.

### 3.3 MP ingestion assessment and quantification in 3-dpf larvae at different time intervals
<figure><img src="image_9.png" /><figcaption>**Figure 3.** Microscopic pictures ZEISS Axio Observer Z1, magnification x20 (a) and microscopic zoomed pictures on 10 mg MP. L⁻¹, ZEISS Axio Observer Z1, magnification x60 (b) of 3-dpf oyster D-larvae exposed at different time intervals (1, 3 and 5 hours) and facing different concentrations of fluorescent MPs.</figcaption></figure>

The ingestion of fluorescent MPs by 3-dpf oyster D-larvae was assessed. Larvae ingested beads after 1, 3 and 5 hours of fluorescent MP exposure (Figure 3.a). We noted that the tested MPs tended to accumulate in the larvae digestive tract (Figure S2). However, there was no
difference between the exposure time intervals for a given concentration (Figure 3.b). When reconstructing the wells of microplates, we managed to quantify the number of D-larvae that ingested MPs and the ones who did not (Figure 3). A clear dose-response effect was observed. However, no impact was highlighted on the exposure time intervals (Figure 4). The larvae that were exposed to 10 mg MP. L⁻¹ were more likely to have ingested MPs (98%) than the ones exposed to 1 mg MP. L⁻¹ (84%) or 0.1 mg MP. L⁻¹ (47%).

<figure><img src="image_10.png" /><figcaption>**Figure 4.** MP ingestion by oyster D-larvae according to exposure time intervals (1, 3 and 5 hours) and MP concentrations (0; 0.1; 1; 10 mg MP. L⁻¹). Bars are expressed as mean values ± SEM for the three replicates. A variation in letters indicates significant differences (p<0.05, ANOVA).</figcaption></figure>

These results could be directly linked to the number of beads ingested by the oyster D-larvae and present in their digestive tract. Assessing the fluorescent content and intensity in larvae enabled to determine the number of beads ingested by each individual (Figure 5.a). Almost all
292 oyster larvae exposed to 10 mg MP. L⁻¹ ingested MPs (98%, red circle, Figure 5.a). The different

293 exposure time intervals were grouped in the graph as there was no significant difference for

294 this parameter. Larvae exposed to 0.1 mg MP.L⁻¹ ingested an average of 14.7±0.8 MP.larvae⁻¹

295 (grey circle, Figure 5.a), while the ones exposed to 1 mg MP.L⁻¹ (blue circle, Figure 5) and 10 mg

296 MP.L⁻¹ respectively ingested 35.7±4.1 and 141.8±5.2 MP.larvae⁻¹. There was no significant

297 difference between the three MP concentrations for the amount of microbeads ingested by the

298 D-larvae.
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**Figure 5.** Relationship between the number of oyster D-larvae that ingested MPs (%) and the mean number of microbeads ingested per larva (a) after 1, 3 and 5 hours of exposure (n=18). Asterisks indicate significant differences (Kruskal Wallis Test). Levels of significance: p<0.1="."; p<0.05="*"; p<0.01="**"; p<0.001="****" and (b) correlation plot (PCA-Biplot) between different variables: individuals, MPs ingested by larvae and percentage of larvae (%) presenting MPs in their digestive tract.

To correlate the 3 variables, a PCA-Biplot was created (Figure 5.b). Individuals who ingested high concentrations of MPs (fluorescence found in their digestive tract) were correlated to the percentage of larvae that ingested MPs. Groups of individuals (depending on their exposure conditions) were well distributed on the PCA. Moreover, "contribution" of variables highlighted that the number of MPs ingested by larvae was the variable that contributed the most to creating correlations.

## 3.4 Effects of MP exposure on 3-dpf larvae

For this experiment, D-larvae were bred for 3 dpf (72 hours). Then, they were exposed to MPs (1-5 µm) for 1, 3 and 5 hours. The effects of MP exposure to the larvae swimming behaviour was then analysed (Figure 6).
<figure><img src="image_13.png" /></figure>

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321 **Figure 6.** Maximum speed recorded and swimming trajectories of 3-dpf oyster D-larvae after a 1 (a), 3 (b) and 5-hour (c) exposure to fluorescent MPs at different concentrations (0; 0.1; 1; 10 mg MP. L⁻¹). Values are expressed as mean values ± SEM for the three replicates. A variation in letters indicates significant differences between different concentrations (p<0.05, Kruskal-Wallis test).

325
326 After a 1-hour exposure to different MP concentrations, the maximum speed recorded
327 increased significantly for exposed D-larvae (Figure 6.a) in comparison to control larvae.
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329 However, there are no differences observed in terms of MP concentrations. After 3 and 5 hours
330 of exposure (Figure 6.b and 6.c), the results were quite heterogeneous; no trend could be
331 defined. Regarding their swimming trajectories, control larvae had predominant circular
332 trajectories (Figure 6.a). After 3 hours, exposed larvae tended more towards circular
333 trajectories when increasing MP concentration, although this is not significant. After a 5-hour
334 exposure, we noted a significant difference (p>0.05) in swimming. Indeed, there was an
335 increase in motionless trajectories (p> 0.05). At this stage of development (72-hour D-larvae),
336 swimming trajectories seemed not to be as strongly impacted by MPs.

336
337 **4. DISCUSSION**
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For the whole study, we examined two different developmental stages of oysters exposed to 1-
339 5 µm fluorescent MPs: (1) embryos after being exposed for 24 hours and (2) 3-dpf D-larvae
340 after being exposed for 1, 3 and 5 hours.

341 First of all, after a 24-hour exposure, D-larvae exposed at the embryonic stage had MPs stuck
342 on their locomotor cilia and their coat. They did not show any sign of MP ingestion. When
343 embryos were directly in contact with fluorescent MPs, the (24-hour) larvae body length was
impacted. Exposed oyster D-larvae were smaller than unexposed ones. The consequences on larval size were previously documented by Sussarellu *et al.* (2016). When oyster broodstocks were exposed to 2 and 6 µm polystyrene beads, the larvae were smaller in size. Similarly, after being exposed to MPs (20 µm polystyrene beads), copepod eggs (*Calanus helgolandicus*) were smaller (Cole *et al.*, 2015). However, these biometric results and growth abnormalities might be due to developmental delays and malformations directly impacting the D-larvae size. As mentioned, exposed oyster larvae showed abnormalities and developmental arrests. Some studies highlighted that contaminants such as heavy metals and pesticides caused developmental defects (Mai *et al.*, 2012; Gamain *et al.*, 2017; Gamain *et al.*, 2016). We observed similar phenomena when species were exposed to 4-6 µm polyethylene MPs (Bringer *et al.*, 2020). In addition, a recent study has underlined that blue mussel larvae, *Mytilus edulis*, had malformations and developmental defects after being exposed to 100 nm and 2 µm polystyrene microbeads (Rist *et al.*, 2019). In this study, we noted a sharp increase in malformed larval rate from 1 mg MP. L⁻¹ and a dose-dependent increase in developmental arrests up to 15% at 10 mg MP. L⁻¹. These results echoed a previous study using 4-6 µm PE MPs (Bringer *et al.*, 2020). Our findings were consistent with the results of Lei *et al.* (2018), who demonstrated that MP toxicity was size-dependant rather than polymer-dependant. The study highlighted that the swimming behaviour of D-larvae, in particular their maximum speed recorded and trajectories, was impacted by MPs. Exposed larvae swam slower and had predominant circular swimming trajectories. Gamain *et al.* (2019) also reached similar conclusions when studying oyster larvae exposed to copper and pesticides. In addition, a reduced swimming speed was observed in oyster D-Larvae exposed to different sizes of PE MPs
366 (Bringer et al., 2020). According to our microscopic observations, MPs stuck on the larvae cilia.

367 Several studies documented that MPs tended to bind on aquatic species and disturb their

368 swimming behaviour. For instance, brine shrimps *Artemia franciscana* had MPs stuck to their

369 antennas and consequently an abnormal swimming behaviour (Bergami et al., 2016). MPs also

370 stuck on the tentacles of *Hydra attenuata*, which could impair feeding by restricting its ability to

371 move and capture prey (Murphy et al., 2018). The observations on larvae exposed (*H. attenuata*) for 24 hours underlined that MPs *quanta* (Alimi et al., 2018) were likely to impact

372 their swimming behaviour, mainly their speed and trajectories.

373

374 In a second phase, 3-dpf larvae were analysed and exposed to MPs at different time intervals

375 (1, 3 and 5 hours). In previous studies, authors noticed that oyster larvae could ingest and

376 accumulate MPs in their digestive tract (Jones and Jones, 1988; Cole et al., 2015). Capture

377 efficiency of small particles (ca. 1–6 µm) is species-specific and depends on the gill architecture

378 and structure of the latero frontal cilia or cirri (Silverman et al., 1995; Ward and Shumway 2004;

379 Rosa et al., 2017, 2018). We noted that the higher the MP concentration was, the more

380 particles the larvae ingested. However, no significant difference was observed in terms of

381 exposure times. This implies that D-larvae ingested MPs faster than they egested (Cole et al.,

382 2016). The correlations in the PCA-Biplot highlighted that the larvae ingested and stored large

383 quantities of MPs in their digestive tract when the experimental unit was loaded with particles.

384 A previous study demonstrated that oyster (*C. gigas*) and slipper limpet (*C. fornicata*) larvae

385 could deplete the nutrient stocks in their environment in no time (Blanchard et al., 2008).

386 MP ingestion and accumulation in the digestive tract of 3dpf D-larvae indicated an increase in

387 their maximum swimming speed under certain contamination conditions. MP ingestion altered
the maximum swimming speed and thus, to a lesser extent, the trajectories. However, this phenomenon was not observed for all contaminated conditions. One cannot clearly conclude in a trend in duration nor in a dose-response effect.

In this study, we noticed that MPs tended to agglomerate and stick to the locomotor eyelashes of the 24 hour-old larvae, which were not able to filter exogenous food and did not show MP ingestion. However, MPs were ingested by the 72-hour D-larvae (3 dpf). This impacted their swimming behaviour as larvae with MPs clung on their eyelashes showed behavioural abnormalities while larvae that had ingested MPs did not demonstrate behavioural differences.

This study underlined that the swimming behaviour and the development of the oyster were affected when exposed at embryonic stage to small PE MPs. For older larva exposed to MPs at 3 dpf, their swimming activity tended to increase when exposure time lasted up to 1 hour. The larvae did not show significant difference in their swimming trajectories after being exposed to MPs.

Another study demonstrated that egestion rate kinetic of 1-5 µm fluorescent MPs by oyster larvae was fairly rapid (< 3 h) and depended on the particle size (Ward et al., 2019). Indeed, one could investigate the larvae MP egestion and exclude the possibility of re-ingestion (Ris et al., 2019) by developing a relevant protocol.

## 5. CONCLUSIONS

In this study, MPs stuck on the locomotor cilia of the D-larvae exposed for a 24-hour period at the embryo-larval stage, modifying their maximum swimming speed and to, a lesser extent, their trajectories. We concluded that being exposed to small fluorescent MPs at the embryo-
larval stage was detrimental for D-larvae (C. gigas), causing developmental abnormalities, changes in their swimming behaviour (both speed and trajectories) as well as developmental arrests after a 24-hour period. The effects observed were dose-dependent.

In addition, 3-dpf exposed D-larvae ingested MPs, which were then observed in their digestive tracts. The higher the MP concentration was, the more MP particles the D-larvae tended to ingest. Regarding their swimming behaviour, we noted an increase in speed, especially for larvae exposed for a 1-hour period. The swimming trajectories did not seem to be impacted by MPs. Further research on *Crassostrea gigas* and other bivalve species, such as *Mytilus edulis*, is essential to build up on our knowledge. This would help establish recommendations on the ecological risks of MPs in coastal and estuarine ecosystems.

## SUPPORTING INFORMATION

Video: the oyster D-larvae swimming behaviour was recorded on a ZEISS microscope, both after a 24-hour exposure at the embryo-larval stage, after 3 dpf of development and a 3-hour exposure at 0.1 mg MP. L⁻¹.

## AUTHOR INFORMATION

### Corresponding author

Phone: +33 645299739 (France)

Email address: arno.bringer1@univ-lr.fr

## ACKNOWLEDGMENT
Arno Bringer received a PhD grant (Comité Régional de la Conchyliculture de la Charente-Maritime CRC17) as well as financial support from the Région Nouvelle Aquitaine and the Comité Départemental de la Charente Maritime (CD17) to develop his research. The AQUAECOs project (Amélioration de la Qualité Environnementale sur les zones Conchylicoles des Pertuis Charentais), a partnership with CRC17 and the AFB (French Biodiversity Agency) contributed to this study. Funds were also given by the University of La Rochelle, the University of Bordeaux and the Centre National de la Recherche Scientifique (France). We are grateful to Oriane Titeca for proofreading the language. We wish to thank our collaborators for their work and help.

440

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613 ADDITIONAL DATA

<figure><img src="image_19.png" /><figcaption>**Figure S1.** Counting method (superimposing pictures to reconstruct wells) to determine MP presence/absence in D-larvae using a zooming function.</figcaption></figure>
<figure><img src="image_20.png" /><figcaption>**Figure. S2.** Microscopic pictures (ZEISS Axio Observer Z1) of oyster-D larvae exposed to fluorescent MPs (1-5 µm). On the left picture: fluorescent microbead in the larva digestive tract (described by Jones and Jones, 1988). On the right picture: larva with agglomerates MP microbeads with exposure of 1 mg MP. L⁻¹.</figcaption></figure>

<table><thead><tr><td>Conditions</td><td>Body length</td><td>Abnormal</td><td>Developmental arrest</td><td>Maximum speed</td></tr></thead><tbody><tr><td>Control</td><td>73.6±6.7 <sup>a</sup></td><td>14.8±1.2 <sup>a</sup></td><td>4.1±0.4 <sup>a</sup></td><td>167.4±5.1 <sup>a</sup></td></tr><tr><td>0.1 mg MP.L<sup>-1</sup></td><td>60.3±0.7 <sup>b</sup></td><td>17.5±0.9 <sup>a</sup></td><td>6.8±0.6 <sup>ab</sup></td><td>148.5±3.7 <sup>b</sup></td></tr><tr><td>1 mg MP.L<sup>-1</sup></td><td>58.3±3.7 <sup>bc</sup></td><td>35.2±2.2 <sup>b</sup></td><td>8.5±0.7 <sup>b</sup></td><td>135.6±3.0 <sup>c</sup></td></tr><tr><td>10 mg MP.L<sup>-1</sup></td><td>55.9±6.3 <sup>c</sup></td><td>37.3±2.0 <sup>b</sup></td><td>15.3±1.6 <sup>b</sup></td><td>137.9±3.1 <sup>c</sup></td></tr></tbody></table>
<figure><img src="image_21.png" /><figcaption>**Figure S4.** Screenshots taken from the recorded videos (ZEISS Axio Observer Z1) of oyster D-larvae exposed to fluorescent MPs for 24 hours (1 mg MP. L⁻¹) at the embryonic stage. Presence of fluorescent MP as agglomerates close to the eyelashes and the larval mantle.</figcaption></figure>