# Microalgae colonization and trace element accumulation on the plastisphere of marine plastic debris in Monastir Bay (Eastern Tunisia)

Mondher Tarchi, Noureddine Zaaboub, Majed Alsubih, Bochra Brik, Maria

Virgínia Alves Martins, Lotfi Aleya, Lamia Trabelsi

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Mondher Tarchi, Noureddine Zaaboub, Majed Alsubih, Bochra Brik, Maria Virgínia Alves Martins, et al.. Microalgae colonization and trace element accumulation on the plastisphere of marine plastic debris in Monastir Bay (Eastern Tunisia). Environmental Science and Pollution Research, 2023, 30 (12), 10.1007/s11356-022-23930-9. hal-05139222

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# Microalgae colonization and trace element accumulation on the plastisphere of marine plastic debris in Monastir Bay (Eastern Tunisia)

**Mondher Tarchi**

National Institute of Marine Sciences and Technologies: Institut National des Sciences et Technologies de la Mer

**Noureddine Zaaboub**

National Institute of Marine Sciences and Technologies: Institut National des Sciences et Technologies de la Mer

**Majed Alsubih**

college of Engineering, King khaled university

**Bochra Brik**

National Institute of Marine Sciences and Technologies: Institut National des Sciences et Technologies de la Mer

**Maria Virgínia Alves Martins**

Universidade de Aveiro

**Lotfi Aleya**

Université Bourgogne Franche-Comté: Universite Bourgogne Franche-Comte

**Lamia Trabelsi** (✉ lamiatABELSIakremi@gmail.com)

National Institute of Marine Sciences and Technologies: Institut National des Sciences et Technologies de la Mer https://orcid.org/0000-0001-5168-691X

## Research Article

**Keywords:** Marine plastic debris, plastisphere, epiplastic microalgae, trace elements, Monastir Bay, Tunisia

**Posted Date:** September 1st, 2022

**DOI:** https://doi.org/10.21203/rs.3.rs-1915380/v1

**License:** © 🌐 This work is licensed under a Creative Commons Attribution 4.0 International License.
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**Abstract**

In this study, we examined the epiplastic microalgal community that developed on low-density polyethylene (LD-PE) plastic debris found in two distinct regions of the Monastir Bay (Tunisia): the coast exposed to anthropogenic discharges and the open sea in front of the Kuriat Islands. The analysis of trace elements accumulated in the plastisphere was also carried out. The collected plastispheres were predominantly composed of cyanobacteria. Chlorophytes and diatoms were also found. Euglenophyceae was also recorded but with comparatively low relative abundance (RA), whereas dinoflagellates were almost absent. The diatom showed a high diversity of species (25 species). Potentially harmful microalgae (PHM) were more abundant in the plastisphere than in seawater at all stations. For all coastal area stations, most of the PHM colonizing the plastisphere are potentially harmful cyanobacteria. For S1 (offshore station), the PHM community profile was characterized by the dominance of potentially harmful diatoms. The main potentially harmful cyanobacterium identified on the plastisphere was *Phormidium* sp. *Pseudo-nitzschia* spp. was the only potentially harmful diatom observed on the plastisphere. Accumulation of trace elements in plastisphere was higher than that in sediments. Trace elements in plastisphere are ranked with a very high contamination factor at all sites, according to the sequence Pb > Cu > Cd > Ni > Zn. The accumulation of trace elements in the plastisphere is highest near areas with harbor and industrial activities and the urban rejects zone.

**Introduction**

The Mediterranean Sea has been described as the sixth gyre area of marine litter accumulation (Cózar et al. 2015) with plastic debris averaging 423 g/km². Indeed, due to surface currents entering into the Mediterranean Sea via the Strait of Gibraltar, while deep currents exit via the Atlantic Ocean (Ben Ismail et al. 2012), plastic debris (23,150 tons) are trapped in the basin and carried by currents in a closed loop (Eriksen et al. 2014).

Currently, between 1,000 and 3,000 tons of plastic debris float on the surface of the Mediterranean (fragments of bottles, bags, packaging, fishing lines...). According to a study by the World Wide Fund for Nature (WWF), Tunisia is the fourth largest per capita consumer of plastic products in the Mediterranean. In 2016, Tunisia generated 0.25 million tons of plastic waste, of which 0.05 million tons (20%) was uncollected and 0.20 million tons (80%) was collected for treatment. About 0.15 million tons (60%) of this waste was sent to landfills; 0.04 million tons (16%) was openly discarded; and only 0.01 million tons (4%) was recycled. The Tunisian economy loses about $20 million annually due to plastic pollution, which affects tourism, shipping, and fishing (World Wildlife Fund report, 2019).

Moreover, environmentalists had warned that, in the years of this global pandemic, the plastic pollution from masks and gloves could pose a severe threat to the marine environment and life as the pollution mounting rapidly (Amaral-Zettler et al. 2020; Das et al. 2021). For example, biofilms (bacteria, viruses, phytoplankton, micropredators) and marine invertebrates use plastic debris of various sizes (macro-, meso-, and microplastics) as a habitat. These substrates have also been shown to bind pathogens
(Zettler et al. 2013; Amaral-Zettler 2020) and chemical, metallic, and organic contaminants that accumulate into the food chain after ingestion by aquatic organisms, eventually causing infections, strangulation, internal injuries, and hormonal disruption (Andrady 2011; Rochman et al. 2013).

Biofilms deposited by algae or even crustaceans on large floating debris increase the density of plastic particles which promote the sedimentation of this mobile "plastisphere" into deep layers and ultimately threaten the biological benthic activity (Stefatos et al. 1999; Zettler et al. 2013). Recent studies have confirmed not only the colonization of plastic surfaces by specific communities compared to those in the surrounding seawater (Zettler et al. 2013; Amaral-Zettler 2020), but also those in the sediment (Harrison et al. 2014; Reisser et al. 2014; Oberbeckmann et al. 2016; De Tender et al. 2017 and Debroas et al. 2017). They have also reported the presence of invasive species, potentially toxic microalgae (Masó et al. 2003; Masó et al. 2016; Casabianca et al. 2019), marine insect eggs (Ribeiro-Brasil et al. 2022) and halobates (Zettler et al. 2011).

Plastic pollution is a universal problem whose solution requires universal commitments.

Although scientific research on the quantity, distribution, degradation and sequestration processes of plastic debris and characterization of microbial communities that colonize plastic debris (macro-, meso- and micro-organisms) has increased spectacularly in recent decades, studies remain limited to the European continent and some countries in America and Asia, and none from Africa. Furthermore, just a few researchers have looked at communities of environmentally collected plastic in the coastal and/or open ocean; the majority have concentrated on incubation experiments.

This study is the first investigation of the plastisphere that has been developed on marine plastic debris in Tunisia, a country at the northern coast of Africa and southern area of the Mediterranean Sea. Located in the eastern shore of Tunisia, Monastir Bay is a semi-enclosed ecosystem with weak hydrodynamic regime due to its sub-marine topography (Souissi et al. 2014), which makes it a suitable environment for plastic debris contamination and plastisphere development.

In this work we aimed to (i) identify the microalgal community attached to the plastisphere along with comparing its structure to that of the free microalgal community of the surrounding seawater and (ii) evaluate the capacity of the plastisphere to accumulate trace elements along with that of the marine sediment.

# Material And Methods

## Study site

Monastir Bay is considered as a semi-enclosed area, bounded on the northwest by the Fast Island of Monastir and on the southeast by the shoals connecting the Fast Island of Thapsus with the Kuriat Islands (Fig. 1).
Monastir Bay is the seat of increasing urbanization. The population density (approximately 600,000 inhabitants in 2018) is one of the highest in the country. Several buildings, municipalities, industries, and sewage treatment plants line the bay coastline, exerting strong environmental pressure on it. Currently, there are 500 companies in the textile sector, including about 35 specialized in the chemical treatment of fabrics (especially jeans), scattered throughout the region. It is also the site of a large-scale coastal development and important fishing activities with 4 fishing ports: the fishing ports of Monastir, Ksibet El Mediouni, Sayada, and Teboulba. In addition, aquaculture is well developed in the area and has a significant environmental impact on the bay (Challouf et al. 2018; Damak et al. 2020). Monastir Bay has become the most important offshore aquaculture area, where all farms raise perch and sea bream.

## Sampling

In Monastir Bay, eight sampling sites (S1-S8) were chosen: seven (S2-S8) along the coast and one (S1) in the internal offshore area. Three different matrices were selected for sampling, namely coastal seawater, plastisphere, and sediment.

From the coastal seawater, 3 L were collected: 2 L were utilized for chlorophyll a and nutrient analysis; and 1 L was utilized for microalgae enumeration. A 3‰ formaldehyde solution was used to preserve the microalgae samples. For each station, between 3 and 6 white LD-PE (low-density polyethylene # 4) plastic bags floating on the sea surface and exhibiting a "mature" plastisphere (green-yellow patches conspicuous to the eye) were collected using a stainless steel grab. Each plastic bag is cut into two pieces with stainless steel scissors. Two batches were obtained for each sample. The first batch was placed in a glass bottle containing sterile seawater previously filtered onto a 0.22 µm porosity filter. This batch was preserved with a 3‰ formaldehyde solution and used to count microalgae in the plastisphere. The second batch was placed into a glass bottle previously rinsed with HCl solution (1M) and Milli-Q water. To stop biological reactions, three drops of high purity nitric acid was added to each bottle for the second batch. This batch was used for trace element analysis on the plastisphere. Three sediment replicates with cylindrical plastic cores (7 cm in diameter and 10 cm long) were collected at each site. The top 3 cm of the core were carefully cut off and placed in a plastic container for trace element analysis. All samples were stored in the dark at 4°C before being taken to the laboratory.

During sample collection, all relevant parameters were recorded including coordinates, GPS, and weather conditions. Chemical parameters of coastal water (temperature, pH, salinity, and dissolved oxygen (DO) were recorded *in situ* at each station using a multiparameter kit (Multi 3400 i/SET; sensitivity (± 1digit) especially important for pH (0.01).

## Nutrients and chlorophyll a analysis

*Nutrient analyses were performed using the BRAN and the LUEBBE type III Autoanalyzer (APHA, 1992). 1 L of seawater*
adjacent to each LDPE plastisphere sample was collected,
then filtered through a fiberglass filter (Whatmann GF/F)
and stored at -20°C until analysis. The determination of
nutrient concentrations was performed using a colorimetric
assay according to the method of Grasshoff (1983). The
quantification of total chlorophyll a concentration was
performed as described by Lorenzen et al. (1965) and
Aminot and Chaussepied (1983). The absorbance of
pigment extract was measured at 665 nm and 750 nm both
before and after acidification. The equation used was: [Chl
a] µg/L = 26.7 × (λ 665 - λ 750) - (λ' 665 - λ' 750) × 10 / V,
where λ: absorbance before acidification and λ':
absorbance after acidification.

Plastisphere extraction

Different techniques were used to detach the plastisphere from the plastic surface: successive washes with a spray of filtered seawater, ultrasound baths at different intensities, and manual scrapings with glass slides, toothbrushes, or cytological brushes. To evaluate the effectiveness of each technique, observations of the plastic surface before and after were made using optical and electron microscopy. The microscopic observations revealed that brushing with a cytological brush is the most efficient and least efficient (Fig. 2).

FTIR-ATR analysis

An Attenuated total reflectance (ATR) and Fourier transform infrared (FTIR), study, was conducted to confirm that the debris from the recovered plastic bags is LDPE. Before being subjected to ATR-FTIR analysis, the samples were cleaned with a clean room towel dampened with deionized water from an LDPE bottle (Jung et al. 2018).

The ATR-FTIR spectra of the plastic debris were examined using a 45° incidence Bruker FTIR spectrometer (Equinox 55, Bruker Co., Ettlingen, Germany). Spectra were recorded between 4000 and 400 cm⁻¹, with 32 scans total and a resolution of 4 cm⁻¹. Peak height algorithms were used to identify the absorption bands, which were then recorded and compared to the literature-reported absorption bands for each polymer (Chércoles Asensio et al. 2009; Jung et al. 2018; and Nandiyanto et al. 2019).

SEM-EDX Analysis
Scanning electron microscopy with energy dispersive X-ray was used to observe the microalgal community growing on the plastisphere and to evaluate the effectiveness of plastisphere extraction techniques. To prepare samples for SEM the methodology described by Kirstein et al. (2018) was used with some modifications. Briefly, 0.5 cm² strips of LD-PE covered with plastisphere were fixed at 4°C for 10 days in phosphate buffer solution (0.1M), pH = 7.2, containing 3% glutaraldehyde. Strips were dehydrated in ethanol baths of increasing concentration (30%, 50%, 70%, and 90%) for 10 min, followed by three 100% baths. Samples were dried under a vacuum using a paddle pump and then observed using a Q 250 Analytical SEM (Thermo Scientific™), which gives qualitative and quantitative information as to elemental composition.

## Microalgae: morphotaxonomic characterization and cells enumeration

To recover the epiplastic microalgae from the plastisphere, we used 10 g of LD-PE covered with plastisphere for each sample. The sample was placed into a glass crystallizer, and 200 ml of seawater filtered through a 0.22 µm filter was added. The sample was gently scraped off with a sterile cytologic brush. After detachment, a volume of filtered seawater was added to the plastisphere solution to reach a final volume of 1L. Recovered samples were stored in a 3‰ solution of formaldehyde in darkness and at room temperature for subsequent enumeration.

Morphotaxonomic characterization ensures microalgal enumeration in both matrices, thus allowing quantification of groups and species along with the calculation of relative abundance to compare the variation between the two matrices and among different stations. We used the classical microalgal identification technique (Uthermöhl 1958) to determine the composition of the microalgal community growing on the plastisphere and in seawater. Samples were counted with an inverted microscope (Nikon Eclipse TS100) at a final magnification of 200 x. Cell abundance was expressed per g fresh weight for plastisphere (cells $g^{-1}$ fw) and per litter (cells $L^{-1}$) of seawater. Microalgal taxonomy was made according to identification keys (Hasle et al. 1996; Komárek et Anagnostidis 2005).

## Trace element analysis

After being oven dried at 50°C, the samples were homogenized and sieved through a 2 mm mesh screen. To determine total trace elements, 0.5 g of plastisphere and 0.5 g of sediment samples were digested completely with a mixture of concentrated solutions of HF, $HClO_4$ and $HNO_3$, as described by Mudroch et al. (1996). The final solution was diluted to 100 mL in volume. The concentrations of Cd, Co, Cu, Cr, Ni, Pb and Zn were determined using ICP-OES (Perkin Elmer ICP optima 8000). The ICP-OES was calibrated using multi-element calibration standard (Perkin Elmer). The two certified reference materials, BCR-32 (Natural Moroccan Phosphate Rock: Phosphorite) and BCR 414 (plankton) were used to estimate the recovery of the method. The measured recovery rates were between 91.50% and 101.90% for the BCR 32 and between 93.07% and 106.29% for the BCR 414. Each sample was analysed 3 times to verify the accuracy of the instrument. The relative standard deviation of the three replicates was calculated. For all elements the RSDs were between 0.16% and 4.9% (Table 1).
# Geoaccumulation Index

In coastal area use of Geoaccumulation Index (Igeo) introduced by Müller and Suess (1979), was calculated using the equation:

$$Igeo = \log_2 (C_m/ 1.5 C_b).$$

Where C_m is the metal concentration and C_b is the geochemical background concentration of the metal (Turekian and Wedepohl 1961). The Igeo values are classified into seven categories, each representing a degree of pollution: Igeo < 0: no pollution; 0 < Igeo < 1: no pollution or moderate pollution; 1 < Igeo < 2: moderate pollution; 2 < Igeo < 3: moderate to strong pollution; 3 < Igeo < 4: heavy pollution; 4 < Igeo < 5: strong to extreme pollution; and Igeo > 5: extreme pollution.

## Contamination factor

To describe the contamination of toxic trace elements in a marine environment, we may define a contamination factor (Cᵢᵣ) accordingly:

$$C_{f}^{i} = C_{m}^{i} / C_{n}^{i}$$

The mean content of the trace elements (Cᵢᵣ for Cd, Pb, Cu, Zn) from plastisphere was calculated in µg/g; Cᵢᵣ = concentration reference value for those metals were taken from distribution of the elements in some major units of the earth's crust, background value or uncontaminated sediment. The classification of the contamination factors was estimated according to the following scale: Cᵢᵣ < 1 - Low contamination factor; 1 < Cᵢᵣ < 3 - Moderate contamination factor; 3 < Cᵢᵣ < 6 - Considerable contamination factor; 6 ≥ Cᵢᵣ - Very high contamination factor

## Statistical analysis

The data were logarithmically transformed with log (X + 1) prior to statistical analysis. The data was presented as the mean ± standard deviation. The considered variables were tested using Pearson's coefficient. The results were statistically verified using the STATISTICA 12 software package. For all statistical purposes, the criterion of significance was set at P < 0.05.

# Results And Discussion

## Physicochemical and nutriments characteristics

The results of the physicochemical parameters and nutrients in Monastir Bay were presented in Table 2. Temperature, pH, and salinity indicated very close and similar concentrations between all stations. The average temperature was 30.06 ± 2.7°C, the pH was 8.34 ± 0.79, salinity ranged from 37.7 ± 3.39 mg/l to 38.9 ± 3.89 mg/l. Temperature plays a role in the settlement and growth of the colonizing community, with higher temperature leading to stimulate cell metabolism and thus the rapid development of the
attached organisms (Nava and Leoni, 2021). Similarly, salinity values recorded in Monastir Bay ranged within those known to promote biological development (Dussud et al. 2018b; Kesy et al. 2019; Oberbeckmann et al. 2018). Concerning oxygen concentrations, stations S3, S4, S5 and S6 (extending from Khniss to Lamta) exhibit the lowest values with 14.50 ± 1.16, 13.10 ± 1.31, 13.60 ± 0.95 and 13.60 ± 1.08 mg/l, respectively. The rest of the stations showed values around 16 mg/l, mainly Kuriat Island (S1) and Ras Dimas (S8), with 16.80 ± 1.68 mg/l. This variability in oxygen concentrations among stations will accordingly influence metals bioavailability within the water sediment interface (Zhang et al. 2014; Zaaboub et al. 2015).

Concerning nutrients, stations S4 (in front of Ksibat El Mediouni) and S5 (in front of the wastewater of Lamta) exhibited significant values compared to the other stations. The highest concentrations of $NO_2^-$ (4.59 ± 0.4 µmol/l); $NO_3^-$ (15.49 ± 1.54 µmol/l); silicates (10.54 ± 1.05 µmol/l); total nitrogen (39.45 ± 3.94 µmol/l) and total phosphorus (16.04 ± 1.60 µmol/l) are recorded at S5. The enhanced nitrogen concentrations may reflect the intense mineralization of organic matter since Monastir Bay is a receptacle of huge urban and industrial discharges (Ben Amor et al. 2020). This anthropogenic interference may also explain the increased of phosphate concentrations at S4. When combined to both temperature and salinity, nutrients can strongly shape the plastic-attached community with greater biodiversity usually associated with high nutrient levels (Oberbeckmann et al. 2018; Li et al. 2019).

Chlrophyll a distribution illustrates that Khniss station (S3) registered the highest concentration with 6,81 ± 0.68 mg/m³, followed by S5 in front of the wastewater Lamta with a value of 5.60 ± 0.50 mg/m³ and S4 with a value of 4.98 ± 0.44 mg/m³. The stations with low oxygen levels (S3, S4, and S5) exhibited the highest concentrations of Chl a and nutrients since they receive intense waste inputs from domestic wastewater discharges, which promote bacterial oxygen consumption. Our suggestions may be supported by other studies on Monastir Bay (Sassi et al. 1998; Souissi et al. 2014; Damak et al. 2019) showing the importance of eutrophication observed between Khniss (S3) and Ksibat El Mediouni (S4) with anoxic conditions and very high levels of nutrients. We infer that all combined these factors will contribute to shaping the platisphere and determine the accumulation rate of metals (Nava and Leoni 2021).

## FTIR-ATR

The infrared spectra and absorption bands of the LDPE plastic bag labelled with resin code 4 (control) and the LDPE plastic bag debris collected from all stations (Fig. 3) show that the corresponding spectra exhibit a general allure similar to LDPE (Jung et al. 2018). This allure is characterized by asymmetric stretching of $CH_2$ at 2920 cm⁻¹, symmetric stretching of $CH_2$ at 2845 cm⁻¹, bending deformation of $CH_2$ at 1465 cm⁻¹, symmetric deformation of $CH_3$ at 1377 cm⁻¹, and rocking deformation of $CH_2$ at 717 cm⁻¹.

In particular, the spectra of stations S3 to S8 showed the formation of two new functional groups. The first is represented by a shoulder in the region of 1680–1630 cm⁻¹. This corresponds to amide, a form of
carbonyl group (Nandiyanto et al. 2019). The second is represented by a shoulder in the 1100–1070 cm⁻¹ region and a peak at 825 cm⁻¹. This corresponds to C-O stretching and C-H stretching, respectively, a form of oxy compound (Coates, 2006). The formation of a carbonyl compound and oxy compounds is a sign of oxidation of LDPE (Rajandes et al. 2012). It is noteworthy that the shoulder in the 1100–1070 cm⁻¹ range can be attributed to expanded polystyrene (EPS) secreted by microorganisms. Indeed, FTIR spectra in the 900–1300 cm⁻¹ range are characteristic of functional groups in polysaccharides and nucleic acids (Brian-Jaisson et al. 2016).

## SEM-EDX analysis

Scanning Electron Microscopes (SEM) with an Energy dispersive X-ray (EDX) microphotograph highlights the richness and the diversity in the shapes and sizes of prokaryotic and eukaryotic microorganisms living on the plastisphere along the coast of Monastir Bay. For example, we found rod-shaped, spherical, and coccoid bacteria, filamentous cyanobacteria, and centric and araphid pennate diatoms (Fig. 4). However, no dinoflagellates were found using SEM, presumably because they were washed away during SEM processing. The fibrous and mucilaginous structures of extracellular polymeric substances secreted by microorganisms can also be seen. Indeed, EPS is a necessary compound for the formation of the plastisphere and can be referred to as "the cement" of this new habitat. EPS ensures cell attachment to their plastic support and nutrient chelation for microorganism growth.

Moreover, SEM-EDX is also useful to confirm the effectiveness of the plastisphere extraction technique. Indeed, through brushing the extracted plastisphere (Fig. 2) will be easily subjected to assessment of the diversity of the epiplastic microalgal community and the quantification of the accumulated trace elements. For example, the elemental analysis results show that the amount of metal traces in the brushed plastic was zero after plastisphere extraction. SEM has been already used by Zettler et al. (2013) to characterize plastisphere-colonizing organisms. Soon after, scientists used SEM to track the progression and kinetics of plastisphere formation (Amaral-zettler et al. 2020; Ramsperger et al. 2020 and Cheng et al. 2021).

This technique provides a complete description of microorganism diversity and activity during the three stages of biofilm formation on plastics (primo-colonization, growing, and maturation phases). In this sense, our preliminary results of the elemental analysis and the concentration of trace elements determined by ICP-OES showed a good correlation between the mass percent of elements and the concentration of heavy metals (Data not displayed). These results are encouraging as to explore the possibility of using elemental analysis to follow the accumulation kinetics of heavy metals on the plastisphere.

## Microalgae: community composition, morpho-taxonomic characterization and cell counts in the two matrices: seawater and plastisphere
Several parameters influence the composition and structure of the community of microorganisms that harbour plastic debris. Among these parameters, the plastic size, the habitat, the physicochemical quality of the water, the type of plastics, the successive phases of maturation of the biofilm and even the sampling period (De Tender et al. 2015; Oberbeckmann et al. 2016; Hoellein et al. 2017; Frère et al. 2018).

Moreover, different modern techniques are currently used to study the structural complexity of microbial communities growing on the plastisphere. Some scientists have used variety molecular methods (Clone-libraries, Denaturing Gradient Gel Electrophoresis System - DGGE, Amplicon sequencing, and Shotgun metagenomics). Others used microscopic techniques such as SEM and light microscopy coupled with FISH. Each of these methods has its advantages (good taxonomic, multiple samples analysed simultaneously) and disadvantages (a minor part of microorganism diversity identified, expensive instrumentation, and limit in the taxonomic resolution) (De Tender et al. 2017).

To investigate the composition and the spatial structure of the microalgae community growing on the plastisphere of plastic debris collected in July 2020 in the coastal marine waters of Monastir Bay, we limited our work to white low-density polyethylene (LD-PE; floating plastic), and we used the classical techniques of microalgae identification based on microalgal cells counting by a reverse-phase microscope.

Morphotaxonomic characterization ensures enumeration of microalgal cells in both matrices, which facilitates group and species quantification. It also allows the calculation of relative abundance to compare the variation between the two matrices and between different stations.

To analyse and compare the microalgae community present in the two matrices: plastisphere (P) and seawater (SW), the relative abundance (RA) of the different microalgae groups (cyanobacteria, diatoms, chlorophytes, dinoflagellates and euglenophyceae) to total microalgae was calculated (Fig. 5a).

The redundancy analysis (RA) shows dissimilarities between the different groups in the two matrices, which confirm the specificity of the community structure living on the plastisphere, compared to seawater and revealed the spatial variation of the microalgae community according to the sampling stations. In parallel, the morphological-taxonomic characterization revealed the specificity of the group structures formed in the plastisphere compared to seawater.

The matrix sea water (SW) was composed of different groups, i.e., cyanobacteria, diatoms, and dinoflagellates. Euglenophyceae were also observed with a relatively low relative abundance (RA) compared to the other groups, while Chlorophyta were absent. The highest RA for cyanobacteria (91%) was observed at station S2, while the highest RA for diatoms (79.03%) was observed at S1. For dinoflagellates, the highest RA (49.74%) was found at S8.

The matrix plastisphere (P) was predominantly composed of cyanobacteria. Six of the eight sampling stations (S2-S4; S6-S8) had an RA > 50%. The highest RAs for cyanobacteria (99.35%, 96.80%, and 91.80%) were observed at stations S8, S7, and S6, respectively. As has been depicted in Fig. 5b, the
cyanobacterial population, which developed on the plastisphere, is more diversified than that the surrounding seawater. Indeed, 16 dominant cyanobacteria strains were observed in the plastisphere (*Anabaena* sp., *Calothrix* sp., *Chroococcus* sp., *Closterium* sp., *Coelosphaerium* sp., *Gloeocystis* sp., *Gomphosphaeria* sp., *Leptolyngbya* sp., *Limnothrix* sp., *Lungbya majuscula*, *Lyngbya* sp., *Merismopedia* sp., *Phormidium* sp., *Planktothrix* sp., *Rivularia* sp., *Tolypothrix* sp.) versus only 5 in the surrounding waters.

The RA analysis of cyanobacteria species both matrices P and SW shows that the seawater was dominated by the spherical *Merismopedia* sp. for stations S1 (99.65%), S2 (86.61%), S6 (80%), and S7 (70.58%) with an average abundance of 5.40 x 10³ cells L⁻¹ (Table 3). While the filamentous cyanobacterium *Anabaena* sp. dominated the two stations, S1 and S8, with RAs equal to 100%. At stations S3, S4, and S5 potentially harmful cyanobacteria were observed with the dominance of *Lymgbya majuscule* with an average abundance of 5.40 x 10³ cells L⁻¹. *Lymgbya majuscule* has the highest RA (69.23%) at station S4.

In the plastisphere, the number of cyanobacterial species varied from 3 to 8, depending on the station. Although the white LD-PE was the support for all the plastispheres investigated in this work, the taxonomic composition was highly variable among stations. There is no clear dominance of one or two species, as was the case for the seawater matrix. *Anabaena* sp. was the only cyanobacterium observed on all the collected plastispheres with an average abundance of 3.25 x 10⁷ cells g⁻¹. Its RA varied from 1.3% for station S7 to 41.66% for station S4. Some cyanobacteria species are observed only on one plastisphere. *Calothrix* sp., *Closterium* sp., *Leptolyngbya* sp. and *Tolypothrix* sp. are only observed on the plastisphere of station S3. While, *Gloeocystis* sp. and *Gomphosphaeria* sp., are only present on the plastisphere of stations S8 and S6, respectively.

Our analysis also shows that the eight plastispheres collected were hosted by the potentially harmful cyanobacteria: *Lungbya majuscula* and *Phormidium* sp. with a clear dominance of *Phormidium* sp. with an average abundance of 3.24 x 10⁷ cells g⁻¹. Our results are consistent with the earlier observations in marine studies that identified cyanobacteria as one of the most common photoautotrophic prokaryotes found on marine plastic debris (Zettler et al. 2013; Oberbeckmann et al. 2014; Bryant et al. 2016; Oberbeckmann et al. 2016; De Tender et al. 2017; Dussud et al. 2018a; Muthukrishnan et al. 2019). Yokota et al. (2017) explained the important role of cyanobacteria in ensuring the buoyancy of plastic debris. Indeed, cyanobacteria endowed with gas vacuoles can temporarily increase the overall buoyancy of plastic debris until they die or are displaced or consumed by higher density organisms.

The two other groups namely, chlorophytes and diatoms were also found on the plastisphere matrix. The occurrence of Euglenophyceae was also recorded with a relatively low RA. The highest RA for Euglenophyceae (11.98%) was registered for station S4. Chlorophyceae were recorded from S1 to S5. The highest RA for Chlorophyta (89.27%) was noted for station S1 near the Kuriat Islands. The two Chlorophyceae found on are two non-mobile green microalgae found in freshwater environments, namely *Scenedesmus* sp. and *Pediastrum* sp. Their abundance averaged 2.16 x 10⁵ and 1.3 x 10⁵ cells g⁻¹,
respectively. Chlorophycea have rarely been observed on the plastisphere, with a few data reported in both freshwater and seawater plastispheres (Zettler et al. 2013; Kettner et al. 2017; Weig et al. 2021).

Our findings show that dinoflagellates are almost absent (an RA ≤ 0.90%) in all the sampling plastispheres. Regardless of the sampling stations, the analysis of the relative abundance of dinoflagellate species (Fig. 5c) demonstrated that the seawater had a dinoflagellate "mosaic" of 26 species, whereas the plastisphere was characterized by low species diversity (only 4 species and unknown Dinocystis). As can be seen, the plastispheres collected at stations S2, S4, and S8 do not contain dinoflagellate species. Although dinoflagellates made up a larger portion of the community throughout the water column, these taxa became more common in the plastispheres during the course of the time series (Masó et al. 2003; Zetter et al. 2013; Reisser et al. 2014; Kettner et al. 2017; Dudek et al. 2020). However, not all dinoflagellates were related to both plastic-attached and water column communities.

The taxonomic composition of the dinoflagellates was dissimilar to that of the plastisphere and the surrounding seawater. For example, for station S1, the seawater matrix is composed of 9 species of dinoflagellates: *Alexandrium minutum*, *Alexandrium pseudogonyaulax*, *Gymnodinium catenatum*, *Peridinium* sp., *Polykrikos* sp., *Prorocentrum cordatum*, *Prorocentrum rathymum*, *Protoperidinium minutum*, and *Protoperidinium pyriforme*, and a few hundred dinocysts with an RA = 5%, while the plastisphere matrix contains only Indeterminate dinocysts. Resting cystis of unidentified dinoflagellates and both temporary cysts and vegetative cells of *Alexandrium taylori* were enumerated on floating plastic debris collected in summer during a bloom of harmful algae (HAB) *Alexandrium taylori* along the Catalan coast (north-western Mediterranean) (Masó et al. 2003). The production of temporary cysts has typically been linked to unfavourable environmental factors. This enables the population to endure the worst possible weather circumstances and survive. According to Garcés et al. (2002), stickiness may also be important for the dispersal of phytoplankton species that have an adhesive stage in their life cycle.

The planktonic species *Alexandrium minutum* and the epibenthic species *Coolia monotis*, *Prorocentrum concavum* and *Prorocentrum lima* are the only dinoflagellates species found on the plastisphere with an average abundance of 6.00 x 10², 1.2 x 10³, 1.5 x 10² and 1.8 x 10³ cells g⁻¹, respectively. All dinoflagellate species found on the collected plastisphere are autotrophic. However, Dudek et al. (2020) demonstrated that the plastisphere communities observed in the Caribbean coastal site were dominated by the heterotrophic dinoflagellate *Amphidinium* sp. at weeks 1 and 3, but shifted to the autotrophic strains *Prorocentrum* sp. and *Alexandrium* sp. at week 6 of incubation. This means that the population dynamics of the dinoflagellate community fluctuate with time. Dinoflagellates include many species inhabiting various aquatic ecosystems. Their nutritional mode (autotrophic, mixotrophic, heterotrophic, symbiotic or parasitic) and chemical defences may affect their ecological niches (bottom of water bodies: the benthos or the water column: the plankton). Heterotrophic species constituted 49% of total species. They are also reported to be slightly dominant in marine waters (Gómez 2012). This prevalence of heterotrophic dinoflagellates may be explained by their competitive features. For example, they can be raptors feeding on prey using "temporal gradient sensing" chemotaxis, in which cells move along a
chemical gradient in a directed manner toward the higher concentration. As the extracellular
concentration of the attractant increases, a corresponding increase in the ratio of net to gross
displacement results in an overall movement toward the stimulus (Cancellieri 2001). Moreover,
heterotrophic dinoflagellates are major grazers of diatoms in the sea, and constitute also an important
food resource for mesozooplankton (Sherr and Sherr 2007).

All these observations make the explanation of the presence and/or absence of one or more species of dinoflagellate on the plastisphere a difficult mission. Certainly, their flexibility in the trophic mode, their mobility, as well as their life cycle make them an "intelligent colonizer" and a "crucial piece of the puzzle" to understand the interaction between the different organisms sheltering the plastisphere. Therefore, more research on the dinoflagellate community growing on the plastisphere is needed.

Our results indicate that diatoms were present in all plastispheres, with RAs ranging from 0.19% at S8 to 40% at S5. The diatom group showed a high diversity of species observed on the two matrices: 25 species on the plastisphere and 23 species in the seawater (Fig. 5d). Some species were only observed on the plastisphere (*Cocconeis placentula*, *Grammatophora marina*, *Skeletonema costatum*, *Striatella unipunctata*, *Synedra ulna*, and *Pseudo-nitzschia* spp.), while others were only in seawater (*Asterionellopsis glacialis*, *Bacteriastrum delicatulum*, *Coscinodiscus* sp. and *Guinardia* spp.).

Both *Navicula* sp. and *Nitzschia longissima* were observed in both matrices but with different relative abundances. In seawater, the highest RA of *Navicula* sp. (100%) was observed for station S2. For the plastisphere, the highest RA of *Navicula* sp. (58.06%) was observed for station S6. Other studies showed the importance of diatoms dominant colonizers of plastic debris that are exposed to sunlight (Oberbeckmann et al. 2014; Masó et al. 2016; Michels et al. 2018; Kettner et al. 2019; Amaral-Zettler et al. 2020). It was reported that after two weeks of plastic colonization, diatoms multiply on plastic polymers and can reach concentrations up to 10 times higher compared to the surrounding environment (Eich et al. 2015).

The crucial role of diatoms during biofilm formation has already been demonstrated (Patil and Anil 2005). However, the general process of biofilm formation, which starts with electrostatic attraction and repulsion between surfaces and negatively charged bacterial cell walls (Renner and Weibel 2011), may apply to diatoms even though the biological process of diatom adhesion to plastic debris has not yet been thoroughly investigated. In fact, negatively charged diatom cell surfaces have been seen (Konno 1993; Gélabert et al. 2006). The diatoms were more readily observed on the film than the small microplastics. This suggests that the rigidity morphology of diatoms requires a flatter surface area for colonization compared to bacteria (Cheng et al. 2021).

**Potentially Harmful microalgae in the two matrices: seawater and plastisphere**

To better analyse the abundance of potentially harmful microalgae (PHM) in both matrices P and SW, we calculated the mean relative abundance of total PHM (RA<sub>PHM</sub>), the relative abundance of different groups
of potential harmful microalgae (RA<sub>G</sub>) in relation to total PHM, the relative abundance of potential harmful cyanobacteria species in relation to total potential harmful cyanobacteria (RA<sub>Cya</sub>), the relative abundance of the potential harmful dinoflagellate species in relation to total potential harmful dinoflagellates (RA<sub>Din</sub>) and the relative abundance of potential harmful diatoms species in relation to total potential harmful diatoms (RA<sub>Dia</sub>).

The analysis of the results of the mean relative abundance of total PHM (RA<sub>PHM</sub>) in both matrices (Fig. 6) showed that there is a significant difference (p ≤ 0.05) between the stations. On the other hand, PHM are more abundant in the plastisphere than in seawater at all stations. Four of the eight study stations contained plastic debris with plastispheres containing more than 40% PHA. The RA<sub>PHM</sub> ranged from 1.42 ± 0.26 for S1 to 55.82 ± 3.02 for S8. For SW, the highest RA<sub>PHM</sub> did not exceed 22.57 ± 1.10% (station S4).

The mean relative abundances of PHM for the plastisphere are remarkably high for stations S8, S7, and S2. They are 97, 26, and 28 times higher than those for seawater, respectively. These sites are characterized by shallow water, low hydrodynamics and clandestine dumping. Station S1, unlike all others, has low RA<sub>PHM</sub> for both matrices (1.19 ± 0.36% and 1.42 ± 0.26%, in seawater and plastisphere, respectively). Station S1 is located in the open sea of the Monastir Bay, in front of the Kuriat Islands. This station is the deepest (20–25 m depth) and the marine dynamics are quite active, which reduces the amount of plastic floating on its surface and the duration of its stay. Compared to the coastal area, relatively little plastic waste was found during the sampling event in this station. Even the samples that were taken did not reveal a dense plastisphere, which suggests that the debris were just recently thrown out.

The RA<sub>G</sub> analysis (Fig. 7a) reveals that for all coastal area stations (except S1: offshore station, Kuriat Islands), the most PHM colonizing the plastisphere are potentially harmful cyanobacteria. For station S1, the PHM community profile was characterized by the dominance of potentially harmful diatoms with a RA<sub>G</sub> of 81.5%. Filamentous and colonial cyanobacteria are frequently the causes of HAB in inland waters. Their interaction with plastic debris in wastewater effluent may influence HAB formation and persistence (Lee 2008). In fact, filamentous cyanobacteria that grow on the plastisphere of floating plastic can aggregate and contribute to the formation of surface scum. Many HAB-forming cyanobacteria boost buoyancy via gaseous vacuoles (Oliver 1994). This reduces the density of the plastic and slows its sequestration in the sediment. In contrast, when attached to neutrally buoyant plastic debris, specialized cyanobacterial cells such as heterocysts and akinetes (which lack gas vacuoles) can increase the sequestration of its polymers in the sediment. Cyanobacterial cells "hitching" on rapidly sinking plastic may provide a new pathway to escape unfavourable conditions and thus may increase the size of the cyanobacterial "seed bank" in the sediment and possibly the frequency and/or severity of future HABs (Yokota et al. 2017).

The potentially harmful cyanobacteria observed in seawater are mostly composed of the species *Lyngbya majuscule*, except at station S2 where *Phormidium* sp. dominates with a RA<sub>Cya</sub> of 100%.
Whereas, the main potentially harmful cyanobacterium identified on the plastisphere was *Phormidium* sp. (Fig. 7b). The RAcya of *Phormidium* sp. was equal to 100% for S1, S5, and S6. The genus *Phormidium* has been found on the plastisphere of various types of plastic debris from various geographical origins (Zettler et al. 2013; Oberbeckmann et al. 2014; Bryant et al. 2016; Oberbeckmann et al. 2016, Du et al. 2022 and Dey et al. 2022). *Phormidium* and related hydrocarbon-degrading taxa molecular signatures were discovered in plastics by two studies: Zettler et al. (2013) on microplastics collected from the North Atlantic and Oberbeckmann et al. (2014) on PET bottles incubated in the North Sea. This raises the intriguing possibility that the cyanobacteria *Phormidium* that live in the plastisphere actively hydrolyse the plastic rather than relying on inorganic nutrients found in the surrounding environment or released by heterotrophic bacteria (the primo-colonizers of plastisphere) (Harrison et al. 2014; McCormick et al. 2014; Yokota et al. 2017). *Phormidium* has the capacity to grow large benthic mats in lakes and rivers. Since some strains produce the potent neurotoxic metabolites (anatoxin-a, dihydroanatoxin-a, homoanatoxin-a, and dihydrohomoanatoxin-a) also known as anatoxins (Gugger et al. 2005; Heath et al. 2010; Wood et al. 2012). Worldwide, *Phormidium* mats have been linked to several dog poisonings and deaths (Puschner et al. 2010; Faassen, et al. 2012). To the best of our knowledge, few studies have demonstrated the toxicity of marine *Phormidium* strains. Williamson et al. (2002) has highlighted the ability of the marine cyanobacterium *Phormidium* sp. to synthesize a toxin: phormidolide. The latter is very highly toxic to brine shrimp with an LC<sub>50</sub> = 1.5 µM. Phormidolide is one of only a few macrolide-type natural products to be reported from marine cyanobacteria.

The cyanobacterium *Lyngbya majuscula* has also been observed but with low RA<sub>Cya</sub>. The highest RA<sub>Cya</sub> (42.85%) was detected on station S4. The toxicity of marine cyanobacteria *Lyngbya majuscula* has been observed on various marine organisms: *Brine shrimp, Biomphalaria glabrata* mollusc, and *Chelonia mydas* green turtle (Gerwick et al. 1994; Arthure et al. 2008).

Dinoflagellates are the most potentially harmful microalgae found in seawater. While no potential harmful diatoms were found (Fig. 7a). The RA<sub>G</sub> of dinoflagellates was 100% for four stations (S1, S6, S7, and S8) (Fig. 7c). Regardless of sampling stations, 11 strains of potentially harmful dinoflagellates were identified in seawater (*Alexandrium minutum, Alexandrium pseudogonyaulax, Amphidium carteri, Coolia monotis, Gymnodium catenatum, Heterocapsa circularisquama, Ostreopsis* sp. *Prorocentrum concavum, Prorocentrum cordatum, Prorocentrum lima* and *Prorocentrum rathymum*). The dinoflagellate *Alexandrium minutum* is found in 6 stations, while *Ostreopsis* sp. was only observed in station S2.

In contrast to the seawater, potentially harmful dinoflagellates were found in low abundance (3.21%) on the plastisphere. *Prorocentrum lima* is the most dominant. It has been identified in three stations (S3, S6, and S7). Three other species were also identified: *Alexandrium minutum, Coolia monotis*, and *Prorocentrum convacum*. The LD-PE associated dinoflagellates we identified could be associated with HABs.

Species of the genus *Alexandrium* cause paralytic shellfish poisoning in humans. The first evidence of plastic HAB association was described by Masó et al. (2003), who found temporary cysts of *Alexandrium*
*taylori* adhering to plastic bottles. Other potentially toxic dinoflagellates in our plastisphere samples included *Coolia monitis*, considered a potential cause of seafood poisoning (Faust 1995; Lenoir et al. 2004) and the genus *Prorocentrum*, which contains several toxic species, some of which can generate toxins that play a role in ciguatera syndrome.

The second group of potentially harmful microalgae found on the plastisphere are diatoms. Five stations recorded their existence (S1, S2, S6, S7, and S8). The station S1 (Kuriat island) recorded the highest $RA_G$ (81.5%), while the station S8 recorded the lowest (0.62%). Our findings also showed that there were no potentially harmful diatoms in the seawater matrix. *Pseudo-nitzschia* spp. was the only potentially harmful diatom observed on the plastisphere (Fig. 7d). Our results also demonstrated the total absence of potentially toxic diatoms in the seawater matrix. Our results are consistent with other work that has shown the dominance of *Pseudo-nitzschia* spp. in the marine plastisphere (Casabianca et al. 2018; Dudek et al. 2020; Cheng et al. 2021). The diatom genus *Pseudo-nitzschia* is found all over the world and some of its species generate domoic acid (DA), the neurotoxin that leads to amnesic shellfish poisoning. At least three people were killed by this poison in 1987, which led to a plethora of studies on the physiology, distribution, ecology, and identification of *Pseudo-nitzschia* species. Since prior assessments in 1998, more understanding has been obtained regarding the fate of DA, including how marine species accumulate it and how light and microbes break it down. Review of the biotic and abiotic elements that affect DA production is done with an emphasis on how recent findings have altered our initial theories regarding the control mechanisms. Recent research has confirmed that silicate and phosphate deficiency cause the formation of DA. However, recently identified triggers for stress, such as high or low copper or iron concentrations, point to DA's potential function in trace-metal chelation (Lelong et al. 2012).

## Metal accumulation in sediment

Table 4 shows the trace elements in surface sediments with a large difference between the values for contaminant accumulation in the sediment. Sediment has been considered the central accumulator of trace elements in the marine environment (Tessier et al. 1979). The degree of trace metal enrichment in the sediment of the Monastir Bay coastal area can be evaluated using NOAA sediment quality guidelines (SQG) known as effects range-low (ERL) and effects range-median (ERM) (Long and Morgan 1990; Long et al. 1995). This is the enrichment of trace elements above which toxic effects in benthic organisms can be expected with a frequency of at least 50% (ERM), and below which effects can be expected only rarely (ERL). A toxic effect can be detected for Cd in the central part of Monastir Bay (S6), which is known as the outfall of the agricultural zone where pesticides are used, and in the coastal zone, there are sites for ships and ship repairs with the installation of offshore aquaculture cages. This cadmium accumulation in sediment has been observed in recent decades (Khiari et al. 2022).

The Geoaccumulation Index (Igeo) (Table 5), calculated using as background values the trace elements estimations of Turekian and Wedepohl (1961), confirms the extreme Cd pollution at this site and strong to extreme pollution in northern coastal areas S3, S4 and, S5 (Fig. 8a). For Pb, Igeo shows no or moderate pollution in the central part of the bay.
The middle part is known to be derived from waste from boat painting (S3), industrial waste (S4), and water treatment plants (S5); areas indicated in Fig. 1.

The contamination factor is used to evaluate the potential risk of heavy metal accumulation in biological systems (Hakanson 1980). This index is commonly used to evaluate the accumulation of heavy metals (Rutkowski et al. 2020; Yan et al. 2016). This study used this index to assess heavy metals in the plastisphere. The origin of the plastisphere is one of the most important parameters of our study. It should be noted that, the LDPE samples were collected from the tidal fluctuation zone, with some of them in contact with the sediment. Because there is no metal enrichment in the sediment, we used it as the reference background value. Only for Cd, this enrichment is considered.

The enrichment of metals in the plastisphere represents a significant accumulation of trace elements with a contamination factor index higher than 6 (Table 4), indicating that most industrial, port, and aquaculture activities at stations S1, S2, S7, and S8 have a high contamination factor. At station S1 there are aquaculture activities; the other stations are affected by urban and port waste. The average contamination factor at all sites has a higher average value for the following sequence of metals: Pb > Cu > Cd > Ni > Zn. Cobalt is not included in this classification because it occurs only in the central part of the bay (S4 and S5).

A comparative view of metal accumulation in the sediment and plastisphere is shown in Fig. 8. Metal accumulation appears to be influenced by waste from urban industrial areas and ports (S3, S4, and S5). A particular process was observed at station S6, where antifouling paint is used for aquaculture cages in this coastal area. According to Singh and Turner (2009), trace elements Cd, Cr, Ni, and Pb have maximum concentrations in the antifouling residues in the coastal sediment, indicating a heterogeneous source of metallic contaminants in the marine environment where cage cleaning and maintenance activities occur.

Metal accumulation in the plastisphere is most common near Ras Dimas and Ksibet El Madiouni, where there is a specific morphological feature with a very shallow water depth, not exceeding 0.5 m depth and with very low hydrodynamics, as explained by Khiari et al. (2022). As shown in Fig. 8, trace elements in the plastisphere have accumulated mainly in this zone, which is characterized by low bathymetry. Due to the marine barrier (Dahr) and Ras Dimas spit, which have canceled the hydrodynamics in this area, these pollutants, brought by the runoff and released by the port and fishing activities, have been deposited on the plastisphere without any mechanical dynamics (Khiari et al. 2022).

In this context, the interaction of metals with another matrix is the main cause of plastic-metal ion interaction in terms of competitive processes. The contact between plastic, other solid phases, and the water phase was subject to both continental and marine influences in the coastal zone. Water chemistry, especially salinity, pH, temperature, redox conditions, and other dissolved ions, are the main parameters affecting the properties of the plastic matrix and associated elements (organic or inorganic). An important factor in this complex is the degradation rate and biofouling of polymers, which affect the bioavailability of metals, wettability, and reactive surface area (Binda et al. 2021). Environmental processes have a great influence on the way plastics and metals interact and associate.
# The source of trace elements in epiplastic microalgae associations

The accumulation of plastic debris and trace elements in aquatic and terrestrial environments is a major concern. The adsorption of trace elements onto the plastisphere in the marine environment is generally a result of adsorption onto plastic surfaces found in freshwater and terrestrial environments. On the other hand, there is adsorption on plastic debris present in seawater, which is less common as the continental environment is the main source of plastic waste. Plastic debris act as vectors to transport trace elements and can increase the chemical exposure of ingesting organisms and the resulting toxicity (Bank 2022).

Interactions between metals and plastic debris occur through a variety of modes, primarily direct adsorption of cations (physical); other forms of association of metals to the plastisphere by complexation on charged sites or neutral regions of the plastic surface; and complexation on charged sites or neutral regions of the particulate plastic surface that will be the site of chemical co-precipitation. Other types of the metal-plastisphere association include adsorption onto hydrated oxides (Ashton et al. 2010).

## Conclusion

This study, which is the first one carried out on this subject in Africa, has shown that the plastisphere can be considered an "extreme ecosystem" and a "new field of study for bioremediation." In fact, it has demonstrated its great ability of the epiplastic microalgal community to bioaccumulate trace metals at very high concentrations, despite their toxicity. It is thus a "trace metal sponge" community. Moreover, the plastic substrate, which is a synthetic support of hydrocarbons, can develop a diversified community of epiplastic microalgae with the presence of potentially toxic species. This will result in a double toxicity of the plastisphere towards the whole food chain. Cyanobacteria and diatoms are the dominant species in the microalgal community that has been observed on the plastisphere. These typically phototrophic microalgae are well known for playing significant roles in biofilm formation because of their capacity to secrete and produce extracellular polymeric substances (EPS). Isolation and characterization of epiplastic microalgal communities is a prospective research area to investigate the bioremediation potential of these microorganisms because it is well known that heavy metals and these polymers interact. In addition, it is imperative to start prioritizing and identifying processes that occur in the plastisphere, such as biodegradation or pathogenicity. To accurately interpret what is occurring in nature, future studies must include going "back to the bench" and discovering the mechanisms involved in degradation processes.

In the end, "nothing is lost, nothing is created, everything is transformed" (Antoine de Lavoisier, inspired by Anaxagore de Clazomènes).

## Declarations
**Acknowledgements** The authors would like to thank the Deanship of Scientific Research at King Khalid University for funding part of this work through the Large Group RGP. 2/95/43. We express our gratitude to Mrs. Mabrouka Mahfoudi for her cooperation in microalgae enumeration and identification.

**Authors Contributions** All authors contributed to the study conception and design. Mondher Tarchi: Material preparation, investigation, data collection and analysis and first draft writing. Noureddine Zaaboub: Conceptualization, trace element analysis contribution and supervision, Majed Alsubih: data collection, analysis and writing. Bochra Brik: software and formal analysis. Maria Virgínia Alves Martins: review, and editing. Lotfi Aleya: analysis, review, and editing. Lamia Trabelsi: Conceptualization, supervision, validation, writing, review, and editing. All authors read and approved the final manuscript.

**Funding** This study is part of the PhD research work of Mr. Mondher Tarchi. This study was financed by the National Institute of Marine Sciences and Technologies and by the Deanship of Scientific Research at King Khalid University through the Large Group RGP. 2/95/43.

**Availability of data and materials** Data generated or analyzed during this study are included in this published article. Additional datasets are available from the corresponding author on reasonable request.

**Ethical Approval** Not applicable

**Consent to Participate** Not applicable

**Consent to Publish** Not applicable

**Competing Interests** The authors declare no competing interests.

**References**

1. Amaral-Zettler LA, Zettler ER, Mincer TJ (2020) Ecology of the plastisphere. Nat Rev Microbiol 18(3):139–151. https://doi.org/10.1038/s41579-019-0308-0

2. Aminot A, Chaussepied M (1983) Manuel des analyses chimiques en milieu marin. Editions Jouve, CNEXO, Paris, p 395

3. Andrady AL (2011) Microplastics in the marine environment. Mar Pollut Bull 62(8):1596–1605. https://doi.org/10.1016/j.marpolbul.2011.05.030

4. Arthur K, Limpus C, Balazs G, Capper A, Udy J, Shaw G, Keuper-Bennett U, Bennett P (2008) The exposure of green turtles (Chelonia mydas) to tumour promoting compounds produced by the cyanobacterium Lyngbya majuscula and their potential role in the aetiology of fibropapillomatosis. Harmful Algae 7(1):114–125

5. Ashton K, Holmes L, Turner A (2010) Association of metals with plastic production pellets in the marine environment. Mar Pollut Bull 60(11):2050–2055. https://doi.org/10.1016/j.marpolbul.2010.07.014
6. Bank MS (2022) Microplastic in the Environment: Pattern and Process. University of Massachusetts Amherst, MA, USA, Springer edition

7. Ben Amor R, Jerbi H, Abidi M, Gueddari M (2020) Assessment of trace metal contamination, total organic carbon and nutrient accumulation in surface sediments of Monastir Bay (Eastern Tunisia, Mediterranean Sea). Regional Studies in Marine Science https://doi.org/10.1016/j.rsma.2020.101089

8. Ben Ismail S, Sammari C, Gasparini GP, Béranger K, Brahim M, Aleya L (2012) Water masses exchanged through the Channel of Sicily: Evidence for the presence of new water masses on the Tunisian side of the channel. Deep-Sea Research I 63 (2012) 65-81

9. Binda G, Spanu D, Monticelli D, Pozzi A, Bellasi A, Bettinetti R, Carnati S, Nizzetto L (2021) Unfolding the interaction between microplastics and (trace) elements in water: A critical review. Water Research 204

10. Brian-Jaisson F, Molmeret M, Fahs A, Guentas-Dombrowsky L, Culioli G, Blache Y, Cérantola S, Ortalo-Magné A (2016) Characterization and anti-biofilm activity of extracellular polymeric substances produced by the marine biofilm-forming bacterium Pseudoalteromonas ulvae strain TC14. Biofouling 32(5):547-560. https://doi.org/10.1080/08927014.2016.1164845

11. Bryant JA, Clemente TM, Viviani DA, Fong AA, Thomas KA, Kemp P, Karl DM, White AE, DeLong EF (2016) Diversity and Activity of Communities Inhabiting Plastic Debris in the North Pacific Gyre. mSystems 1(3):e00024-16 https://doi.org/10.1128/msystems.00024-16

12. Cancellieri PJ (2001) Chemosensory Attraction of Pfiesteria spp. to Fish Secreta. A thesis submitted to the graduated Faculty of North Carolina State University. Pp64

13. Casabianca S, Capellacci S, Giacobbe MG, Dell'Aversano C, Tartaglione L, Varriale F, Narizzano R, Risso F, Moretto P, Dagnino A, Bertolotto R, Barbone E, Ungaro N, Penna A (2018) Plastic-associated harmful microalgal assemblages in marine environment. Environ Pollut 244:617-626. https://doi.org/10.1016/j.envpol.2018.09.110

14. Challouf R, Hamza A, Yahia A, Ghozzi K, Saidi B, Bradai MN (2018) Assessing Potential Impacts of Offshore Fish Farming on Small-Scale Fishery in Monastir Bay (Eastern Shore of Tunisia). Saudi J Life Sci (SJLS). DOI:10.21276/haya.2018.3.3.11

15. Cheng J, Jacquin J, Conan P, Pujo-Pay M, Barbe V, George M, Fabre P, Bruzaud S, Ter Halle A, Meistertzheim AL, Ghiglione JF (2021) Relative Influence of Plastic Debris Size and Shape, Chemical Composition and Phytoplankton-Bacteria Interactions in Driving Seawater Plastisphere Abundance, Diversity and Activity. Front Microbiol 11:610231. https://doi.org/10.3389/fmicb.2020.610231

16. Chércoles Asensio R, San Andrés Moya M, de la Roja JM, Gómez M (2009) Analytical characterization of polymers used in conservation and restoration by ATR-FTIR spectroscopy. Anal Bioanal Chem 395(7):2081-2096. https://doi.org/10.1007/s00216-009-3201-2

17. Coates J (2006) Interpretation of Infrared Spectra, A Practical Approach. Encyclopedia of Analytical Chemistry https://doi.org/10.1002/9780470027318.a5606

18. Cózar A, Sanz-Martín M, Martí E, González-Gordillo JI, Ubeda B, Gálvez J, Irigoien X, Duarte CM (2015) Plastic accumulation in the Mediterranean sea. PLoS ONE 10(4):e0121762.
19. Damak M, Fourati R, Elleuch B, Kallel M (2020) Environmental quality assessment of the fish farms' impact in the Monastir Bay (eastern of Tunisia, Central Mediterranean): a benthic foraminiferal perspective. Environ Sci Pollut Res 27:9059-9074. https://doi.org/10.1007/s11356-019-07523-7

20. Damak M, Frontalinic F, Elleuch B, Kalle IM (2019) Benthic foraminiferal assemblages as pollution proxies along the coastal fringe of the Monastir Bay (Tunisia). J Afr Earth Sci 150:379-388. doi:10.1016/j.jafrearsci.2018.11.013

21. Das S, Vera M, Gandin V, Singer RH, Tutucci E (2021) Intracellular mRNA transport and localized translation. Nat Rev Mol Cell Biol 22(7):483-504. https://doi.org/10.1038/s41580-021-00356-8

22. Debroas D, Mone A, Ter Halle A (2017) Plastics in the North Atlantic garbage patch: A boat-microbe for hitchhikers and plastic degraders. Sci Total Environ 599-600:1222-1232. https://doi.org/10.1016/j.scitotenv.2017.05.059

23. De Tender C, Devriese LI, Haegeman A, Maes S, Vangeyte J, Cattrijsse A, Dawyndt P, Ruttink T (2017) Temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea. Environ Sci Technol 51:7350-7360. https://doi.org/10.1021/acs.est.7b00697

24. De Tender CA, Devriese LI, Haegeman A, Maes S, Ruttink T, Dawyndt P (2015) Bacterial Community Profiling of Plastic Litter in the Belgian Part of the North Sea. Environ Sci Technol 49(16):9629-9638. https://doi.org/10.1021/acs.est.5b01093

25. Dey S, Rout AK, Behera BK, Ghosh K (2022) Plastisphere community assemblage of aquatic environment: plastic-microbe interaction, role in degradation and characterization technologies. Environ Microbiome 17:32. https://doi.org/10.1186/s40793-022-00430-4

26. Du Y, Liu X, Dong X, Yin Z (2022) A review on marine plastisphere: biodiversity, formation, and role in degradation. Comput Struct Biotechnol J 20:975-988 https://doi.org/10.1016/j.csbj.2022.02.008

27. Dudek KL, Cruz BN, Polidoro B, Neuer S (2020) Microbial colonization of microplastics in the Caribbean Sea. Limnol Oceanogr Lett 5:5-17. https://doi.org/10.1002/lol2.10141

28. Dussud C, Hudec C, George M, Fabre P, Higgs P, Bruzaud S, Delort AM, Eyheraguibel B, Meistertzheim AL, Jacquin J, Cheng J, Callac N, Odobel C, Rabouille S, Ghiglione JF (2018a) Colonization of Non-biodegradable and Biodegradable Plastics by Marine Microorganisms. Front Microbiol 9:1571. https://doi.org/10.3389/fmicb.2018.01571

29. Dussud C, Meistertzheim AL, Conan P, Pujo-Pay M, George M, Fabre P, Coudane J, Higgs P, Elineau A, Pedrotti ML, Gorsky G, Ghiglione JF (2018b) Evidence of niche partitioning among bacteria living on plastics, organic particles and surrounding seawaters. Environ Pollut 236:807-816. https://doi.org/10.1016/j.envpol.2017.12.027

30. Eich A, Mildenberger T, Laforsch C, Weber M (2015) Biofilm and Diatom Succession on Polyethylene (PE) and Biodegradable Plastic Bags in Two Marine Habitats: Early Signs of Degradation in the Pelagic and Benthic Zone? PLoS ONE 10(9):e0137201. https://doi.org/10.1371/journal.pone.0137201
31. Eriksen M, Lebreton LC, Carson HS, Thiel M, Moore CJ, Borerro JC, Galgani F, Ryan PG, Reisser J (2014) Plastic Pollution in the World's Oceans: More than 5 Trillion Plastic Pieces Weighing over 250,000 Tons Afloat at Sea. PLoS One 9(12):e111913.
https://doi.org/10.1371/journal.pone.0111913

32. Faassen EJ, Harkema L, Begeman L, Lurling (2012) First report of (homo)anatoxin-a and dog neurotoxicosis after ingestion of benthic cyanobacteria in The Netherlands. Toxicon 60(3):378-384.
https://doi.org/10.1016/j.toxicon.2012.04.335

33. Faust MA (1995) Observation of sand-dwelling toxic dinoflagellates (Dinophyceae) from widely differing sites, including two new species. J Phycol 31:996-1003. https://doi.org/10.1111/j.0022-3646.1995.00996

34. Frère L, Maignien L, Chalopin M, Huvet A, Rinnert E, Morrison H, Kerninon S, Cassone AL, Lambert C, Reveillaud J, Paul-Pont I (2018) Microplastic bacterial communities in the Bay of Brest: Influence of polymer type and size. Environ Pollut 242(Pt A):614-625.
https://doi.org/10.1016/j.envpol.2018.07.023

35. Garcés E, Masó M, Camp J (2002) Role of temporary cysts in the population dynamics of Alexandrium taylori (Dinophyceae). J Plankton Res 24:681-686.
https://doi.org/10.1093/plankt/24.7.681

36. Gélabert A, Pokrovsky OS, Viers J, Schott J, Boudou A, Feurtet-Mazel A (2006) Interaction between zinc and freshwater and marine diatom species: Surface complexation and Zn isotope fractionation. Geochim Cosmochim Acta 70(4):839-857. https://doi.org/10.1016/j.gca.2005.10.026

37. Gerwick WH, Proteau PJ, Nagle DG, Hamel E, Blokhin A, Slate DL (1994) Structure of Curacin A, a Novel Antimitotic, Antiproliferative and Brine Shrimp Toxic Natural Product from the Marine Cyanobacterium Lyngbya majuscula. J Org Chem 59(6):1243-1245.
https://doi.org/10.1021/jo00085a006

38. Gómez F (2012) A quantitative review of the lifestyle, habitat and trophic diversity of dinoflagellates (Dinoflagellata, Alveolata). Syst Biodivers 10(3):267-275.
http://dx.doi.org/10.1080/14772000.2012.721021

39. Grasshoff K, Ehrhardt M, Kremling K (1983) Methods of Seawater Analysis. 2nd Edition, Verlag Chemie Weinhein, New York, 419 p

40. Gugger M, Lenoir S, Berger C, Ledreux A, Druart JC, Humbert JF, Guette C, Bernard C (2005) First report in a river in France of the benthic cyanobacterium Phormidium favosum producing anatoxin-a associated with dog neurotoxicosis. Toxicon 45(7):919-928.
https://doi.org/10.1016/j.toxicon.2005.02.031

41. Harrison JP, Schratzberger M, Sapp M, Osborn AM (2014) Rapid bacterial colonization of low-density polyethylene microplastics in coastal sediment microcosms. BMC Microbiol 14:232.
https://doi.org/10.1186/s12866-014-0232-4

42. Hakanson L (1980) An ecological risk index for aquatic pollution control. A sedimentological approach. Water Res 14:975-1001
43. Hasle GR, Lange CB, Syvertsen EE (1996) A review of Pseudo-nitzschia, with special reference to the Skagerrak, North Atlantic, and adjacent waters. Helgol Meersunter 50:131-175

44. Heath MA, Wood SA (2010) Benthic Cyanobacteria and Anatoxina and Humanatoxin-a Concentrations in Five Southland Rivers. Cawthron Report No. 1841, New Zealand, 15 pp

45. Hoellein TJ, McCormick AR, Hittie J, London MG, Scott JW, Kelly JJ (2017) Longitudinal patterns of microplastic concentration and bacterial assemblages in surface and benthic habitats of an urban river. Freshw Sci 36(3):491-507. https://doi.org/10.1086/693012

46. Jung MR, Horgen FD, Orski SV, Rodriguez CV, Beers KL, Balazs GH, Jones TT, Work TM, Brignac KC, Royer SJ, Hyrenbach KD, Jensen BA, Lynch JM (2018) Validation of ATR FT-IR to identify polymers of plastic marine debris, including those ingested by marine organisms. Mar Pollut Bull 127:704-716. https://doi.org/10.1016/j.marpolbul.2017.12.061

47. Kesy K, Oberbeckmann S, Kreikemeyer B, Labrenz M (2019) Spatial environmental heterogeneity determines young biofilm assemblages on microplastics in baltic sea mesocosms. Front Microbiol 10:1-18. https://doi.org/10.3389/fmicb.2019.01665

48. Kettner MT, Rojas-Jimenez K, Oberbeckmann S, Labrenz M, Grossart HP (2017) Microplastics alter composition of fungal communities in aquatic ecosystems. Environ Microbiol 19(11):4447-4459. https://doi.org/10.1111/1462-2920.13891

49. Kettner MT, Oberbeckmann S, Labrenz M, Grossart HP (2019) The eukaryotic life on microplastics in brackish ecosystems. Front Microbiol 10:538. https://doi.org/10.3389/fmicb.2019.00538

50. Khiari N, Khalil N, Charef A, Atoui A (2022) Spatial variability of heavy metals in the coastal area of Monastir and origin of pollution. Arab J Geosci 15:297. DOI: 10.1007/s12517-022-09586-2

51. Kirstein IV, Wichels A, Krohne G, Gerdts G (2018) Mature biofilm communities on synthetic polymers in seawater - Specific or general? Mar Environ Res 142:147-154. https://doi.org/10.1016/j.marenvres.2018.09.028

52. Komárek J, Anagnostidis K (2005) Cyanoprokaryota. 2. Teil: Oscillatoriales. In: Büdel B, Gärtner G, Krienitz L, Schagerl M (eds) Süßwasserflora von Mitteleuropa, vol 19. Elsevier GmbH, München, pp 1-759. 2

53. Konno H (1993) Settling and coagulation of slender type diatoms. Water Sci Technol 27:231

54. Lee RE (2008) Phycology. Cambridge Univ. Press

55. Lelong A, Hégaret H, Soudant P, Bates SS (2012) Pseudo-nitzschia (Bacillariophyceae) species, domoic acid and amnesic shellfish poisoning: revisiting previous paradigms. Phycologia 51(2):168-216. https://doi.org/10.2216/11-37.1

56. Lenoir S, Ten-Hage L, Turquet J, Quod JP, Bernard C, Hennion MC (2004) First evidence of palytoxin analogues from an Ostreopsis mascarenensis (dinophyceae) benthic bloom in southwestern indian Ocean1. J Phycol 40:1042-1051. https://doi.org/10.1111/j.1529-8817.2004.04016.x

57. Li W, Zhang Y, Wu, Zhao Z, Xu W, Ma Y, Niu Z (2019) Colonization characteristics of bacterial communities on plastic debris influenced by environmental factors and polymer types in the haihe
estuary of Bohai Bay. China Environ Sci Technol 53:10763-10773.
https://doi.org/10.1021/acs.est.9b03659

58. Long ER, MacDonald DD, Smith SL, Calder FD (1995) Incidence of adverse biological effects within ranges of chemical concentrations in marine and estuarine sediments. Environ Manage 19:81-97.
https://doi.org/10.1007/BF02472006

59. Long ER, Morgan LG (1990) The potential for biological effects of sediment-sorbed contaminants tested in the national status and trends program. NOAA Tech. Memo. NOS OMA 52. US National Oceanic and Atmospheric Administration, Seattle, Washington, p 175

60. Lorenzen CJ (1966) A method for continuous measurement of "in vivo" chlorophyll concentration. Deep-Sea Res 13:223-227

61. Masó M, Fortuño JM, de Juan S, Demestre M (2016) Microfouling communities from pelagic and benthic marine plastic debris sampled across Mediterranean coastal waters. Sci Mar 80S1:117-127.
http://dx.doi.org/10.3989/scimar.04281.10A

62. Masó M, Garcés J, Pagès F, Camp J (2003) Drifting plastic debris as a potential vector for dispersing Harmful Algal Blooms (HAB) species. Sci Mar 67:107-111.
https://doi.org/10.3989/scimar.2003.67n1107

63. McCormick A, Hoellein TJ, Mason SA, Schluep J, Kelly JJ (2014) Microplastic is an abundant and distinct microbial habitat in an urban river. Environ Sci Technol 48(20):11863-11871.
https://doi.org/10.1021/es503610r

64. Michels J, Stippkugel A, Lenz M, Wirtz K, Engel A (2018) Rapid aggregation of biofilm-covered microplastics with marine biogenic particles. Proc Biol Sci 285(1885):20181203.
https://doi.org/10.1098/rspb.2018.1203

65. Mudroch A, Azcue JM, Mudroch P (1996) Manual of Physico-Chemical Analysis of Aquatic Sediments. CRC Press Inc., FL, U.S.A., p 356

66. Müller P, Suess E (1979) Productivity, sedimentation rate and sedimentary organic matter in the oceans_I. Organic carbon preservation. Deep-Sea Research, 26 A, 1347-1362.
https://doi.org/10.1016/0198-0149(79)90003-7

67. Muthukrishnan T, Al Khaburi M, Abed RMM (2019) Fouling Microbial Communities on Plastics Compared with Wood and Steel: Are They Substrate- or Location-Specific? Microb Ecol 78(2):361-374.
https://doi.org/10.3389/2Ffmicb.2021.673553

68. Nandiyanto ABD, Oktiani R, Ragadhita R (2019) How to Read and Interpret FTIR Spectroscope of Organic Material. Indonesian J Sci Technol. https://doi.org/10.17509/ijost.v4i1.15806

69. Nava V, Leoni B (2021) A critical review of interactions between microplastics, microalgae and aquatic ecosystem function. Water Res 188:116476. https://doi.org/10.1016/j.watres.2020.116476

70. Oberbeckmann S, Kreikemeyer B, Labrenz M (2018) Environmental factors support the formation of specific bacterial assemblages on microplastics. Front Microbiol 8:1-12.
https://doi.org/10.3389/fmicb.2017.02709
71. Oberbeckmann S, Osborn AM, Duhaime MB (2016) Microbes on a Bottle: Substrate, Season and Geography Influence Community Composition of Microbes Colonizing Marine Plastic Debris. PLoS One 11(8):e0159289. https://doi.org/10.1371/journal.pone.0159289

72. Oberbeckmann S, Loeder MG, Gerdts G, Osborn AM (2014) Spatial and seasonal variation in diversity and structure of microbial biofilms on marine plastics in Northern European waters. FEMS Microbiol Ecol. 2014 Nov;90(2):478-92. https://doi.org/10.1111/1574-6941.12409

73. Oliver RL (1994) Floating and sinking in gas-vacuolate cyanobacteria. J Phycol 30:161-173.
http://dx.doi.org/10.1111/j.0022-3646.1994.00161.x

74. Patil JS, Anil AC (2005) Quantification of diatoms in biofilms: Standardisation of methods. Biofouling 21(3-4):181-188. https://doi.org/10.1080/08927010500241726

75. Puschner B, Hoff B, Tor ER (2008) Diagnosis of anatoxin-a poisoning in dogs from North America. J Vet Diagn Invest 2008 Jan 20(1):89-92. https://doi.org/10.1177/104063870802000119

76. Ramsperger AFRM, Stellwag AC, Caspari A, Fery A, Lueders T, Kress H, Löder MGJ, Laforsch C (2020) Structural Diversity in Early-Stage Biofilm Formation on Microplastics Depends on Environmental Medium and Polymer Properties. Water 12(11):3216. https://doi.org/10.3390/w12113216

77. Reisser J, Shaw J, Hallegraeff G, Proietti M, Barnes DK, Thums M, Wilcox C, Hardesty BD, Pattiaratchi C (2014) Millimeter-sized marine plastics: a new pelagic habitat for microorganisms and invertebrates. PLoS ONE 9:e100289. https://doi.org/10.1371/journal.pone.0100289

78. Renner LD, Weibel DB (2011) Physicochemical regulation of biofilm formation. MRS Bull 36(5):347-355. https://doi.org/10.1557/mrs.2011.65

79. Ribeiro-Brasil DRG, Brasil LS, Veloso GKO, Matos TP, Lima ES, Dias-Silva K (2022) The impacts of plastics on aquatic insects. Sci Total Environ 813:152436.
https://doi.org/10.1016/j.scitotenv.2021.152436

80. Rochman C, Hoh E, Kurobe T, Teh SJ (2013) Ingested plastic transfers hazardous chemicals to fish and induces hepatic stress. Sci Rep 3:3263. https://doi.org/10.1038/srep03263

81. Rutkowski P, Diatta J, Konatowska M, Andrzejewska A, Tyburski Ł, Przybylski P (2020) Geochemical Referencing of Natural Forest Contamination in Poland. Forests 11:157,1-18.
https://doi.org/10.3390/f11020157

82. Sassi R, Souissi F, Soussi N, Boukaaba M, Belayouni H (1998) Géochimie de la matière organique pour l'analyse de la dégradation du littoral de Monastir-Ksibet el Mediouni (Tunisie orientale). C R Acad Sci Paris 303-308. DOI 10.1016/s1251-8050(98)80047-5

83. Sherr EB, Sherr BF (2002) Significance of predation by protists in aquatic microbial food webs. Antonie Van Leeuwenhoek 81(1-4):293-308. https://doi.org/10.1023/a:1020591307260

84. Singh N, Turner A (2009) Trace metals in antifouling paint particles and their heterogeneous contamination of coastal sediments. Mar Pollut Bull 58:559-564.
https://doi.org/10.1016/j.marpolbul.2008.11.014

85. Souissi R, Turki I, Souissi F (2014) Effect of submarine morphology on environment quality: Case of Monastir bay (Easter Tunisia). Carpathian Journal of Earth and Environmental Sciences, North
University Center of Baia Mare, Romania, 2014, 9 (3), pp.231-239. https://hal.archives-ouvertes.fr/hal-01550712

86. Stefatos A, Charalampakis M, Papatheodorou G, Ferentinos G (1999) Marine Debris on the Seafloor of the Mediterranean Sea: Examples from Two Enclosed Gulfs in Western Greece. Mar Pollut Bull 36:389-393. https://doi.org/10.1016/S0025-326X(98)00141-6

87. Tessier A, Campbell PGC, Bisson M (1979) Sequential extraction procedure for the speciation of particulate trace metals. Anal Chem 51:844-851. http://dx.doi.org/10.1021/ac50043a017

88. Turekian KK, Wedepohl KH (1961) Distribution of the Elements in Some Major Units of the Earth's Crust. Geol Soc Am Bull 72:175-192. http://dx.doi.org/10.1130/0016-7606(1961)72[175:DOTEIS]2.0.CO;2

89. Uthermohl H (1958) Zur Vervollkommnung der quantitativen Phytoplanktonmethodik. Mitteilungen Inst. Ass. Theor. Appl. Limnologie, Stuttgard, 9, I-38

90. Weig AR, Löder MGJ, Ramsperger AFRM, Laforsch C (2021) In situ Prokaryotic and Eukaryotic Communities on Microplastic Particles in a Small Headwater Stream in Germany. Front Microbiol 12:660024. https://doi.org/10.3389/fmicb.2021.660024

91. Williamson RT, Boulanger A, Vulpanovici A, Roberts MA, Gerwick WH (2002) Structure and absolute stereochemistry of phormidolide, a new toxic metabolite from the marine cyanobacterium Phormidium sp. J Org Chem 67(23):7927-7936. https://doi.org/10.1021/jo020240s

92. Wood SA, Smith FMJ, Heath MW, Palfroy T, Gaw S, Young RG, Ryan KG (2012) Within-mat variability in anatoxin-a and homoanatoxin-a production among benthic Phormidium (Cyanobacteria) Strains. Toxins 4:900e912. https://doi.org/10.3390/2Ftoxins4100900

93. Yan N, Liu W, Wie H, Gao L, Han Y, Wang M, Li H (2016) Distribution and assessment of heavy metals in the surface sediment of Yellow River, China. J Environ Sci 39:45-51. https://doi.org/10.1016/j.jes.2015.10.017

94. Yokota K, Waterfield H, Hastings C, Davidson E, Kwietniewski E, Wells B (2017) Finding the missing piece of the aquatic plastic pollution puzzle: Interaction between primary producers and microplastics. Limnol Oceanogr Lett 2(4):91-104. https://doi.org/10.1002/loL2.10040

95. Zaaboub N, Martins MVA, Dhib A, Bejaoui B, Galgani F, El Bour M, Aleya L (2015) Accumulation of trace metals in sediments in a Mediterranean Lagoon: usefulness of metal sediment fractionation and elutriate toxicity assessment. Environ Pollut 207:226-237. https://doi.org/10.1016/j.envpol.2015.09.033

96. Zettler ER, Mincer TJ, Amaral-Zettler LA (2013) Life in the "Plastisphere": microbial communities on plastic marine debris. Environ Sci Technol 47:7137-7146. https://doi.org/10.1021/es401288x

97. Zettler ER, Mincer T, Proskurowski G, Amaral-Zettler LA (2011) The "Plastisphere": A new and expanding habitat for marine protists. J Phycol 47:pS45-pS45

98. Zhang C, Yu ZG, Zeng GM, Jiang M, Yang Z, Fang Cui, Zhu MY, Shen LQ, Hu L (2014) Effects of sediment geochemical properties on heavy metal bioavailability. Environment International 53 (2014) 270-281. https://doi.org/10.1016/j.envint.2014.08.010
## Tables

Table 1 to 5 are available in the Supplementary Files section.

## Figures

<figure><img src="image_2.png" /><figcaption>**Figure 1**</figcaption></figure>

Monastir Bay study area and sampling stations (S1: Kuriat Island, S2: Agba City, S3: Khniss, S4: Ksibat El Mediouni, S5: Lamta wastewater treatment station, S6: Aquaculture cage cleaning station, S7: Bekalta, S8: Ras Dimas)

<caption>**Figure 2**</caption>

Scanning electron microscopy images and elemental analysis of the LDPE surface before and after plastisphere extraction (scraping).

<caption>**Figure 3**</caption>
ATR FTIR spectra of LDPE plastic bag labelled with resin code # 4 (control) and LDPE plastic bags debris collected from the eight sampling stations

Figure 4

Scanning electron micrographs showing the diversity and structural complexity of the plastisphere that develops on LDPE collected from different stations in Monastir Bay.

Figure 5

Bar blots of relative abundance (%) of: different microalgae groups (to total microalgae) (a), cyanobacteria species (relative to total cyanobacteria) (b), dinoflagellates species (relative to total dinoflagellates) (c) and diatoms species (relative to total diatoms) (d) in plastisphere (P) and sea water (SW) at the eight sampling stations.
<figure><img src="image_3.png" /><figcaption>Figure 6</figcaption></figure>

Mean (± SD) relative abundance (%) of total potential harmful microalgae (to total microalgae) in plastisphere and sea water at the eight sampling stations.

<caption>Figure 7</caption>

Bar plots of relative abundance (%) of: different groups of potential harmful microalgae (to total potential harmful microalgae) (a), potential harmful cyanobacteria species (relative to total potential harmful cyanobacteria) (b), potential harmful dinoflagellates species (relative to total potential harmful dinoflagellates) (c) and potential harmful diatoms species (relative to total potential harmful diatoms) (d) in plastisphere (P) and sea water (SW) at the eight sampling stations.
Figure 8

Trace element concentrations in surface sediments and plastic debris plastispheres of Monastir Bay: (a) Cd, Co, Cu, and Cr concentrations; (b) Ni, Pb, and Zn concentrations.

**Supplementary Files**

This is a list of supplementary files associated with this preprint. Click to download.

* Table1.docx

* Table2.docx

* Table3.docx

* Table4.docx

* Table5.docx