# Microplastics in the insular marine environment of the Southwest Indian Ocean carry a microbiome including antimicrobial resistant (AMR) bacteria: A case study from Reunion Island

Loik Sababadichetty, Guillaume Miltgen, Bryan Vincent, François Guilhaumon, Véronique Lenoble, Margot Thibault, Sophie Bureau, Pablo Tortosa, Thierry Bouvier, Philippe Jourand

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Loik Sababadichetty, Guillaume Miltgen, Bryan Vincent, François Guilhaumon, Véronique Lenoble, et al. Microplastics in the insular marine environment of the Southwest Indian Ocean carry a microbiome including antimicrobial resistant (AMR) bacteria: A case study from Reunion Island. Marine Pollution Bulletin, 2024, 198, pp.115911. 10.1016/j.marpolbul.2023.115911. ird-04348345

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Distributed under a Creative Commons Attribution 4.0 International License
1 Title: Microplastics in the insular marine environment of the Southwest Indian Ocean carry a
2 microbiome including antimicrobial resistant (AMR) bacteria: a case study from Reunion Island.

3

4 Author names: Loik Sababadichetty¹,²a, Guillaume Miltgen²,³a, Bryan Vincent⁴a, François Guilhaumon⁵a, Veronique Lenoble⁶, Margot Thibault¹,⁷,⁸, Sophie Bureau¹, Pablo Tortosa³, Thierry Bouvier⁹ and Philippe Jourand⁵, *.

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8 Authors affiliations :

9 ¹Université de La Réunion, UMR ENTROPIE, 15 Avenue René Cassin, CS 92003, 97744, Saint Denis
10 Cedex 9, La Réunion, France.

11 ²CHU, Laboratoire de Bactériologie, CHU Félix Guyon, Allée des Topazes, 97400, Saint-Denis, La Réunion, France.

13 ³Université de La Réunion, UMR PIMIT Processus Infectieux en Milieu Insulaire Tropical, CNRS
14 9192, INSERM 1187, IRD 249. Plateforme de recherche CYROI, 2 rue Maxime Rivière, 97490 Ste
15 Clotilde, La Réunion, France

16 ⁴CIRAD, UMR040 LSTM, Campus Agro Environnemental Caraïbe, BP 214-97285, Cedex 2 le
17 Lamentin, Martinique, Antilles Françaises, France

18 ⁵IRD, UMR ENTROPIE, 15 Avenue René Cassin, CS 92003, 97744, Saint Denis Cedex 9, La Réunion, France.

20 ⁶Université de Toulon, Aix Marseille Université, CNRS, IRD, UMR MIO, 83 Toulon, France

21 ⁷The Ocean Cleanup, Rotterdam, The Netherlands

22 ⁸CNRS, Université Toulouse III, Laboratoire des Interactions Moléculaires et Réactivité Chimique et
23 Photochimique (IMRCP), UMR 5623, Toulouse, France

24 ⁹UMR MARBEC, Université Montpellier, CNRS, Ifremer, IRD, Montpellier, France

25 *Corresponding author : Philippe Jourand, IRD, UMR ENTROPIE, 15 Avenue René Cassin, CS 92003, 97744, Saint Denis Cedex 9, La Réunion, France. Tel: +262 0262 52 89 19; E-mail: philippe.jourand@ird.fr

28 ªThese authors contributed equally to this work.
29 Highlights

30 • Severe marine plastic pollution impacts Southwest Indian Ocean insular ecosystems

31 • Plastic debris from Southwest Indian Ocean host rich microbiomes

32 • Proteobacteria dominate such marine plastic microbiomes

33 • These debris carry a consequent culturable bacterial flora including potential pathogens

34 • AMR bacteria hitchhike on these plastics

35

36 Abstract

37 The increasing threats to ecosystems and humans from marine plastic pollution require a
38 comprehensive assessment. We present a plastisphere case study from Reunion Island, a remote
39 oceanic island located in the Southwest Indian Ocean, polluted by plastics. We characterized the
40 plastic pollution on the island's coastal waters, described the associated microbiome, explored viable
41 bacterial flora and the presence of antimicrobial resistant (AMR) bacteria. Reunion Island faces plastic
42 pollution with up to 10,000 items/km² in coastal water. These plastics host microbiomes dominated by
43 Proteobacteria (80%), including dominant genera such as *Psychrobacter*, *Photobacterium*,
44 *Pseudoalteromonas* and *Vibrio*. Culturable microbiomes reach 10⁷ CFU/g of microplastics, with
45 dominance of *Exiguobacterium* and *Pseudomonas*. Plastics also carry AMR bacteria including β-
46 lactam resistance. Thus, Southwest Indian Ocean islands are facing serious plastic pollution. This
47 pollution requires vigilant monitoring as it harbors a plastisphere including AMR, that threatens
48 pristine ecosystems and potentially human health through the marine food chain.

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50 Keywords: Reunion Island, plastic marine pollution, microbiome, Proteobacteria, AMR bacteria
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# 1. Introduction

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Marine pollution by plastic wastes and debris is an important source of anthropogenic contamination in the oceans (Thushari & Senevirathna, 2020). This pollution is increasingly seen as a major concern not only for the environment, i.e., contamination of pristine ecosystems with loss of biodiversity, but also for human health through contamination of marine trophic networks (Smith et al., 2018; Wright et al. 2020; Kumar et al., 2021). Plastics, when they enter the marine ecosystem, influence the marine food chain differently in different species due to factors such as size, shape, polymer composition and chemical additives (Tuuri and Laterme, 2023). Plastics contaminate marine life at many levels, from zooplankton (Cole et al., 2013) to bivalves (Van Cauwenberghe and Janssen, 2014), fishes (Li et al. 2021), reptiles such as turtles (Thibault et al., 2023) to marine mammals (Panti et al., 2019) and, at the end of the marine food chain, humans (Waring et al., 2018). For example, microplastics have been found in most marine organisms along trophic levels in various organs such as intestinal contents, gills or even flesh (Walkinshaw et al., 2020). In addition, the physical and chemical properties of plastic debris allow them to absorb numerous contaminants such as chemicals, metals and bacteria (Imran et al., 2019), acting as vectors for these contaminants when ingested by organisms. Therefore, bioaccumulation of plastics for higher trophic organisms has wider impacts on marine food webs with potential impacts on human health (Carbery et al., 2018).

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Plastics slowly degrade over time into smaller and smaller particles including those called "microplastics" (particle size between 0.1 - 5 mm) through weathering and physical processes as well as microbial activities (Galloway et al., 2017; Rummel et al., 2017; Jacquin et al., 2019). As a result, plastics remain present in the marine environment over very large time scales and accumulate, with an estimation of microplastic particles in 2014 reaching up to 50 trillion particles and weighing over 200,000 metric tons (van Sebille et al., 2015). Microplastic contamination of coastal and marine ecosystems reaches up to 140 particles/m³ in water and 8766 particles/m³ in sediments (Thushari & Senevirathna, 2020).

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These microplastics are durable, often floating substrates with physical and chemical properties that can have negative impacts on entire marine ecosystems over all bathymetric zones (Rochman, 2015; Masry et al., 2021). The physical traits of microplastics negatively impacting the
ecosystems and marine organisms are the plastic size, their shape, the polymer composition and the surface chemistry (Lambert et al., 2017). Moreover, the potential harm of microplastics in marine environments is linked to their ability to accumulate contaminants and transport toxic substances and pathogens (Imran et al., 2019; Chouchene et al., 2023). Microplastic substrates offer new habitats and very effective dispersal ways to microbial communities that can attach through the formation of biofilms and drift along oceanic currents (Oberbeckmann & Labrenz 2020). In addition, microorganisms' communities associated with plastic, the so-called "plastisphere" (Zettler et al., 2013), have been shown to be enriched with pathogenic bacteria, including members of the genus Vibrio (Oberbeckmann & Labrenz 2020). Thus, pathogen-enriched floating microplastics have the potential to disperse over long distances and spread pathogenic bacteria to new marine areas and ecosystems and should therefore be considered as a threat to marine ecosystems as well as to animal and human health (Bowley et al., 2021; Stabnikova et al., 2021).

Among the pathogens underscored on microplastics, many bacterial strains have been found to harbor antimicrobial resistance (AMR) as well as resistance to heavy metals (MRG) (Bowley et al., 2021). These multidrug resistances are correlated with the presence of heavy metals, organic pollutants, and traces of antibiotics in the marine environment, which can adsorb onto plastic biofilms (Imran et al., 2019). These substances are known to promote horizontal gene transfer (HGT) of virulence and resistance *via* mobile genetic elements (MGE) within bacterial communities (Sobecky & Hazen, 2009). Therefore, microplastics and their associated adsorbed chemicals, by promoting horizontal gene transfer in bacteria, contribute to the selection and dispersal of antimicrobial and metal resistance (Arias-Andres et al., 2018; Marathe & Bank, 2022). Finally, microplastics can have a significant impact on the spread of multidrug-resistant (MDR) pathogens, which may represent an additional threat (in terms of dangerousness compared to simple bacterial contamination) to the entire marine-related trophic network, including humans (Wright et al. 2020).

In the Indian Ocean (IO), recent reports highlight plastic accumulation along the coasts from Australia to India, in the Arabian Peninsula, along the coasts of East Africa and of the IO islands (Pattiaratchi et al., 2022). In the open ocean, the authors reported the absence of any rubbish patch in the northern IO, while a significant patch was identified in the southern IO in relation to the South
107 Atlantic Ocean (Pattiaratchi et al., 2022) and the South Pacific Ocean (Maes et al., 2018). According to Pattiaratchi et al. (2022), Reunion Island, an oceanic island located in the southwest part of the IO, is also impacted by this marine plastic pollution. This island, located at the crossroads of southern Africa and the Indian subcontinent, is also strongly affected by the phenomenon of AMR. In the human sphere, Reunion Island is subject to a very high pressure of importation of MDR and extensively drug-resistant (XDR) pathogens, linked to the population flow in the area (Miltgen et al., 2020; Miltgen et al., 2021; Kamus et al., 2022). However, there is very little data on the environmental spread of these pathogens from human excreta after the discharge of these effluents from the wastewater treatment plants into the ocean (Miltgen et al., 2022). The same is true for microplastics that are subject to human pollution in other territories and that can drift via the oceans to Reunion Island.

118 Thus, the microbial communities existing on the plastic marine pollution that reaches the coasts of Reunion Island could be affected by this AMR phenomenon. Therefore, it is of utmost importance to determine whether the microbial communities colonizing the marine microplastic debris drifting off the coast of Reunion Island host pathogenic bacteria, potentially resistant to several antimicrobials, which should then be considered as a threat to public health.

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The present study aims at (i) characterizing the microbiome hosted by marine microplastics drifting in the coastal waters of Reunion Island and (ii) addressing the presence of AMR potential pathogens carried by these microplastics. To our best knowledge, this study is the first in the southwest Indian Ocean islands, including physico-chemical, genomic, and microbiological approaches. It integrates public health concerns and local environmental issues with the aim of shedding light on the role of microplastics and the consequences that this new human-induced niche may have, not only on the marine environment and island ecosystems, but also potentially on the entire marine food chain, up to humans, in a One Health approach (Wright et al., 2020).

131 2. Materials and methods

132 2.1 Site description and sampling methods

133 Reunion Island is located at 55° East 21° South, 700 km east of Madagascar (Fig. 1). The two
134 selected sites (see map in Fig. 1) are distinct in terms of anthropogenic disturbance and oceanic
influence: the first site, Livingstone (21°05'02.5"S 55°13'33.6"E), is located on the leeward west coast, at the level of the Ermitage lagoon, in Saint-Gilles municipality, while the second site i.e. the Tremblet beach in Saint-Philippe municipality (21°17'38"S 55°48'19"E) is located on the windward east coast. The 1st site is heavily impacted by local anthropogenic activities (Tourrand et al., 2013; Guigue et al., 2015; Lemahieu et al., 2017) while the 2nd site is a newly formed beach, almost untouched by human activity, resulting from a volcanic eruption that occurred in 2007 (Staudacher et al., 2009). The collected samples were, on one hand, the plastics from the coastal seawater (PSW) and the sand beach (PS), and on the other hand, the substrates i.e. the coastal sea-water (SW) and the beach sand (S). beach sand.

Plastic debris were collected from the seawater surface at 200 m from the shoreline using a manta net (mesh size: 500 µm; mouth area: 1.125 m²) provided by the non-governmental organization (NGO) "The Ocean Clean Up" (Rotterdam, The Netherlands; https://theoceancleanup.com) (Virsek et al., 2016)). The speedboat was sailing at 2 knots and the sampling time was 20 minutes, with sampling days chosen in fair weather. Three transects were set at each site to generate replicates per area (GESAMP, 2019). Between each replicate, the manta net was rinsed externally with a jet of seawater and all plastic particles were collected. Plastic concentration was calculated following Kukulka et al. (2012). Plastic debris collection numbered by sample according to site and substrate are reported in Supplemental Table 1. At the same time, seawater samples collected from the coastal area (3 replicates of 2 L samples per collection) were processed according to the protocol of Hinlo et al. (2017). In parallel to seawater collection, plastic debris were collected from the beach at each site following the protocols of Besley et al. (2016) and using a 1 m × 1 m sampling quadrat. Three 50 x 10 m corridor transects running parallel to the sea were conducted for each site. We report the density of collected plastic wastes in items/km². At the same time, beach sand samples were collected in triplicates according to the protocol reported by Almeida et al. (2019). From these plastic sample sorting, sub-samples of 50 microplastics particles (size < 5 mm) were randomized (Löder & Gerts, 2015) to carry out optimal DNA extraction as suggested by Debeljack et al. (2017). To avoid air contamination, the separation of the microplastics (< 5 mm) was carried out in a clean room and under a binocular
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magnifier under sterile conditions. These randomized subsamples were assembled in triplicates for both DNA and living microbe extractions (see further for the pre-treatment detailed protocol).

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## 2.2. Sub-samples pre-treatments

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Each microplastic sub-sample of 50 particles was treated according to the protocol of Trachoo (2004) by gentle abrasion to extract DNA and cultivable bacteria while keeping the polymer structure of the plastic intact. For this, 10 g of washed and sterile (see below) silica sand (Sigma-Aldrich, Darmstadt, Germany) was added to a sterile 50 ml Falcon tube (Fischer, Illkirch, France) containing 50 particles of microplastic. Silica sand was treated before use as follows: sand was first washed for 10 min with 2% hydrochloric acid (Merck, Darmstadt, Germany), then rinsed 3 times with MilliQ water. Silica sand was then washed for 10 minutes with sodium hypochlorite solution Emplura at final concentration of 2% (Merck, Darmstadt, Germany) and rinsed 3 times with MilliQ water. Finally, sand was rinsed once for 10 minutes with MilliQ water before being autoclaved (120°C, 20 min, 1 bar). An artificial seawater solution was reconstituted by dissolving 35 g of NaCl (Sigma-Aldrich, Darmstadt, Germany) in 1 L of MilliQ water and then autoclaved (120°C, 20 min, 1 bar). A volume of 20 mL of reconstituted sterile seawater and 20 g of sterile and washed silica sand were added to each microplastic sample. Similarly, the beach sand samples (20 g in a 50 ml Falcon tube) were supplemented with 20 ml of reconstituted sterile seawater. All samples (microplastics and beach sand) were vortexed for 60 seconds. The supernatant was collected and divided into two aliquots of 10 ml for DNA extraction and microbiological analysis. Seawater samples collected from the coastal area (3 replicates of 2 L samples per collection) were processed according to the protocol of Hinlo et al. (2017). For each 2 L sample of sea water, there were two separate filtrations of 1 L on a sterile nitrocellulose membrane (0.22 µm): one filter was stored at -20°C for DNA extraction and the other one underwent resuspension of bacteria by vortexing in 5 ml of reconstituted sterile seawater for microbiological analyses.

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## 2.3 DNA extractions, 16S PCR and library preparation
The 10 ml of supernatants from the plastic abrasion or sediment extractions were filtered and sterilized through 0.22 µm nitrocellulose membranes (Merck Millipore, Cork, Ireland). DNA was extracted from all nitrocellulose membranes resulting from the plastic, sediment supernatant and water filtrations, following the protocols of Debeljack et al. (2017). Briefly, the membranes were placed in Qiagen DNeasy Blood and Tissue kit columns (Qiagen GmbH, Hilden, Germany) and DNA was extracted according to the Qiagen manufacturer's instructions. After extraction, DNA was quantified using a Nanodrop spectrophotometer (Thermo Scientific France, Ilkirch-Graffenstaden, France). The DNA samples were then sent to Macrogen's Next Generation Sequencing (NGS) platform (Macrogen, Seoul, Korea) for mass DNA sequencing. Library construction and sequencing were performed according to Illumina 16S metagenomic sequencing library protocols to amplify the V3 and V4 region of 16S DNA (Bukin et al., 2019). Two nanograms of genomic DNA were amplified by PCR with 5x reaction buffer, 1 mM dNTP mix, 500 nM of each of the universal F/R PCR primers and Herculase II fusion DNA polymerase (Agilent Technologies, Santa Clara, CA). The cycling condition for the 1st PCR was 3 min at 95°C, and 25 cycles of 30 sec at 95°C, 30 sec at 55°C and 30 sec at 72°C, followed by a final extension of 5 min at 72°C. The V3-V4 domain of the 16S rDNA was amplified by PCR using the following primers V3-341F: 5'-CCTACGGGNGGCWGCAG-3', V4-805R: 5'-GACTACHVGGGTATTAATCC-3' (Klindworth et al., 2013) with Illumina adaptor overlays. The PCR product was purified with AMPure beads (Agencourt Bioscience, Beverly, MA) and 2 µl of the purified product was PCR amplified for construction of the final library containing the index using the Nextera XT index primer. The cycling condition for the 2nd PCR was the same as the 1st PCR conditions. The PCR product was purified with AMPure beads. The final purified product was then quantified using qPCR according to the qPCR quantification protocol guide (KAPA library quantification kits for Illumina sequencing platforms) and qualified using the TapeStation D1000 ScreenTape (Agilent Technologies, Waldbronn, Germany).

2.4 NGS analyses.

Paired sequencing (2×300 bp) was performed using the MiSeq™ platform (Illumina, San Diego, USA). Adapter pruning was performed using the fastp program, adapter sequences were
removed and error correction was performed in overlapping sequences (Chen et al., 2018). The read
assembly was performed by assembling pair-end sequences created by sequencing both directions of
the library. The program used in this process is *FLASH (v1.2.11) (Magoc & Salzberg, 2011).
Assembled reads shorter than 400 bp or longer than 500 bp were removed. Next, the preprocessing
and clustering process was performed according to the protocols of Li et al. (2012): data with
sequence errors were removed in order to obtain accurate OTUs. Reads containing ambiguous bases
and chimeric sequences were also removed. After this process, clustering was performed based on
sequence similarity with a cut-off value of 97% using CD-HIT-OTU, a comprehensive program based
on cd-hit-est. Community diversity and taxonomy were analyzed according to Caporaso et al. (2010)
using QIIME (v1.9.0), which is used for OTU analysis and taxonomy information. The main sequence
of each OTU was referenced in the NCBI 16S database, and taxonomic information was obtained with
BLASTN (v2.4.0).

2.4. Microbiological analysis

Successive serial dilutions from 1 to 10⁻² were prepared from the bacterial supernatant
resulting from the plastic / sand abrasion or the seawater 0.22 µm membrane filter resuspension using
the reconstituted sterile seawater. The total bacterial flora was counted by inoculating 100 µl of these
dilutions onto plate agar of Mueller Hinton + PolyViteX (PVX, BioMérieux, Marcy l'Etoile, France)
and Mueller Hinton E (MHE, BioMérieux, Marcy l'Etoile, France) media as previously described
(Miltgen et al., 2020; Bhuyan et al., 2023). Once inoculated, the media were incubated at 35°C ± 2°C
for 24 to 72 h, until the microbial colonies appeared visible. The total bacterial flora was quantified
and expressed as colony forming units (CFU) per g of plastic or sand or ml of seawater. The following
selective media were also used: Columbia Naladixic Acid Agar (CNA, bioMérieux, Marcy l'Etoile,
France) for the identification of the Gram-positive bacteria, Drigalski (DRIG, bioMérieux, Marcy
l'Etoile, France) for Gram negative bacteria and chromID CPS ELITE (CPSE, bioMérieux, Marcy
l'Etoile, France) for a control. For each sample, subcultures were analyzed, and each phenotypically
distinct colony was re-isolated on Mueller Hinton agar (MHE) and incubated at 35°C ± 2°C. After 24
to 72 hours, the individual colonies were identified using MALDI-TOF (Matrix Assisted Laser
Desorption Ionization Time-Of-Flight) mass spectrometry (Bizini & Greub, 2010). After identification, pathogens were counted on selective media to estimate the relative abundance of each bacterial genus. The antimicrobial susceptibility of each pathogen was assessed by the disk diffusion or gradient strips methods (Miltgen et al., 2020). After 18-24 hours, the inhibition diameter around each antibiotic disc or the MIC (minimal inhibitory concentration) were measured and the bacterial/antibiotic pair was categorized susceptible, intermediate or resistant (S/I/R) following the recommendations of The European Committee on Antimicrobial Susceptibility Testing (2020 EUCAST, https://www.eucast.org), while the resistance for isolates belonging to Vibrio spp. was categorized using the Clinical Laboratory Standards Institute (CLSI https://clsi.org/) recommendations.

## 2.5. Microplastic polymer identification

Microplastic particles were retrieved after the sand abrasion as described above. They were characterized by using Fourier Transform InfraRed spectroscopy (FTIR Nicolet i550, Thermo Fisher) in ATR (Attenuated Total Reflection) mode, with wavelengths ranging from 400 to 4000 cm⁻¹ (resolution of 1 cm⁻¹). The pieces were one by one pictured then pressed between diamond and base (Djaoudi et al., 2022). The diamond was cleaned between each particle analysis. Final infrared spectrums (average of 40 scans) were analyzed using SpectraGryph software and its database. Only correspondences higher than 85% were validated. The polymer FTIR identification and frequency per sample are reported in Supplemental Table 1.

## 2.6 Microbial community analyses based on OTUs abundance, taxonomy and statistical methods

A full overview of this analytical approach is presented in Supplemental Fig. 1. We used OTUs abundance data resulting from NGS analyses to compare microbial communities between: (i) sampling coasts (East or West coast), (ii) the matrices from which plastics were sampled (sand beach or seawater) and (iii) the sampled materials (plastic, seawater, or sand beach). The sampling DNA sequencing quality was firstly checked with rarefaction curves (Supplemental Fig. 2a, b, c, d)
computed with iNEXT online (Chao et al., 2016). We used a top-down taxonomic approach to explore the differences between microbial communities by first analyzing the full data at the phylum level then focusing only on the most frequent genera belonging to the Proteobacteria phylum (genera representing more than 1% OTUs total abundance: 15 out of the 405 genera identified in the full database, Supplemental Fig. 3). OTU abundances raw data are available as Supplemental table 2. The composition of microbial communities is presented using (i) barplots of the relative abundance of organisms (cumulated abundances are given in Supplemental tables 2a at phylum level and Supplemental table 3a for genera belonging to the Proteobacteria phylum level) and (ii) Non-Metric Multidimensional Scaling (NMDS) (based on Bray-Curtis dissimilarities) plots using the "vegan" package for the R software (R Core Team 2021, v4.0.4). In both graphical representations, data were grouped according to environmental parameters (combination of matrix and material), with NMDS plots showing ellipses for 95% confidence intervals. Overall differences in bacterial communities' composition were statistically assessed between sample sites, substrates and ecosystems using analysis of similarities (ANOSIM) on the same Bray-Curtis distance matrix (Dixon, 2003). Pairwise differences between groups were statistically assessed using Chi² tests and corrected according to the Bonferroni method. Ecological diversity was measured using several indices: specific richness and the Shannon and Simpson diversity indices. Overall differences in ecological diversity between groups were tested using non-parametric Kruskal Wallis test, followed by Duncan's post-hoc tests when more than two groups were compared and non-parametric Wilcoxon Mann-Whitney test if two groups were compared. Venn diagrams were used to illustrate the dissimilarity in composition between groups (Oliveiros, 2007).

Linear models were used to compare (i) the abundance of plastic debris collected at the sea surface and on beaches between sites (East vs West) and polymer type and (ii) the abundance of culturable bacterial populations isolated from plastic, water and sand samples from different sites or substrates. Tukey signed-rank tests were further used to evaluate pairwise differences.

## 3. Results

### 3.1. Plastic data
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At the sea surface, concentrations of particles were highly variable across samples, ranging from 0 to 7,391 items / km² on the west coast and from 3,561 to 23,692 items / km² on the east coast (Table 1 and Supplemental Table 1). There was no significant difference between the west and east coasts in terms of item concentrations (p.value > 0.05) (Table 1). Most of the debris (85%) have been successfully classified by FTIR analysis. Both on the west and east coasts, the most abundant plastic polymers identified were polyethylene (PE), representing 75% and 84% of particles respectively; and polypropylene (PP) representing the remaining 25% and 16% (Table 1). There was no significant difference between the west and east coast in terms of polymer type (p.value > 0.05). For the beaches, two significant different concentrations of 0.34 ± 0.31 and 0.022 ± 0.008 item/m² were estimated for the west and east coast, respectively. Significant differences between the diversity of polymers on the strandings were found on the west coast, there was 50% PE, 38% PP, 9% polystyrene (PS) and 3% polyvinyl chloride (PVC) while on the east coast, there was 70% PE, 29% PP and 1% PS (Table 1).

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## 3.2. Bacterial microbiome analyses based on the 16S rDNA sequencing.

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### 3.2.1 Total OTUs diversity and abundance

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A total of 4,052,436 reads were retained (184,201 ± 22,611 reads per sample on average) after quality filtering and chimera checking, reads abundances ranged from 138,698 to 213,256 for the plastics sampled from seawater of the West Coast (PSWw) and seawater samples from East Coast (SWe), respectively (Supplemental Tables 2a, b, c). Read numbers did not differ significantly between plastics and substrates and between coasts (Supplemental Tables 2a, b) and no clear grouping was detected when samples were compared pair wisely (Supplemental Table 2c). All rarefaction curves showed an early stationary phase indicating sufficient sequencing depth of the taxa amplified in the microplastic, sand and seawater matrices (Supplemental Fig. 2a, b, c, d). Overall, high-quality sequences were grouped into 1,084 OTUs, 877 OTUs were identified from microplastic samples, 747 from the sand samples and 468 from seawater samples. These 1,084 OTUs (Supplemental Table 3) were tallied at an overall mean of 21,299 ± 5,127 OTUs / sample, with the difference between the East coast and West coast sites being not significant (Supplemental Table 2a, p > 0.05), as the difference between plastics sampled in sea water and on plastics sampled in sand (Supplemental Table 2b, p > 0.05). However, OTUS were significantly more abundant for plastics than for both of their substrates
322 (seawater or sand), with OTUS being 1.46 times more abundant in plastic DNA samples than in
323 seawater DNA samples, and 1.32 times more abundant for plastics sampled in sand (Supplemental
324 Table 2b, p < 0.05). Microplastic debris from seawater shared 28.5% and 60.7% of OTUs with
325 seawater in the western and eastern sites, respectively, while microplastics from the sandy beach
326 shared 50.2% and 61.2% of OTUs with sand beach in the western and eastern sites, respectively (Fig.
327 2).

328 3.2.2 Diversity of OTUs at the phylum level

329 Phylum richness was significantly higher on sand than on water (Table 2). However, there
330 was no significant difference between the study sites (East versus West) nor between plastic versus
331 non-plastic substrates (seawater or sand beach). Shannon and Simpson diversities differed
332 significantly between study sites and between plastic and non-plastic samples (Table 2) but not
333 between sand and water. Shannon and Simpson diversities were higher on the east coast than on the
334 west coast and in non-plastic samples compared to plastic samples (Table 2). The full diversity
335 indexes of the bacterial phylum distribution according to the site and the substrate are presented in
336 Supplemental Table 4a.

337 An overall number of 19 phyla was identified in both plastics and their substrates samples
338 (Fig. 3a, Supplemental Table 4b). All samples were dominated by Proteobacteria (75%), Bacteroidetes
339 (11%), Cyanobacteria (5%) and Fimicutes (4%). Actinobacteria, Planctomycetes and Verrucomicrobia
340 were also present across all the samples but at lower levels (<2%). At the phylum level, bacterial
341 communities associated with plastics harvested in sea water (PSW) stands out from other groups
342 (ANOSIM and Chi² tests, Fig. 3b and Supplemental Tables 4c and d). Height phyla were detected in
343 seawater and sand but not on plastics: Acidobacteria, Chlamydiae, Gemmatimonadetes,
344 Ignavibacteriae, Kiritimatiellaeota, Lentisphaerae, Spirochaetes and Thaumarchaeota. Phylum
345 communities did not differ between sites (West vs. East) and matrices (samples harvested from
346 seawater vs. samples harvested from the sand).

347 3.2.3 Proteobacteria

348 Proteobacteria were the dominant phylum (75.6% of the total of OTUs) and were further
349 analyzed. To better analyze data at the genus level, we filtered out the least frequent Proteobacteria
350 OTUs (frequency <1%). 15 out of 405 genera, representing 79.4 % of the total Proteobacteria OTUs
351 were kept (see Supplemental Fig. 3a and Supplemental Table 3 for full results). Genera richness and
352 Shannon diversity did not differ between sites (East vs. West) and environmental parameters (sand or
353 seawater), nor between plastic and non-plastic samples (Table 2 and Supplemental Table 5a, b and c).
354 Simpson diversity did not differ between sites (East versus West) nor between plastic versus non-
355 plastic samples but differed significantly between substrates (sea water versus sand) (p < 0.05). Across
356 all samples four genera accumulated more than 70% of the overall OTU abundance: *Psychrobacter*
357 (21.9% in all samples and 28.8% in plastics), *Vibrio* (20.2% in all samples and 17.1% in plastics),
358 *Pseudoalteromonas* (17.6% in all samples and 18.1% in plastics) and *Photobacterium* (14.8% in all
359 samples and 19.4 % in plastics) were among the most found abundant genera belonging to the phylum
360 of the Proteobacteria. Nonetheless, the composition of proteobacterial communities differed strongly
361 between the west and the east coasts (Fig. 4a, Supplemental Table 5c). Considering plastic, seawater,
362 and sand samples, on east coast *Vibrio* (33.1%), *Pseudoalteromonas* (25.6%) and *Photobacterium*
363 (19.3%) were dominant whereas the genus *Psychrobacter* (39.7%) was the most abundant on the West
364 coast. Plastic samples showed genera compositions different from seawater or sand samples (Fig. 4a,
365 Supplemental Table 5c) with a dominance of *Psychrobacter* (28.8%), *Photobacterium* (19.5%),
366 *Pseudoalteromonas* (18.1%) and *Vibrio* (17.1%) on plastics, and *Vibrio* (30.1%), *Pseudoalteromonas*
367 (15.8%), *Candidatus Pelagibacter* (12.7%), *Alcanivorax* (12.6%) and *Alteromonas* (10%) in seawater
368 and sand. Proteobacterial communities associated with microplastics or found in the water column or
369 on the beach sand were further differentiated using NMDS ordinations and ANOSIM tests (Fig. 4b).
370 The distribution of proteobacterial communities differed significantly (Supplemental Table 5d)
371 between sites (green symbols vs blue symbols on Fig 4b), material collected (S + SW filled symbols vs
372 PS + PSW empty symbols on Fig 4b), and the type of sample (combination of both site and material,
373 represented with four ellipses on Fig 4b). However, the proteobacterial communities found on sand
374 beaches (PS + S) vs coastal waters (PSW + SW) were similar (Fig 4b; Supplemental Table 5d).

375

376 3.3. Cultivable bacterial flora analysis and antimicrobial resistance
A dense cultivable bacterial flora was found on the microplastics from Reunion Island (3.13 x 10⁷ colony forming units (CFU) /g of microplastics on average, Fig. 5, Supplemental Table 6). The total cultivable flora was significantly denser on microplastics than in seawater (4.82 x 10² CFU/ml of water) or on sand (3.89 x 10⁴ CFU/g of sand) whatever the studied site. On the contrary, no differences were found between the density of this cultivable flora on microplastics collected on the east or the west coast, or between the seawater and the sand. The fraction of potentially pathogenic bacterial microflora was isolated on selective media: a total of 105 bacterial strains could be identified at the genus level using MALDI-TOF analysis: summarized data are presented in Supplemental Table 7 while numeration data of these strains per genus are presented in Supplemental Table 8. Various genera belonging to the phylum Proteobacteria, Firmicutes and Bacillota were found on plastics, seawater, or sand. On plastics these cultivable bacteria reach significantly higher abundances (10³ to 10⁵ CFU/g of plastic) than in seawater (1 CFU/ml of seawater) and on sand (10² CFU/g of sand). Noteworthy, on plastics from both sites (East or West) and both matrices (seawater or sand), the most dominant bacterial genera were *Exiguobacterium* and *Pseudomonas* with several cultivable bacteria scaling from 10³ CFU/g for *Pseudomonas* to 10⁵ CFU/g for *Exiguobacterium*. Acquired resistance profiles could be sorted for 16 of the 105 strains identified by MALDI-TOF (Table 3 and Supplemental Table 9a, b). Microplastics carried bacterial strains with AMR, including strains with non-intrinsic resistances to antibiotics. The main antibiotic resistances detected concerned β-lactams such as penicillin, ampicillin and ticarcillin. On plastics, the AMR potential pathogens detected were strains belonging to *Bacillus*, *Enterococcus*, *Pseudomonas* and *Pantoea* genera.

## 4. Discussion

This study contributes to the knowledge of the health risk associated with the plastisphere in an oceanic region still very little documented.

### 4.1 Reunion Island suffers from plastic pollution

Our observation shows that surface coastal waters around Reunion Island are polluted by plastic. Concentrations of 10,693 ± 1,275 items/km² and 4,025 ± 4,760 items/km² were measured on the West and East coasts respectively. Plastics pollution at Reunion Island may come from local plastic waste sources (Benard et al., 2022) but also from plastic marine debris from the Indian ocean subtropical
405 plastic gyre (Pattiaratchi et al., 2022), with these plastic debris mainly coming from Southeast Asia
406 and carried by the south equatorial current (Thibault et al., 2023). These results are consistent with
407 observations in most other seas and oceans (Thushari & Senevirathna, 2020). In addition, despite of
408 the geographical isolation of Reunion Island in the Indian ocean, the data of the present study argue in
409 favor the hypothesis that (i) marine plastic pollution is a global connected threat to all the world's
410 oceans (Lincoln et al., 2022), and (ii) most scattered oceanic islands are threatened by such a pollution
411 (Derraik et al., 2002).

412 However, the concentrations of these stranded plastics on the beaches of Reunion Island (0,35
413 items/m² on the west coast and 0.0223 items/m² on the east coast) are lower than reported on other
414 Indian Ocean islands such as the Chagos Archipelago (6 items/m2, Hoare et al., 2022) or the Maldives
415 (35.8 items/m², Imhof et al., 2017). Plastic abundance varies with environmental settings such as wind
416 speed, swell intensity, marine currents velocity, seasonality, and the morphology of the island (Imhof
417 et al., 2017). Reunion Island is a young volcanic island (Lenat et al., 2001) harboring few coral reefs
418 and subject to oceanic swell and strong marine currents impacting the few beaches located on the east
419 coast (Pous et al., 2014). Alternatively, the low plastic abundance may result from distinct oceanic
420 influences as well as the remoteness of Reunion Island. The island is mostly under the influence of
421 south Easterly trade winds and currents, and waters circulate from Western Australia to Reunion
422 Island without colliding any important land mass (Schott et al., 2009). Concerning the nature of the
423 plastic polymers found in Reunion Island, whether on the sea surface or in the sand of the beaches,
424 polyethylene and polypropylene are the most abundant polymers. The dominance of these two
425 polymers is hegemonic across the world's seas and oceans as reported in the meta-analysis by Erni-
426 Cassola et al. (2019).

427

428 4.2 Marine microplastic debris reaching Reunion Island host a specific microbiome

429 In the present study we found that a substantial proportion of up to 60% of the operational
430 taxonomic units (OTUs) was shared between microplastics and their environment (seawater or sand
431 beach). Seawater microplastic debris are colonized by planktonic microorganisms forming a biofilm
432 whose composition presents expected high similarities with seawater microbiome (Zettler et al., 2013;
De Tender et al., 2017). Our observation highlights the colonizing role of bacterioplankton but also suggests a relatively new colonization (40% of OTUs are specific to plastics). Such renewal of microbiome composition is known as resulting from the oceanic environmental factors as temperature, salinity, light, and UV radiations: these factors affect the microbial biofilm composition and its renewal with an enrichment in pathogens (Sooriyakumar et al., 2022; Shan et al., 2022).

Similarly, plastic debris reaching beaches, after being introduced into the marine environment, undergo physical and biological processes that break them down into smaller fragments, which can eventually be incorporated as part of the beach sand (Napper & Thompson, 2020). Through this process, the plastic fragments and the sandy beach substrate can contribute to each other's bacterial enrichment, which the high proportion of shared OTUs may reflect. A recent study has shown that a high proportion of OTUs shared between plastic debris and sand samples suggests that plastic debris have a significant impact on microbial communities in marine sediment (Seeley et al., 2020). Nevertheless, there is still an important proportion of OTUs (up to 38%) that are specific to the plastic debris indicating that plastics are also a microbial ecological niche for specific communities compared to the communities from the environment. A recent review suggests that such specific communities result from biofilm formation and evolution processes, with an enrichment in bacteria able to use the plastic polymer as carbon source (Sooriyakumar, et al., 2022; Zhurina et al., 2022).

### 4.3. Proteobacteria and Bacteroidota phyla dominate marine microplastic microbiome

The microbial community structure on Reunion Island's plastic pollution exhibits a notable dominance of Proteobacteria and Bacteroidota phyla across various sample types, including seawater-plastic, sand-plastic, seawater, and beach sand. These phyla, particularly Proteobacteria (75% average relative abundance) and Bacteroidota (11%), remain consistent irrespective of the substrate or site studied, reflecting their adaptability to diverse environmental conditions. Proteobacteria and Bacteroidota are diverse groups of bacteria known for their ability to adapt to a wide range of environmental conditions (Kertsters et al., 2006; Hahnke et al., 2016). They are commonly found in the oceans and their high abundance is well documented in seawater and marine sediments (Stal &
460 Cretiou, 2022). Our observation is consistent with other reports that also observed dominance of
461 Proteobacteria and Bacteroidota phyla associated to plastic biofilm, e.g. Northern Europe seas,
462 Mediterranean Sea and the Atlantic Ocean (Oberbeckmann et al., 2014; Vaksmaa et al., 2021, Debroas
463 et al., 2017).

464 Within the dominant Proteobacteria phylum, specific dominant genera stand out. On seawater
465 plastics, *Photobacterium* (33%), *Pseudoalteromonas* (27%), and *Psychrobacter* (18%) are highly
466 abundant, comprising up to 78% of the Proteobacteria OTUs found on plastics. In surrounding
467 seawater, dominant *Proteobacteria* genera, including *Candidatus* (24%), *Vibrio* (24%), *Alcanivorax*
468 (18%), and *Alteromonas* (11%), collectively represent over 77% of the OTUs. These significant
469 differences between Proteobacteria found on plastics and in surrounding waters suggest that marine
470 microorganisms have adapted to plastics as a colonization surface (Roager & Sonnenschein, 2019).
471 Moreover, several of these dominant Proteobacteria genera (*Photobacterium*, *Pseudoalteromonas*, and
472 *Psychrobacter*) are known for their ability to biodegrade and utilize plastics as a carbon source and
473 nutrient (Raghul et al., 2014; Muriel-Millán et al. 2021; Atanasova et al., 2021). This presence on
474 seawater plastics indicates that the plastics reaching Reunion Island have likely undergone degradation
475 during prolonged floating, in line with the concept of plastic debris transporting microorganisms as
476 suggested by Pattiaratchi et al. (2022).

477 The diversity and abundance of Proteobacterial OTUs on plastics and sand from the East coast
478 resemble each other, with dominant genera such as *Vibrio*, *Pseudoalteromonas*, and *Alteromonas*,
479 suggesting a role for plastics in transporting and enriching the sand bacterial community. In contrast,
480 on the West coast, the Proteobacteria community structure significantly differs between sand plastics,
481 dominated by *Psychrobacter* (>80%), and sandy beach sand, where the Proteobacteria community is
482 led by the genus *Dyella* (>70%). This variation may be linked to the specific types of polymers present
483 on the plastics reaching each coast, with *Psychrobacter* known for its biodegradation of polyurethane
484 and polycaprolactone polyester (PCL) (Atanasova et al., 2021a). The dominance of *Dyella* in beach
485 sand may indicate residual wastewater contamination from forest or cultivated soils, where *Dyella* is
486 primarily isolated (Dar et al., 2020; Huang et al., 2021).
487

488

## 4.4 Cultivable bacteria isolated from marine microplastic debris

489

490

491

492

493

494

495

The study of cultivable bacteria isolated from marine microplastic debris reveals significant microbial activity, with an average concentration of 3 x 10⁷ colony forming units (CFU)/g of plastic. Notably, this demonstrates that microplastic serve as an ecological niche where bacteria remain metabolically active and cultivable. Comparatively, recent research found that plastic microbeads could harbour up to 60 x 10¹⁰ CFU/cm² of plastic (Türetken et al., 2020). In the present study at Reunion Island, the number of cultivable bacteria on plastics exceeded those in beach sediments by 800 times and coastal waters by up to 65 x 10³ times.

496

497

498

499

500

501

502

503

504

505

Similarly, potential pathogens cultivable bacteria were significantly more abundant on plastic debris (ranging from 10³ to 10⁵ CFU/g of plastic) than in surrounding environments like seawater or sediment (1 to 1000 CFU/g of sand or ml of seawater). This is consistent with Wu et al. (2019) work that showed higher abundance of pathogenic bacterial families on microplastics, compared to substrates. The most encountered potentially pathogens cultivable bacterial strains belong to the *Bacillus*, *Exiguobacterium*, and *Pseudomonas* genera, raising concerns about the potential transmission of infectious pathogens to humans or animals (Bergan, 1981; Mandic-Mulec et al., 2015; Chen et al., 2017). Remarkably, many of these potentially pathogenic bacteria also possess the capacity to degrade plastics, exemplified by the genus *Pseudomonas*, which is both abundant (up to 10⁵ CFU/g of plastic) and proficient in plastic degradation (Wilkes & Aristilde, 2017).

506

507

## 4.5 AMRs bacteria

508

509

510

511

512

513

514

Marine plastics have been shown to enrich the plastisphere with pathogens, including genera like *Vibrio*, *Pseudomonas*, *Acinetobacter*, *Aeromonas*, and the Enterobacterales family, many of which have acquired resistance to antibiotics (Junaid et al., 2022). This poses a significant threat to public health and the environment (Marathe & Bank, 2022). In the Western Indian Ocean, this study represents one of the first reports of isolating cultivable pathogens from plastics for potential antimicrobial susceptibility testing (AST). Resistance to antibiotics was observed in strains of *Bacillus*, *Enterococcus*, *Pseudomonas*, and *Vibrio* isolated from Reunion Island's seawater or sandy
beaches. The majority of observed resistance was directed against molecules in the β-lactam family, such as ampicillin, penicillin, and ticarcillin. This is consistent with recent findings that residual beta-lactams, including ampicillin, are common emerging pollutants that chemically adsorb onto plastic, promoting resistance among the plastisphere's bacterial community (Imran et al., 2019; Wang et al., 2021). Furthermore, beta-lactam resistance genes (bla genes family) are often associated with mobile genetic elements (MGEs), facilitating horizontal gene transfer within the plastisphere (Wang et al., 2021; Zhang et al., 2022; Silva et al., 2023). This data, collected in a high-income country in the Southwest Indian Ocean, underscores that microplastic debris serves as a reservoir of potentially antibiotic-resistant pathogens (AMR), aligning with similar reports from around the world (Yang et al., 2019; Bowley et al., 2021; Liu et al., 2021). Importantly, this should be viewed from a One Health perspective, acknowledging that highly anthropized coastal environments can be contaminated by enteric bacteria from human or animal sources, potentially including MDR bacteria, based on regional epidemiology (Fernandes et al., 2020; Miltgen et al., 2022).

528

## 5. Conclusions

In this study, we presented a first case of the plastisphere assessment from Reunion Island, a remote oceanic island located in the Southwest Indian Ocean, polluted by plastic debris from various geographical origins. The characterization of plastic pollution in the island's coastal waters and beaches indicates that Reunion Island is facing plastic pollution with up to 10,000 objects/km² in the coastal waters, mainly consisting of polyethylene (up to 75%) and polypropylene (up to 25%). Plastic debris host dense microbiomes, dominated by Proteobacteria (80%). In addition, the cultivable microbiotes reached 10⁹ CFU/g of microplastics, with a dominance of bacteria from genera *Exiguobacterium* (10⁵ CFU/g of plastic) and *Pseudomonas* (10³ CFU/g of plastic). This plastic debris also carries β-lactam resistant AMR bacteria such as certain strains of the genera *Bacillus*, *Enterococcus* and *Pantoea* resistant to ampicillin, penicillin and ticarcillin. Overall, our results confirm, as it has already been described for other islands in other oceans and seas, that the islands of the Indian Ocean are facing severe marine plastic pollution, the debris of which host a dense
542

plastisphere including AMR bacteria. Our data also suggests potential risks associated with a plastic-
specific microbiome for Southwest Indian Ocean socio-ecosystems.

544

545

6. The following list is the Supplemental data related to this article.

546

Supplemental Fig. 1: Diagram of NGS data analysis process.

547

Supplemental Fig. 2a, b, c, d: Alpha rarefaction (Chao1) curves showing the observed sampling effort. (a) East coast Plastic Sea water (PSW) + Sea water (SW); (b) East coast Plastic Sand (PS) + Sand (S)

549

550

Supplemental Fig 3: Proteobacteria OTUs abundances according to genera. The full list of OTUs
repartition according to the Proteobacteria genera including the rare Proteobacteria genera (<1%) is
reported as an Excel file in Supplemental Table 3.

551

552

553

Supplemental Table 1: Plastic debris collection number by sample according to site and environmental parameters (seawater or sand beach) and polymer IFTR identification frequency per sample.

554

555

Supplemental Table 2a, b, c: NGS full data: number of total reads and OTUs per sample including means and standard deviation calculations. Two-way Anovas were carried out to compare OTUs abundances data by site (a) or substrate (b) or samples (c) and letters indicate significantly different means according to a test of Duncan (at p < 0.05). Abbreviations: S: sand; SW: sea water; PSW: plastic from sea water; PS: plastic from sand.

556

557

558

559

560

Supplemental Table 3: Excel file presenting the full taxonomic list of the 1084 identified OTUs and their sample distribution at Phylum level and Proteobacteria genera level.

561

562

Supplemental Table 4a: Diversity indexes of the bacterial phylum distribution according to the site and the substrate. Data are mean ± standard errors. Letters indicate significantly different means, according to a post-hoc Dunn's multiple comparison test (at p < 0.05).

563

564

565

Supplemental Table 4b: Total sample OTUs number and distribution per Phylum. Abbreviations: S: sand; SW: sea water; PSW: plastic from sea water; PS: plastic from sand.

566

567

Supplemental Table 4c: Phylum OTUS distribution Chi2 test results and associated p.values.

568

Supplemental Table 4d: Analysis of similarity tests (ANOSIM) results for phylum. The groups tested are i) the site (East or West), ii) the matrix (sand beach or sea water), iii) the material harvested (sand,

569
water, or plastic); and iv) the sample type (combination of site and material). The results are given as ANOSIM R values and its significance.

Supplemental Table 5a: Diversity indexes of the Proteobacteria genus distribution according to the site and the substrate. Data are means ± standard errors. Letters indicate significantly different means, according to a post-hoc Dunn's multiple comparison test (at p < 0.05).

Supplemental Table 5b: Total sample OTUs number and distribution per Proteobacteria genus of which frequency was >1%. Abbreviations: S: sand; SW: sea water; PSW: plastic from sea water; PS: plastic from sand.

Supplemental Table 5c: Proteobacteria genera (15 genera > 1%) distribution Chi2 test results and p. values.

Supplemental Table 5d: Analysis of similarity tests (ANOSIM) results for Proteobacteria genera >1%. The groups tested are (i) the site (East or West), (ii) the matrix (sand beach or sea water), (iii) the material harvested (sand, water or plastic) and (iv) the sample type (combination of site and material). The results are given as ANOSIM R values and its significance.

Supplemental Table 6: Total culturable bacterial flora in CFU / g or ml of substrate, Abbreviations: PSW: plastics from coastal sea-water; SW: coastal sea-water; PS: plastics from beach sand; S: beach sand. Values are means ±standard deviation of means (n = 3) of three independent samplings, A two-anova was carried out and the different letters indicate significant differences as determined by Tukey HSD test (p ≤ 0.05).

Supplemental Table 7: Excel file presenting the list of the 105 cultivable bacterial strains isolated from selective media and identified at genus level by MaldiTof.

Supplemental Table 8a: Numbering of culturable bacterial strains according to the genus identified by Maldi Tof and expressed as CFU per g or ml of substrate i.e. plastic from sea water (PSW), plastic from sand beach (PS), sea water (SW) or sand beach (S) and associated literature indicating the bacterial genus potential pathogenicity and plastic degradation capacities.

Supplemental Table 8b: List of the bibliographic references of the Supplemental Table 8a.

Supplemental Table 9a and b: Full antibiograms data. Antibiograms were carried out according to antibiotic specific spectrum related to the bacterial genus tested. From 8 to 16 ATB were tested
according to the specific ATBiograms carried out to test the bacterial strains genus. In bold: ATB
resistance, in non-bold: ATB sensitivity.

Data accessibility: NGS raw data 16SrDNA sequences are deposited in zenodo data bank:
https://doi.org/10.5281/zenodo.8063253

7. Funding

This study was supported by funds from the Structure Fédérative de Recherche Biosécurité en milieu
Tropical (BIOST), Université de la Réunion (France) and Institut de Recherche pour le
Développement (IRD). Project ID: BMRPLAST.

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996 Authors contributions: Conceptualization P.J.; Data curation B.V., F.G., L.S., P.J., M.T., V.L. and
997 G.M.; Formal analysis B.V., L.S., P.J., M.T., V.L. and G.M.; Funding acquisition: P.J and G.M.;
998 Investigation: S.B., M.T., V.L., L.S., and P.J. Methodology: P.J., M.G. and G.M.; Project
999 administration: P.J.; Supervision: P.J.; Writing - original draft: B.V., L.S. and P.J. Writing - review
1000 & editing: M.G., V.L., F.G., P.T., T.B. and G.M.

1002 Funding: This work was supported by the Structure Fédérative de Recherche Biosécurité en milieu
1003 Tropicale (BIOST) de l'Université de la Réunion (Project BMRPLAST, 2021) and the Institut de la
1004 Recherche pour le Développement (IRD).

1006
1007
<figure><img src="image_2.png" /><figcaption>Fig. 1.(a) Location map of Réunion island in the southwest Indian Ocean close to Madagascar and Mauritius island; (b) Map of the Réunion island including the two major cities and the study site locations.</figcaption></figure>
a East Coast : (604 OTUs)

<figure><img src="image_3.png" /></figure>

b West Coast : (835 OTUs)

<figure><img src="image_4.png" /><figcaption>Fig. 2. Venn diagrams of the Operational Taxonomic Unit (OTUs) distribution showing shared and specific OTUs according to site i.e. East coast (a) or West coast (b) in all sample types i.e. PSW: plastics from coastal sea-water; SW: coastal sea-water; PS: plastics from beach sand; S: beach sand.</figcaption></figure>
a

<figure><img src="image_5.png" /><figcaption>Bacterial phylum OTUs relative abundance</figcaption></figure>

b

<figure><img src="image_6.png" /></figure>

Figure 3: OTUs distribution of the bacterial community according to the analysis at phylum level for the different substrates: PSW: plastics from coastal sea-water; SW: coastal sea-water; PS: plastics from beach sand; S: beach sand. (a) Relative abundance of bacterial phyla identified in all samples. (b) Two-dimensional NMDS ordination of bacterial community structure. Stress = 0.18. Ordination was based on the distance dissimilarity matrix. Ellipsoids represent the standard error confidence limit (95%) per substrate.
a

OTUs relative abundance of genera belonging to the Pseudomonadota phylum

<figure><img src="image_7.png" /></figure>

b

<figure><img src="image_8.png" /></figure>

Figure 4: OTUs distribution of the phylum of the Proteobacteria according to the analysis at genera level for the different substrates: PSW: plastics from coastal sea-water; SW: coastal sea-water; PS: plastics from beach sand; S: beach sand. (a) Relative abundance of Proteobacteria genus identified in all samples. (b) Two-dimensional NMDS ordination of Proteobacterial community structure. Stress = 0.18. Ordination was based on the distance dissimilarity matrix. Ellipsoids represent the standard error confidence limit (95%) per substrate.
<figure><img src="image_9.png" /><figcaption>Figure 5 : Total culturable bacterial flora in CFU / g or ml of substrate.
Abbreviations: PSW: plastics from coastal sea-water; SW: coastal sea-water; PS: plastics from beach sand; S: beach sand. Plots represent means, and error bars represent standard deviation of means (n = 3) of three independent samplings.
The different letters above plots indicate significant differences as determined by Tukey HSD test (P ≤ 0.05).</figcaption></figure>
<table><caption>Table 1: Quantification and characterization of plastic debris collected on East coast and West coast, in seawater or on sand beach. Except for plastic polymer nature, all data are means ± stdv of samples (n = 3) / site and substrate collection.</caption><thead><tr><th rowspan="2">Site</th><th rowspan="2"></th><th colspan="2">East</th><th colspan="2">West</th></tr><tr><th>Seawater</th><th>Sand beach</th><th>Seawater</th><th>Sand beach</th></tr></thead><tbody><tr><td rowspan="2">Substrate</td><td>Plastic item concentration (mean ± stdv)</td><td>4,025 ± 4,760<br>item/km²</td><td>0.022 ± 0.008<br>item/m²</td><td>10,693 ± 11,275<br>item/km²</td><td>0.34 ± 0.31<br>item/m²</td></tr><tr><td>Weight of plastic item in mg / item (mean ± stdv)</td><td>1.4 ± 1.3</td><td>31.4 ± 51.2</td><td>2.6 ± 7.4</td><td>56.5 ± 43.2</td></tr><tr><td></td><td>Weight of microplastic in mg / sample (mean ± stdv)</td><td>24.7 ± 0.1</td><td>202 ± 90</td><td>46.7 ± 0.7</td><td>1,798 ± 110</td></tr><tr><td rowspan="4">Plastic polymer (%)</td><td>Polyethylene</td><td>75</td><td>70</td><td>84</td><td>50</td></tr><tr><td>Polypropylene</td><td>25</td><td>29</td><td>16</td><td>38</td></tr><tr><td>Polystyrene</td><td>0</td><td>1</td><td>0</td><td>9</td></tr><tr><td>Polyvinyl Chloride</td><td>0</td><td>0</td><td>0</td><td>3</td></tr></tbody></table>
<table><caption>Table 2: Diversity indexes of the marine bacterial phyla and Proteobacteria genera found according to the combination of the site, the substrate, and the presence of plastic. Data are pooled according to three parameters: the study site (East or West), the type of substrate (Sea Water or Sand) and the presence of plastic (Plastic or non-plastic substrate i.e., Seawater or Sand). Data are reported as means ± standard errors. Letters indicate significantly different means according to a test of Wilcoxon-Mann-Whitney (at P < 0.05). Abbreviations: S: sand; SW: sea water; PSW: plastic from sea water; PS: plastic from sand. Full data of the diversity indexes per site and per substrate are presented in supplemental table 3.</caption><thead><tr><th rowspan="2">Parameters</th><th rowspan="2">Groups</th><th colspan="2">Richness</th><th colspan="2">Shannon diversity</th><th colspan="2">Simpson diversity</th></tr><tr><th>Phylum</th><th>Proteobacteria</th><th>Phylum</th><th>Proteobacteria</th><th>Phylum</th><th>Proteobacteria</th></tr></thead><tbody><tr><td rowspan="2">Site</td><td>East (S + SW + PSW)</td><td>9.8 ± 3.1 a</td><td>8.9 ± 0.6 a</td><td>2.7 ± 0.8 a</td><td>1.05 ± 0.06 a</td><td>2.0 ± 0.5 a</td><td>0.53 ± 0.03 a</td></tr><tr><td>West (S + SW + PSW)</td><td>8.4 ± 2.7 a</td><td>10.3 ± 0.4 a</td><td>1.9 ± 0.9 b</td><td>1.21 ± 0.11 a</td><td>1.6 ± 0.7 b</td><td>0.56 ± 0.06 a</td></tr><tr><td rowspan="2">Substrate</td><td>Sand (S + PS)</td><td>11.8 ± 1.8 a</td><td>10 ± 0.6 a</td><td>2.4 ± 1.1 a</td><td>1.00 ± 0.08 a</td><td>1.7 ± 0.6 a</td><td>0.48 ± 0.04 a</td></tr><tr><td>Water (W + PS)</td><td>6.9 ± 1.3 b</td><td>9.3 ± 0.5 a</td><td>2.2 ± 0.8 a</td><td>1.24 ± 0.09 a</td><td>1.8 ± 0.6 a</td><td>0.61 ± 0.04 b</td></tr><tr><td rowspan="2">Plastic</td><td>Plastic (PSW + PS)</td><td>8.6 ± 2.8 a</td><td>9.7 ± 0.6 a</td><td>1.9 ± 0.6 b</td><td>1.03 ± 0.09 a</td><td>1.5 ± 0.4 b</td><td>0.51 ± 0.05 a</td></tr><tr><td>Non-Plastic (SW or S)</td><td>9.8 ± 3.0 a</td><td>9.5 ± 0.6 a</td><td>2.8 ± 1.0 a</td><td>1.24 ± 0.09 a</td><td>2.1 ± 0.7 a</td><td>0.60 ± 0.04 a</td></tr></tbody></table>
<table><caption>Table 3: Antibiotic multiresistances detected among the culturable bacterial strains. In bold are noticed non-natural antibiotic resistances. Abbreviations: PSW : plastic from sea water; PS : plastic from sand beach; SW : sea water; S : sand beach; TS: Trimethoprim/ sulfamethoxazole; PT: Piperacillin/ Tazobactam; AC: Amoxicillin / Clavulanic acid.</caption><thead><tr><th>Site</th><th>Substrate</th><th>Bacterial strain MT code</th><th>Bacterial genus</th><th>ATB1</th><th>ATB2</th><th>ATB3</th><th>ATB4</th></tr></thead><tbody><tr><td rowspan="12">East</td><td rowspan="2">PSW</td><td>T2-14</td><td>Pantoea</td><td>Ampicillin</td><td>Cefadroxil</td><td>Ticarcillin</td><td></td></tr><tr><td>T3-42</td><td>Pseudomonas</td><td>TS</td><td></td><td></td><td></td></tr><tr><td rowspan="3">SW</td><td>T5-26</td><td>Staphylococcus</td><td>Erythromycin</td><td>Penicillin</td><td></td><td></td></tr><tr><td>T4-54</td><td>Bacillus</td><td>Amoxicillin</td><td>Imipenem</td><td>Penicillin G</td><td>Vancomycin</td></tr><tr><td>T4-52</td><td>Vibrio</td><td>Ampicillin</td><td></td><td></td><td></td></tr><tr><td rowspan="3">PS</td><td>A1-2</td><td>Pseudomonas</td><td>Ticarcillin</td><td>TS</td><td></td><td></td></tr><tr><td>A2-15</td><td>Bacillus</td><td>Amoxicillin</td><td>Imipenem</td><td>Penicillin G</td><td></td></tr><tr><td>A1-13</td><td>Vibrio</td><td>Ampicillin</td><td></td><td></td><td></td></tr><tr><td rowspan="2">S</td><td>A4-9</td><td>Bacillus</td><td>Amoxicillin</td><td>Penicillin G</td><td></td><td></td></tr><tr><td>A6-6</td><td>Enterobacter</td><td>Ampicillin</td><td>AC</td><td>Cefalexin</td><td></td></tr><tr><td rowspan="12">West</td><td rowspan="2">PSW</td><td>MEEP141</td><td>Pseudomonas</td><td>TS</td><td></td><td></td><td></td></tr><tr><td>E5-28</td><td>Pseudomonas</td><td>TS</td><td></td><td></td><td></td></tr><tr><td>SW</td><td>E4-13</td><td>Bacillus</td><td>Amoxicillin</td><td>Clindamycin</td><td></td><td></td></tr><tr><td>PS</td><td>B1-3</td><td>Enterococcus</td><td>Rifampicin</td><td>Vancomycin</td><td></td><td></td></tr><tr><td rowspan="2">S</td><td>B6-14</td><td>Aeromonas</td><td>Piperacillin</td><td>Ticarcillin</td><td>Ticarcillin / Clavulanic acid</td><td>PT</td></tr><tr><td>B5-17</td><td>Staphylococcus</td><td>Clindamycin</td><td>Fusidic acid</td><td>Penicillin</td><td></td></tr></tbody></table>