Salt marsh zonation and substrate type modulation for plastisphere: an experimental assessment in the Lagoon Patos estuary in extreme south of Brazil
Lara Mesquita Pinheiro, Larissa Tomasin Andreola, Carlos Rafael Borges Mendes, Mikael Luiz Pereira Morales, Vanessa Ochi Agostini, Grasiela Lopes Leães Pinho
TL;DR
This study explores how salt marsh zones and plastic properties influence the development of microbial communities on plastics in a Brazilian estuary.
Contribution
The study reveals that environmental factors like flooding regimes have a stronger influence on plastisphere communities than plastic properties.
Findings
Flooded and intermediate zones showed higher microbial colonization than dry zones.
Smaller plastics favored microbial growth, while larger ones supported more macrofouling.
EVA polymer substrates had higher biofilm accumulation compared to other plastics.
Abstract
The accumulation of plastics in aquatic environments is a growing global concern, as biofouling on plastic debris leads to the formation of the plastisphere, an ecological niche for diverse microbial and macrofouling organisms. Although plastic characteristics such as size, color, and polymer type may influence plastisphere development, there is no consensus regarding their relative importance, and studies in estuarine environments remain scarce. Here, we investigated plastisphere formation across three salt marsh zones (dry, intermediate, and flooded) in the Patos Lagoon estuary (southern Brazil), considering variations in plastic size (6 × 2, 30 × 10, and 60 × 20 mm), color (white, black, and red), and polymer type (Polypropylene—PP, Polystyrene—PS, and Ethylene Vinyl Acetate—EVA). Three independent 21-day field experiments were conducted, and plastisphere development was assessed…
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Figure 9- —Universidade Federal Do Rio Grande
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Taxonomy
TopicsCoastal wetland ecosystem dynamics · Coastal and Marine Dynamics · Geological formations and processes
Introduction
Currently, one of the growing global environmental concerns is the excessive accumulation of plastic waste in aquatic environments, thus being considered a pollutant of emerging concerns (Amaral-Zettler et al. 2021; Gonçalves et al. 2025). These residues can fragment and generate smaller particles that can be classified according to their size into: > 1 m (megaplastics), 25–1000 mm (macroplastic), 5–24 mm (mesoplastics), 1 μm–5 mm (microplastics) and < 1 μm (nanoplastics) (Frias and Nash 2019; Hanvey et al. 2017). Once in the water, plastics provide a solid surface for organisms (e.g., bacteria, algae, fungi, and macroorganisms) to colonize, forming the plastisphere (Zettler et al. 2013).
The plastisphere is a specific ecological niche for the microbial community, serving as an excellent substrate that promotes metabolic adaptations and survival (Zhai et al. 2023). The composition of this community within the plastisphere is reported to be more diverse than the community found in its surrounding environment (e.g., water and sediment) (Vannini et al. 2021). The plastisphere can facilitate microbial transport within and between ecosystems (Silva et al. 2023a, b), acting as a disperser of pathogenic microorganisms (Ormsby et al. 2023). It can also adsorb heavy metals (e.g., zinc and copper) (Richard et al. 2019) and antibiotics (Silva et al. 2023a, b), consequently promoting microbial evolution by enhancing the horizontal transfer of antibiotic (Yang et al. 2019) and heavy metals resistance genes (Wu et al. 2024). Consequently, plastics cause several effects on the biota and the ecosystem (Battisti et al. 2023; Gonçalves et al. 2025).
Due to the roughness and hydrophobic properties of different types of plastic polymers, studies indicate that different plastic types of influence plastisphere formation, facilitating or inhibiting some taxa in the formation of biofilm and the colonization of microbial communities (Miao et al. 2019; Sooriyakumar et al. 2022). For example, in marine environments, bacteria tend to colonize polystyrene (PS) more easily, while algae prefer to colonize polyvinyl chloride (PVC) and polyethylene terephthalate (PET) surfaces (Miao et al. 2021). In addition, bacterial abundance has been reported to be higher on PS surfaces compared to polypropylene (PP) and polyethylene (PE) (Sooriyakumar et al. 2022). However, some studies have shown that the plastispheric community is not influenced by the different types of polymers (Sérvulo et al. 2023a, b; Wallbank et al. 2022). For salt marsh environments, there is a difference between the plastisphere between PET, PE and PP, with PP and PE colonized more by nitrifying bacteria and PET by cyanobacteria (Hou et al. 2025). These authors also reinforce that the difference in the formation of the plastisphere is mainly due to the difference in the characteristics of the plastics, due to their roughness and structure (Hou et al. 2025), but that their colonization can still vary in relation to the geographic region studied. Thus, it is important to investigate the formation of the plastisphere in different characteristics of the polymers (e.g., ethylene–vinyl acetate–EVA, PP and PE) and in geographic regions not yet studied (Rosato et al. 2022), and comparison approaches between different polymers are recommended (Pinto et al. 2019; Sooriyakumar et al. 2022).
In this context, the color and size of polymers can also influence plastisphere formation. Dark-colored substrates tend to support higher plastisphere densities, whereas white substrates typically exhibit lower densities (Dobretsov et al. 2013). A similar pattern was observed by Satheesh and Wesley (2010), who reported that red and blue substrates harbored higher densities of macroorganisms compared to white ones. Conversely, Wen et al. (2020) reported that polymer color does not affect plastisphere density; however, some species exhibit preferences for specific colors, with blue being more attractive to certain microbial species and yellow favoring pathogenic and biodegrading species. The same authors also found that polymer size (ranging from 0.22 to 3 mm) does not influence plastisphere formation. Similarly, Frère et al. (2018) reported no significant influence of polymer sizes between 0.3 and 5 mm on plastisphere development. However, Debroas et al. (2017) observed differences in plastisphere composition when comparing plastic sizes ranging from 0.3 mm to 20 cm. Therefore, no consensus has been reached regarding whether polymer color and size are deterministic factors for organism colonization, representing a research gap that warrants further investigation. Furthermore, we emphasize that most of these studies were conducted in marine environments (Dobretsov et al. 2013; Debroas et al. 2017; Satheesh and Wesley 2010; Wen et al. 2020), rather than in estuarine environments (Frère et al. 2018), highlighting an additional research gap in estuarine systems.
Several studies indicate that environmental conditions in each region have a greater influence on plastisphere formation than the intrinsic characteristics of polymers (Marsay et al. 2023; Nguyen et al. 2023; Philippe et al. 2023; Sooriyakumar et al. 2022; Tang and Li 2025). To date, the marine environment is by far the most extensively studied in relation to plastic pollution and plastisphere development, while research in estuarine systems remains limited (Delacuvellerie et al. 2022). Salt marshes, which are located within estuarine environments, experience fluctuations in flooding, salinity, and temperature that directly influence the composition and diversity of local flora and fauna. These ecosystems are important transitional zones with high ecological connectivity (Laruelle et al. 2013; Odebrecht et al. 2010). Notably, salt marshes often act as sinks for plastic debris, resulting in high levels of plastic contamination in these environments (Pinheiro et al. 2021).
In the south extreme region of South America, substantial gaps remain regarding how the plastisphere forms and varies across estuarine salt marshes, particularly in systems that function simultaneously as sinks and transformers of plastic debris. Although pioneering studies in the Patos Lagoon estuary have described the presence and seasonal dynamics of the plastisphere (Pinheiro et al. 2021; Sérvulo et al. 2023a, b), no research has yet been tested whether its composition is shaped by the physical characteristics of plastics, such as polymer type, color, and size, despite the contradictory evidence reported in the global literature. Moreover, it is unclear whether environmental gradients intrinsic to salt marshes, especially flooding regimes, exert a stronger influence on plastisphere assembly than the intrinsic properties of plastics themselves.
These knowledge gaps led us to propose the following hypotheses: (H1) plastisphere composition differs among polymer types due to variations in surface structure and roughness, as previously reported for different synthetic materials (Miao et al. 2019; Sooriyakumar et al. 2022; Hou et al. 2025); (H2) flooding zones structure microbial colonization by imposing distinct environmental pressures, such as differences in immersion time, salinity, and exposure, which are known to shape estuarine communities (Laruelle et al. 2013); (H3) dark-colored polymers support higher microbial densities than lighter-colored ones, following patterns observed for biofouling on colored substrates (Dobretsov et al. 2013; Satheesh and Wesley 2010); and (H4) larger substrates support different communities than smaller substrates due to greater surface heterogeneity and settlement area (Debroas et al. 2017; Fazey and Ryan 2016). Addressing these hypotheses represents an important and still underexplored research frontier in subtropical South America estuaries. Therefore, this study provides the first multifactorial and spatially explicit assessment of plastisphere formation in a salt marsh of the Patos Lagoon estuary, integrating three flooding zones (flooded, intermediate, and dry) and three key plastic attributes: size (6 × 2 mm, 30 × 10 mm, and 60 × 20 mm), color (white, red, and black), and polymer type (PP, PS, and EVA). This approach represents a novel contribution by simultaneously testing multiple plastic characteristics in a highly dynamic estuarine environment, offering new insights into the mechanisms that drive organism colonization on plastics in salt marsh ecosystems.
Material and methods
Study area
The Patos Lagoon is in the coastal plain of South Brazil, and it is the largest coastal lagoon in South America with 10,360 km^2^ of area (Kjerfve 1986). The entire lagoon watershed encompasses an area of 201,626 km^2^, and it is connected to the Atlantic Ocean by a narrow channel (700 m wide) in its estuarine region located at its southern end (António et al. 2020). The climate and hydrodynamics in this region are mainly defined by the wind systems, precipitation, and fluvial discharge, while microtidal system plays a secondary role (Moller et al. 2009; Pereira et al. 2011). Northern winds are predominant in the summer, favouring the flood regime, while southern winds prevail in the winter and favour the ebb regime (Odebrecht et al. 2010). These cause the seasonal variations in salinity and temperature that directly affect the flooding rates in the lagoon’s margins (Barros et al. 2014). The estuarine region hosts several different environments such as mud flats, beaches, salt marshes and artificial substrates that serve as habitat to many species of ecological and economical importance (Odebrecht et al. 2010). In addition, several anthropogenic activities are developed in this region such as fishing, industries, agriculture, feedstock, and port activities, which in turn affect the surrounding environmental quality (Odebrecht et al. 2010).
A total of 24 salt marsh environments occur in the Patos Lagoon estuary, differing in size, vegetation, flooding rates, and natural and anthropogenic impacts (Costa et al. 1997). Among these, the Molhe Oeste salt marsh is in the city of Rio Grande, southern Brazil (Fig. 1A), covering 0.16 km^2^ at the mouth of the Patos Lagoon estuary (32° 09′ 09.3″ S; 52° 06′ 03.1″ W). Due to its position at the lagoon–ocean interface, this salt marsh potentially receives plastic inputs from both systems. The Molhe Oeste salt marsh is subdivided into four zones based on vegetation and flooding regime: High Marsh–HM (flood index 3.1%), Middle Marsh–MM (20.1%), Low Marsh–LM (64%), and Mud Flat–MF (100%) (Perillo 1999). The flood index corresponds to the percentage of time each zone remains inundated during the tidal cycle and was derived from long-term tidal and elevation data (Perillo 1999). Vegetation in the HM–MM zone includes Bolboschoenus maritimus, Juncus kraussii, Hydrocotyle bonariensis, and Myrsine parvifolia, whereas the MM–LM zone is dominated by Spartina densiflora and S. alterniflora. The LM–MF zone is partially unvegetated. For all experiments, three zones were considered: flooded (MF–LM; flood index 64–100%), intermediate (LM–MM; 20.1–63%), and dry (MM–HM; 3.1–20%), representing the flooding gradient of the Molhe Oeste salt marsh (Fig. 1B). This site was selected due to previously reported plastic contamination (Pinheiro et al. 2021).Fig. 1. Location of the experimental site, salt marsh of Molhe Oeste, Rio Grande, Rio Grande do Sul, Brazil (A) and experimental frames—floating metal structure—in each zone of the salt marsh (B)
Experimental setup
Three separate field experiments were performed to investigate the influence of different plastics characteristics and salt marsh zonation in biofouling. For each experiment, virgin plastic products were purchased and used to fabricate rectangular substrates manually. Three plastics characteristics were tested: size, color, and polymer type. For the size experiment, white polystyrene cups were cut into 6 × 2 mm, 30 × 10 mm, and 60 × 20 mm substrates. For the color experiment, white, black, and red EVA sheets were cut into 30 × 10 mm substrates. For the polymer type experiment, white PP and PS cups and EVA sheets were cut into 30 × 10 mm substrates. The experiments were conducted in three salt marsh zones (flooded, intermediate, and dry), as described above. Plastic items for each experiment were placed in buoyant metallic structures covered by a metallic mesh (500 µm pore size) that allowed initial colonizers to access the substrates (Fig. 1B). Each metallic structure was placed in one zone of the salt marsh, in a total of three experimental units. The substrates were incubated in the salt marsh for 21 days to capture mature biofilm and initial invertebrate settling. Experiments were performed in June–July 2021 (size experiment), October–November 2021 (colour experiment), and February–March 2022 (polymer type experiment). After incubation, all substrates were transported to the laboratory and organized into three replicates, corresponding to three plastic samples for each experiment (color, size, and polymer type) in each zone (intermediate, flooded, and dry), which were used for the analyses described below. That is, each analysis below contained three plastic polymers for each zone and treatment of the experiment.
Plastisphere weight
To determine plastisphere weight, substrates were gently washed three times with sterile saline solution 0.9% to remove non-adhered material/organisms. Then, their weight was measured with an analytical scale (0.001 g precision), then dried in oven at 40 °C until constant weight. To extract the plastisphere from plastics surfaces, substrates were scraped with a glass coverslip and transferred to cryovials containing sterile saline solution 0.9%. Tubes were agitated in vortex for 30 s at 1500 rpm, then sonicated in ultrasound at 40 kHz for 2 min, and vortexed again for 30 s. The solution was then discarded, and plastic substrates were washed three times with sterile saline solution 0.9% to remove loose material. They were dried again in oven at 40 °C until constant weight. The plastisphere weight was then calculated according to the following equation:
\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$PW = \frac{{\left( {W_{i} - W_{f} } \right)}}{{A_{s} }}$$\end{document}where PW is the plastisphere weight (mg.mm^–2^), W_i_ is the initial substrate weight (mg), W_f_ is the final substrate weight (mg), and A_s_ is the substrate area (mm^–2^).
Bacterial density
Bacterial density on the substrates surface was estimated in Epifluorescence microscopy using a methodology adapted from Agostini et al. (2021). The plastisphere was extracted from the plastics surfaces using the same methodology described in Sect. "Plastisphere weight". The resulting solution containing the suspended plastisphere was fixed with formaldehyde to a final concentration of 4% and stored at 8ºC until further analysis. Samples from the dry and intermediate salt marsh zones were diluted to 1:1 and from the flooded zone to 1:10 with distilled water. An aliquot 1 mL of each diluted sample was filtered using a vacuum pump through a nucleopore membrane filter (Whatman Ø 25 mm, 0.2 µm pore size), which were previously darkened with Irgalan Black. Filters were then stained with Acridine Orange 0.1%, washed with distilled water, and filtered again. Filters were then dried at room temperature and placed in microscopy glass slides and covered with a glass coverslip. Slides were analysed under an epifluorescence microscope (Zeiss Axioplan, 1000 × magnification), where five visual fields were randomly selected and photographed. Bacterial cells were counted on a ZEN lite software (Carl Zeiss Microscopy, v2.1), and bacterial density was estimated according to Eq. 2:
\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$BD = \frac{{\left( {C_{{avg}} * A_{f} * V_{s} } \right)}}{{A_{v} * V_{f} * A_{s} }}$$\end{document}where BD is the bacterial density (cells.mm^–2^), C_avg_ is the cells average count per slide, A_f_ is the filter area (mm^–2^), V_s_ is the total sample volume (mL), A_v_ is the microscope visual field area (mm^–2^), V_f_ is the filtered sample volume (mL), and A_s_ is the substrate area (mm^–2^).
Based characterization of pigment composition
The composition of the plastisphere in terms of biofouling microalgae was assessed by quantifying photosynthetic pigments using high-performance liquid chromatography (HPLC) following a methodology developed by Zapata et al. (2000) and adapted by Mendes et al. (2007). The pigments were selected according to the most abundant microalgae groups: chlorophyll a, which is found in all algal groups, was used to estimate total microalgae biomass; lutein and chlorophyll b served as biomarkers for chlorophytes, while fucoxanthin, chlorophylls c1 and c2 were used to identify diatoms. Substrates were initially washed three times with sterile saline solution 0.9% and sonicated for 5 min, then pigments were extracted from individual substrates using 4 mL of methanol 95% buffered with 2% ammonia acetate containing 0.05 mg L^–1^ of internal standard (trans-β-apo-8’-carotenal) for 60 min at −20° C in the dark. Extracts were centrifuged at 1100 rpm for 5 min at 4° C and subsequently filtered in Fluoropore PTFE membrane filters (Millipore Ø 13 mm, 0.2 µm pore size).
Pigment extracts (1 mL) were mixed with 400 µL of Milli-Q water to adjust the extract polarity according to the method used (Zapata et al. 2000), and then were injected on a HPLC (Shimadzu) system constituted by a solvent distribution module (LC-20AD) coupled with a degassing unit (DGU-20A5), a system controller (CBM-20A), a refrigerated auto injector (SIL-20AC), a column oven (CTO-20AC), a photodiode detector (SPD-M20A), and a fluorescence detector (RF-10AXL). A monomeric OS C8 column (Waters® SunFire, 15 cm length, 4.6 mm diameter, 3.5 µm particle size) was used to separate pigments in the chromatography step, using a flow rate of 1 mL.min^–1^, an injection volume of 100 µL, and a run time of 40 min. Peak identification and quantification were performed using commercial reference standards from DHI (Institute for Water and Environment, Denmark). Pigment concentrations were calculated from signals obtained from the photodiode (lutein and fucoxanthin) and/or fluorescence (chlorophyll) detectors. Peaks were integrated using the LC-Solution software and were manually checked and corrected when necessary. Pigment concentrations were normalized using the internal standard to correct for volume variations.
Plastisphere community composition
Macrofouling
Substrates from field experiments were analysed for the presence (attachment) of small invertebrates (macroorganisms) on their surface. Individual substrates were fixed in formaldehyde 4% solution for at least 24 h, when they were analysed visually under a stereoscopic microscope (Leica Wild M3Z and Opticam OPZTS). Both plastic substrates and containing formaldehyde solution were scanned for the presence of macroorganisms. Organisms were identified to the lowest taxonomical level based on its morphology and following literature by Matthews-Cascon et al. (2011), Rios (2009), Ruppert et al. (2005).
Scanning electron microscopy
To visualize the incrusting community on the plastics surface, substrates were analysed with scanning electron microscopy (SEM) following a methodology adapted from Agostini et al. (2021). Substrates were gently washed three times with sterile saline solution 0.9% to remove non-adhered material/organisms. Then, they were fixed in glutaraldehyde 1% for a minimum of 24 h before dried at 40 °C for 24 h. Subsamples of substrates measuring 30 × 10 mm or 60 × 20 mm were taken as larger pieces would not fit the SEM. Samples were then attached to aluminium stubs with carbon tapes and coated with gold powder. Prepared samples were then analysed under a scanning electron microscope (SEM JEOL, JSM-6610LV), and for each substrate three visual fields were randomly selected and photographed at 5000 × magnification. Morphological structures on the substrate’s surfaces were compared with images and descriptions from the literature and individuals were grouped in taxonomical groups: bacteria, fungi, and microalgae (Masó et al. 2016; Ramsperger et al. 2020).
Metabarcoding of procaryotes and fungi
To investigate the composition of prokaryotes and fungi in the Plastisphere, the metabarcoding technique performed by the company ‘Agrega Pesquisa e Desenvolvimento em Biotecnologia LTDA’ by sequencing the 16S and ITS genes was used. For this analysis, composite samples (pool) were prepared with the substrates of the experiments of different characteristics (sizes, colors and polymers) to obtain three final composite samples corresponding to the three areas of the marsh: dry, intermediate and flooded areas. The material was scraped from each substrate with a spatula and packed in a stabilizing solution (Caotropic Buffer Aggregate) in tubes provided by the company. The DNA was extracted using the commercial extraction kit, according to the manufacturer’s recommendations. DNA integrity, concentration and purity were evaluated by 1% agarose gel electrophoresis, QubitTM (Thermo Fisher Scientific) and NanoDrop 2000 (Thermo Scientific). Amplification of the 16S rRNA v4 region was performed using the primers 515 F 5′-GTG CCA GCM GCC GCG GTAA-3′ and 806R 5′-GGA CTA CHV GGG TWT CTA AT-3′ (Caporaso et al. 2011; Sundberg et al. 2013). For ITS markers, the primers used were ITS1 (5′-CTT GGT CAT TTA GAG GAA GTA A-3′) and ITS2 (5′-GCT GCG TTC TTC ATC GAT GC-3′) (Bellemain et al. 2010; White et al. 1990).
The libraries were evaluated using Qubit™ and sequenced on MiSeq (Illumina Inc., USA), using 250 bp paired-end reads and the MiSeq Reagent Kit™ v2. Chimeric sequence removal and clustering of operational taxonomic units (OTUs) were performed using the UPARSE software (Edgar 2013). Taxonomic identifications were carried out using the UNITE v2020 database (Firth et al. 2009), as well as the SILVA reference database v138.2 (Yilmaz et al. 2014). Rarefaction (normalization) of the metagenomic data (OTUs) was performed based on the sample with the lowest number of sequences. Subsequently, based on the relative abundance of sequences (OTUs) in each sample provided by the company, taxa frequency and richness were calculated for each salt marsh zone (flooded, intermediate, or dry). We emphasize that all bioinformatic processing steps (quality filtering, paired-end merging, chimera removal, OTUs clustering, and taxonomic assignment) followed the standard UPARSE pipeline and were performed by the sequencing company.
Data analysis
All data was tested for normality and homoscedasticity using Shapiro–Wilk and Levene’s tests, respectively. Statistical tests were performed for data within the same experiments only, i.e., data was not compared between the three experiments. Additionally, we emphasize that each analysis included three replicates, as detailed in Sect. "Experimental setup", except for the metabarcoding analysis, which was limited by resource constraints and is explained in Sect. "Metabarcoding of procaryotes and fungi". To check for significant differences between plastisphere weight (mg mm^−2^) and bacterial density (cells.mm^−2^) between each salt marsh zone (flooded, intermediate, dry) and plastics characteristics (size, colour, polymer type), two-way ANOVA tests were performed when data followed normality followed by a Tukey’s post hoc test to locate the differences. To check for these differences in microalgae pigments concentration (µg mm^−2^) in the flooded zone (as they could only be detected in this zone) between plastics characteristics, ANOVA tests were performed depending on data normality, followed by Tukey’s pairwise test. For the small invertebrates (macroorganisms) data, a PERMANOVA test based on a Bray–Curtis similarity matrix was used to check for differences in community composition between salt marsh zone and plastics characteristics. A SIMPER analysis was also used to determine the contribution of each taxon to macroorganisms composition in salt marsh zone and plastic characteristics.
NMDS ordination based on Bray–Curtis dissimilarity was applied as an exploratory tool to summarize overall community structure across salt marsh zones, specifically to evaluate the proximity or separation among zones. Core microbiome analysis was conducted to identify bacterial and fungal classes consistently present in all salt marsh zones, based on relative frequency data considering taxa with > 0% occurrence across the three flooding zones. Subsequently, a heatmap based on relative abundance was used to visualize shared and zone-specific taxa. To evaluate the diversity of the plastisphere, the richness of the taxa (n = the families), Shannon Index (H), Simpson Dominance (1-D) Index and Pielou Equitability Index (J) were calculated. All statistical analysis considered a significance limit of 5% and were performed in the PAST software (version 4.10).
Results
Plastisphere weight
In all three experiments (size, color and type of polymer), the weight of the plastisphere was higher in the flooded and intermediate zones compared to the dry zone (Fig. 2). For the size experiment, the highest weight (p < 0.05) was observed in the intermediate zone for the smallest polymer size (6 × 2 mm) and in the flooded zone for the largest polymer size (60 × 20 mm) (Fig. 2A). For the color experiment, in the intermediate zone, the white and black colors presented higher plastisphere weights (p < 0.05), while in the flooded zone it was only white (p < 0.05), with a higher weight when compared to the other zones (p < 0.05) (Fig. 2B). Finally, in the experiment with different polymers, the dry zone presented lower plastisphere weight values (p < 0.05), while the flooded zone presented higher plastisphere weight (p < 0.05) (Fig. 2C). In addition, the EVA polymer showed a higher weight of the plastisphere in each zone (p < 0.05) when compared to the other types of polymers (Fig. 2C).Fig. 2. Mean ± SD of plastisphere weight (mg mm^−2^) for different salt marsh zones and the different size (A), color (B) and polymer (C) experiments. Different letters—statistical differences between treatments and zones (p < 0.05)
Bacterial density
In the dry zone, the bacterial density was higher (p < 0.05) in the size experiment, while for the other experiments (color and type of polymer) the bacterial density was lower (p < 0.05) when compared to the flooded and intermediate zones (Fig. 3). For the size experiment, bacterial density decreased (p < 0.05) with increasing size for the three zones (Fig. 3A). For the color experiment, the bacterial density was lower (p < 0.05) for the dry zone, followed by the intermediate and flooded zones, respectively (p < 0.05), however, the bacterial density did not differ between the colors (p > 0.05) (Fig. 3B). For the experiment with the different polymers, the bacterial density did not differ between the three types of polymers (p > 0.05), however, the bacterial density was lower in the dry and intermediate zones, not differing from each other (p > 0.05) (Fig. 3C). The flooded zone showed higher bacterial density when compared to the other zones (p < 0.05) (Fig. 3C).Fig. 3. Mean ± SD of bacterial density (cells mm^−2^) for different salt marsh zones and the different size (A), color (B) and polymer (C) experiments. Different letters—statistical differences between treatments and zones (p < 0.05)
Based characterization of pigment composition
In the dry zone, chlorophyll a concentrations did not differ significantly among the different sizes, colors, or polymer types (p > 0.05). However, overall chlorophyll a concentration in the dry zone was significantly lower compared to the intermediate and flooded zones (p < 0.05) (Fig. 4). In the size experiment, the smallest size showed higher chlorophyll a concentration in the intermediate and flooded zones (Fig. 4A). In the color and polymer experiments, a similar pattern was observed: the dry and intermediate zones did not differ significantly (p > 0.05), while the flooded zone showed higher values of chlorophyll a (p < 0.05). In all cases, no significant differences were found among treatments within each zone (Fig. 4B, C).Fig. 4. Mean ± SD of chlorophyll a concentration (µg mm^−2^) for different salt marsh zones and the different size (A), color (B) and polymer (C) experiments. Different letters – statistical differences between treatments and zones (p < 0.05)
The concentrations of lutein and chlorophyll b, both biomarkers of chlorophytes (Fig. S1 and S2), followed the same response pattern as chlorophyll a. In contrast, chlorophyll c1 showed a different pattern: in the size (Fig. S3A) and polymer (Fig. S3C) experiments, the flooded zone exhibited higher concentrations of chlorophyll c1, particularly in the smaller size (p < 0.05) and in EVA polymer (p < 0.05). In the color experiment (Fig. S3B), chlorophyll c1 varied only between zones (p < 0.05), with no significant differences among colors (p > 0.05). For chlorophyll c2 and fucoxanthin, the highest concentrations (p < 0.05) were observed in the smallest size within the intermediate and flooded zones (Fig. S4A-S5A), in black and red colors in the flooded zone (Fig. S4B-S5B), and in the PS and PP polymers in the flooded zone (Fig. S4C-S5C).
Macrofouling community
In all experiments (size, color and polymer) the dry zone showed no macroorganisms (Fig. 5), except for the 6 × 2 size treatment, only two Bivalvia individuals were found (Fig. 5A). For the size experiment, the flooded area showed greater abundance, which was more evident in the size of 60 × 20 mm (Fig. 5A). For the color experiment, the intermediate zone was more abundant in terms of the presence of macroorganisms, and it was more representative in the red color (Fig. 5B). For the polymer experiment, the abundance of macroorganisms was higher in the flooded area, with the EVA polymer having the highest and PS the lowest abundance (Fig. 5C). However, in the intermediate zone, PP showed a higher abundance of macroorganisms (Fig. 5C). In all experiments, Bivalvia individuals contributed the most to the abundance of organisms, followed by Cirripedia for the size experiment, Foraminifera for the color experiment, and Gastropoda for the polymer type experiment (Fig. 5). Raw abundance data is available in Table S1.Fig. 5. Total abundance (number of individuals) of macrofouling community taxa for the different salt marsh zones and the different sizes (A), colors (B) and polymer (C)
Microbial community composition
Scanning electron microscopy (SEM) revealed a variety of microorganisms, including isolated and colony-like bacteria in the form of bacilli and cocci, yeasts and filamentous fungi, protozoa, and microalgae such as diatoms (pennate and centric) and Chlorophyceae (Fig. 6). We observed that with the increase in the level of flooding (dry zone – flooded) there was an increase in the colonization of organisms (Fig. 6). However, due to the difficulty in identifying the organisms and their visualization, we were unable to see possible differences between the treatments within each experiment (size, color and polymer).Fig. 6. Scanning electron microscopy images of the biofilm developed on the surface of the plastics exposed at the Molhe Oeste Salt Marsh. In the images, typical morphologies are indicated with letter and arrows showing isolated bacteria (B) or in colonies (C), diatoms frustules (DF), pennate diatoms (PD), filamentous fungi (FF). Scale bar: 5 μm
Taxonomic composition and diversity indices
In total, 28 bacterial classes (Fig. 7A) and 20 fungal classes (Fig. 7B) were found. The Alphaproteobacteria, Bacilli and Gammaproteobacteria classes were the most frequent, varying between the zones (Fig. 7A). Alphaproteobacteria was more frequent in the flooded zone (69.16%), Bacilli in the intermediate zone (30.13%) and Gammaproteobacteria in the dry zone (39.48%) (Fig. 7A). For families of bacteria, 105 families were found, with Moraxellaceae (23.16%) and Rhodobacteraceae (14.84%) being the most frequent for the flooded zone (Table S2). For the intermediate zone, the most frequent families were Enterobacteriaceae (32.68%) and Lactobacillaceae (30.13%), exclusive to this zone (Table S2). While for the dry zone were Rhodobacteraceae (17.80%) and Saprospiraceae (4.29%) (Table S2). The Sphingomonadaceae family was one of the most frequent in the three zones, with 24.16% in the flooded zone, 8.29% in the intermediate zone and 49.95% in the dry zone.Fig. 7. Frequency of taxa (%) in plastisphere samples from different salt marsh zones (flooded, intermediate, dry). A 16S-Bacteria, B 18S-Fungi, Rich—Richness for each zone
Bacterial community diversity indices showed variation along the salt marsh flooding gradient (Table 1). The highest bacterial richness was observed in the intermediate zone (n = 24), followed by the flooded zone (n = 23), while the lowest richness occurred in the dry zone (n = 20). The highest diversity was recorded in the intermediate zone (H = 1.90), decreasing in the dry (H = 1.62) and flooded zones (H = 1.38) (Table 1). Equitability was highest in the intermediate zone (J = 0.60), followed by the dry zone (J = 0.54) and the flooded zone (J = 0.44). The dominance index was highest in the flooded zone (D = 0.49) and lowest in the intermediate zone (D = 0.23) (Table 1).Table 1. Diversity index of the plastisphere in the different zones (dry, intermediate and flooded) of the salt marsh for fungal and bacterial communitiesZonesDryIntermediateFloodedBacteriaRichness (n)202423Shannon (H)1.621.901.38Dominance (D)0.270.230.49Equitability (J)0.540.600.44FungiRichness (n)13148Shannon (H)1.311.581.09Dominance (D)0.410.240.45Equitability (J)0.510.600.53
For fungi, the most frequent class in the dry zone was Dothideomycetes (61.00%), Sordariomycetes (33.48%) for the intermediate zone, and Euromycetes (62.88%) in the flooded zone (Fig. 7B). As for the fungal families, a total of 51 families were found, with Phaeosphariaceae (49.02%) and Trichosphacriaceae (5.92%) being more frequent in the dry zone (Table S3). In the intermediate zone, the most frequent were Mortierellaceae (18.74%) and Phaeosphariaceae (17.78%), while in the flooded zone were Phaeosphariaceae (5.04%) and Dothioraceae (2.43%) (Table S3). The Aspergillaceae family was the most frequent in all zones, with frequencies of 34.99% in the flooded zone, 20.86% in the intermediate zone, and 5.83% in the dry zone (Table S3).
Fungal community diversity indices also varied along the salt marsh flooding gradient (Table 1). The highest fungal richness was observed in the intermediate zone (n = 14), followed by the dry zone (n = 13), whereas the lowest richness occurred in the flooded zone (n = 8). The highest diversity was found in the intermediate zone (H = 1.58), with lower values in the dry (H = 1.31) and flooded zones (H = 1.09) (Table 1). Equitability was highest in the intermediate zone (J = 0.60), followed by the flooded (J = 0.53) and dry zones (J = 0.51). The dominance index was highest in the flooded (D = 0.45) and dry zones (D = 0.41), and lowest in the intermediate zone (D = 0.24) (Table 1).
NMDS ordination and core microbiome
NMDS ordination based on Bray–Curtis dissimilarity indicated differentiation in the composition of bacterial (Fig. 8A, stress = 0) and fungal (Fig. 8B, stress = 0) communities associated with the plastisphere across salt marsh zones. For both communities, the dry, intermediate, and flooded zones showed separation in the ordination space, suggesting variation in microbial community composition along the flooding gradient (Fig. 8). The intermediate zone was more clearly separated from the other zones (Fig. 8), indicating a distinct community composition. Although a stress value of zero indicates an excellent representation of the observed dissimilarities, we emphasize that, due to the limited number of samples, the analysis should be interpreted cautiously and as exploratory.Fig. 8. Non-metric multidimensional scale ordering (NMDS) with Bray–Curtis similarity index for the abundance of plastisphere communities. A Bacterial community composition and B Fungal community composition
The heatmap of the core plastisphere microbiome revealed variation in the relative abundance of bacterial and fungal classes shared among salt marsh zones (Fig. 9). For bacteria (Fig. 9A), Alphaproteobacteria and Bacilli were the most abundant classes, with zone-specific differences, including higher relative abundance of Alphaproteobacteria in the flooded zone and Bacilli in the intermediate zone (Fig. 9A). Other bacterial classes occurred at lower proportions and showed relatively homogeneous distributions across zones (Fig. 9A). For fungi (Fig. 9B), the core microbiome was dominated by Dothideomycetes, Eurotiomycetes, and Sordariomycetes, with shifts in relative abundance along the flooding gradient. Dothideomycetes were more abundant in the dry zone, Sordariomycetes in the intermediate zone, and Eurotiomycetes in the flooded zone (Fig. 9B). Hierarchical clustering reflected these differences in core microbiome composition among zones for both bacterial and fungal communities (Fig. 9).Fig. 9. Heatmap of the core plastisphere microbiome across salt marsh zones. A Bacterial classes and B fungal classes with > 0% occurrence in the dry, intermediate, and flooded zones. Colors indicate relative abundance (%), and hierarchical clustering reflects similarity in community composition among zones
Discussion
Polymer size
Regarding hypothesis H4, our results demonstrated contrasting responses between micro- and macro-organisms. Smaller substrates favored initial microbial colonization and biofilm accumulation, whereas larger substrates supported a higher abundance of macroorganisms. Polymer size significantly influenced plastisphere development. Smaller substrates (6 × 2 mm) supported higher biofilm formation, evidenced by increased bacterial density and elevated concentrations of photosynthetic pigments from microalgae (chlorophyll a), diatoms (chlorophyll c₁, c₂, and fucoxanthin), and chlorophytes (chlorophyll b and lutein). These findings were corroborated by scanning electron microscopy (SEM), which revealed dense microbial colonization and spatial co-occurrence of diatoms and bacteria, suggesting potential biological interactions within the biofilm matrix (Schlundt et al. 2019). This integrative evidence indicates that smaller polymer sizes favor early colonization and biofilm establishment, likely due to their higher surface area to volume ratio and reduced spatial heterogeneity, providing greater opportunities for microbial attachment and metabolic activity.
These patterns were also observed in broader comparisons of microbial communities across polymers ranging from 0.3 to 200 mm (Debroas et al. 2017). However, within narrower size ranges, such as 0.3–5 mm, no clear effect of polymer size on plastisphere composition has been reported (Frère et al. 2018; Wen et al. 2020). This suggests that size dependent responses may occur only when contrasts between substrates are sufficiently large, rather than within the microplastic size range commonly used in ecological assessments.
Most previous studies have relied predominantly on molecular approaches to characterize community structure (Shruti et al. 2024), with limited incorporation of quantitative indicators of biofilm development. As a result, metrics such as cellular density and photosynthetic pigment concentrations remain underexplored in plastisphere research. By integrating community-level data with quantitative measures of biofilm biomass (e.g., bacterial density and chlorophyll profiles), our study provides a complementary perspective on plastisphere development, indicating that microbial composition alone may not fully reflect the magnitude of biofilm accumulation on plastic substrates.
In contrast to the patterns observed for microfouling, macrofouling organisms demonstrated the opposite trend, with higher abundances on the largest substrates (60 × 20 mm). This result was particularly associated with the predominance of Cirripedia and Bivalvia, indicating a preference for substrates offering larger attachment areas. Such preference may be related to the greater availability of space for growth, whereas smaller substrates impose physical constraints that can intensify competition for settlement sites (Fazey and Ryan 2016). Accordingly, plastisphere biomass was highest on the largest polymers, reinforcing the relationship between available surface area and macrofouling accumulation. Although polymer size was the main driver identified here, additional substrate characteristics (e.g., surface roughness, topography, and hydrophobicity) and external factors such as color (Wen et al. 2020) may further modulate colonization patterns (Chen et al. 2020). These features likely operate in combination, suggesting that plastisphere formation reflects the interplay between organism-specific traits and substrate-dependent physicochemical properties.
Polymer color
According to Dobretsov et al. (2013) and Satheesh and Wesley (2010), dark-colored substrates generally support higher plastisphere biomass than lighter ones. Our results only partially align with this pattern: the white polymer showed the greatest overall biomass, whereas only diatom-related pigments (chlorophyll c₂ and fucoxanthin) and small invertebrate abundance were higher on the red polymer. This indicates that color influences specific taxonomic groups rather than uniformly affecting biofilm development, likely reflecting tax on specific settlement preferences (Wen et al. 2020). Color-dependent settlement may be driven by distinct ecological mechanisms, including phototropic behavior (Railkin 2004), thermal properties related to light absorption and reflection (Dobretsov et al. 2013), or the chemical composition of pigments and additives (Hermabessiere et al. 2017). However, the effects of polymer color on plastisphere formation remain poorly constrained, and current evidence suggests that color interacts with other factors, such as polymer type and environmental conditions, rather than operating as an isolated driver.
Hypothesis H3 was only partially supported by our results. Although darker-colored polymers, such as red, showed higher concentrations of some photosynthetic pigments and greater abundance of specific macroorganisms, bacterial density and microbial community composition did not differ consistently among colors. These findings suggest that polymer color may influence particular organism groups but does not exert a dominant effect on the overall structure of the plastisphere.
Polymer type
Different polymer types can modulate colonization by plastisphere associated taxa (Miao et al. 2019; Sooriyakumar et al. 2022). In our experiment, the polymers tested (EVA, PS, and PP) did not influence bacterial density; however, diatoms and chlorophytes showed a clear preference for EVA, as indicated by higher concentrations of photosynthetic pigments (lutein, chlorophyll c₁/c₂). This suggests that polymer type affected microalgal colonization more strongly than bacterial attachment under the conditions tested. One explanation is related to the temporal scale of biofilm development: bacteria are early colonizers and may disperse after maturation, whereas algal adhesion tends to increase in later stages of biofilm succession (Sauer et al. 2022; Ma et al. 2017), which aligns with the 21-day duration of our experiment. Similar patterns were observed by Oberbeckmann et al. (2021) in a 14-day experiment and by Pinto et al. (2019) in a 30-day experiment, where bacterial signals appeared reduced at later stages due to community turnover and dispersion.
SEM observations indicated that EVA and PS exhibit more heterogeneous surface structures (pores, irregularities, and indentations), while PP presented a smoother surface. These characteristics likely increased the availability of microhabitats for algal attachment on EVA, consistent with evidence that surface roughness and structural complexity enhance colonization (Sooriyakumar et al. 2022). This pattern aligns with Da Silva et al. (2023a), who demonstrated that EVA undergoes structural deformation more readily than PP and PS under environmental exposure, increasing its surface heterogeneity. Therefore, the preference observed for EVA is likely linked to its physical properties rather than its chemical composition alone.
On the other hand, small invertebrates exhibited a clear preference for EVA, reflected in the higher plastisphere weight recorded for this polymer. This response may be associated with both organismal traits, such as life history strategies and adaptive structures, and substrate-dependent factors, including roughness and hydrophobicity (Hung et al. 2007; Sooriyakumar et al. 2022). Surfaces with greater structural complexity, such as cracks, crevices, and depressions, can facilitate colonization by providing protected microhabitats and increasing attachment stability (Sooriyakumar et al. 2022), particularly in intertidal systems where wave action and desiccation impose strong selective pressures on colonizing communities. This combination of organism-level and substrate-level mechanisms likely explains the higher invertebrate abundance on EVA, rather than a single isolated driver.
The physical contrasts detected by SEM between polymers (heterogeneous surfaces in EVA and PS versus smoother surfaces in PP) support the biological patterns observed in our experiment. EVA sustained higher abundances of diatoms, chlorophytes, and small invertebrates, consistent with the idea that surface complexity increases available adhesion sites and promotes retention of colonizing organisms (Miao et al. 2021; Sooriyakumar et al. 2022). This convergence between structural observations and colonization patterns indicates that polymer specific microtopography contributes to microhabitat availability, reducing detachment and enabling the persistence of biofilms and associated fauna under estuarine conditions. Together, these results reinforce that the preference for EVA is likely driven by physical surface properties rather than chemical composition alone, aligning with patterns described for environmentally exposed materials (Silva et al. 2023a; Hung et al. 2007). These patterns suggest that surface microtopography may facilitate the retention of biofilms and associated organisms, although analyses of community composition are essential to complement this finding.
With respect to hypothesis H1, our results indicate that polymer type influenced the intensity of colonization and biofilm accumulation, particularly for microalgae and macrofouling organisms, with EVA showing the strongest response. SEM observations revealed greater surface heterogeneity in EVA and PS compared to PP, which likely increased the availability of microhabitats for organism attachment. However, community-level analyses of microbial composition indicated that polymer type had a secondary effect when compared to the flooding regime, suggesting that structural differences among polymers modulate, but do not solely determine, plastisphere community composition.
Flooding zones
Salt marsh zones in the study area differ primarily in flooding regime and associated vegetation structure (Pinheiro et al. 2022, 2021). Across these zones, plastisphere development showed a clear gradient, with the flooded zone exhibiting the highest values for all biological parameters, followed by the intermediate zone, and the lowest levels occurring in the dry zone. A similar trend was described by Pinheiro et al. (2021) for macroorganisms on macroplastics, indicating that greater water availability enhances substrate colonization, particularly for taxa that depend on immersion for attachment and survival. This pattern is consistent with the expectation that flooded conditions facilitate biofilm establishment and reduce desiccation stress, creating a more favorable environment for plastisphere formation.
Environmental gradients across salt marsh zones, such as salinity, nutrient availability, flood frequency, and solar irradiation, are known to influence community structure in estuarine systems (Pinheiro et al. 2021; Seeliger and Odebrecht 2010). Although these parameters were not measured in the present study, our results demonstrate that plastisphere development followed a spatial gradient aligned with the flooding regime, with higher values in the flooded zone and lower values in the dry zone. Previous research has shown that abiotic conditions can modulate the composition and stability of plastisphere communities (Oberbeckmann et al. 2021), suggesting that the patterns observed here may reflect the indirect effects of environmental gradients associated with marsh zonation. However, the absence of direct measurements prevents causal interpretations, and further studies are required to evaluate which specific variables govern plastisphere responses across salt marsh environments.
NMDS ordination revealed clear separation among zones, with the intermediate zone exhibiting the most distinct community composition, as supported by higher richness, diversity, and evenness indices. This pattern may be linked to the physiological characteristics of colonizing organisms and their species-specific tolerances. Algae tend to prefer continuously flooded environments, such as the flooded zone in this study, as these conditions minimize desiccation stress and photoinhibition (Holzinger and Karsten 2013). Diatoms, in turn, exhibit high substrate versatility due to their opportunistic nature, physiological adaptability, and specialized morphological traits that facilitate adhesion (França et al. 2011), even under high hydrodynamic disturbance and fluctuating water levels (Ruggieri et al. 2006).
The heatmap of the core microbiome showed that differences among zones were driven by shifts in relative abundance of shared taxa rather than taxonomic turnover, indicating environmental filtering along the flooding gradient. Proteobacteria was the most abundant bacterial phylum in our samples, a pattern consistent with reports from marine and estuarine plastisphere studies in different regions (Bryant et al. 2016; Lacerda et al. 2022; Sooriyakumar et al. 2022; Wallbank et al. 2022; Zettler et al. 2013). Members of this phylum are recognized as early and versatile colonizers of submerged substrates, displaying high phylogenetic diversity and broad metabolic and ecological adaptability (Dang and Lovell 2016; Salta et al. 2013; Zinger et al. 2011). In our dataset, Alphaproteobacteria dominated in the flooded zone, whereas Gammaproteobacteria were more prevalent in the dry zone, suggesting that flooding regime, and the environmental gradients associated with it, may influence community assembly. Although the absence of direct environmental measurements prevents causal inference, these findings align with previous research indicating that salinity, immersion time, and hydrodynamics can shape plastisphere structure at the phylum level (Oberbeckmann et al. 2021).
At the family level, clear distribution patterns were observed for both bacteria and fungi across zones. Sphingomonadaceae increased toward the flooded zone, whereas Moraxellaceae were more frequent in the dry zone, and Enterobacteriaceae peaked in the intermediate zone. These trends suggest that flooding regime may filter taxa based on tolerance to immersion and desiccation, although the absence of environmental measurements in this study prevents causal interpretation. Moraxellaceae have been reported as key members of microbial communities due to their roles in material transformation and nutrient cycling (Agler et al. 2016; Li et al. 2021), with representatives capable of tolerating low temperature (Dias et al. 2018) and salt stress (Li et al. 2021). Their higher frequency in the dry zone is consistent with this tolerance profile and may indicate better adaptation to reduced immersion. Conversely, Enterobacteriaceae, detected mainly in the intermediate zone, include clinically relevant taxa that can carry antibiotic resistance genes (Gao et al. 2025; Silva et al. 2023b) and are considered a surveillance priority for antimicrobial development (Murray et al. 2022), highlighting the need for closer monitoring of plastisphere associated pathogenic reservoirs.
Sphingomonadaceae, prevalent in the flooded zone, include strictly aerobic and chemoheterotrophic taxa commonly found in aquatic environments (Vaz-Moreira et al. 2011). Their capacity to degrade xenobiotic compounds, including bisphenol A (Oh and Choi 2019), and their consistent detection in plastic associated biofilms (Di Pippo et al. 2020; Oberbeckmann et al. 2016) positions them as potential contributors to plastic biodegradation in salt marshes (Liu et al. 2023). Although this study did not evaluate degradation processes directly, their spatial distribution suggests that inundated areas may facilitate conditions under which biodegradation-associated taxa are more active or competitive.
Knowledge about fungal assemblages within the plastisphere remains limited compared to bacterial communities. In our samples, Ascomycota was the dominant phylum across all zones, represented primarily by Dothideomycetes and Eurotiomycetes. This pattern aligns with previous findings in estuarine plastisphere studies, where Ascomycota has frequently been reported as the most abundant fungal group (Lacerda et al. 2022; Sérvulo et al. 2023a, b). At the family level, Phaeosphaeriaceae increased toward the dry zone, while Aspergillaceae showed higher frequencies in flooded areas, indicating that flooding regime may structure fungal distribution in salt marsh plastisphere communities. Some members of both families have been associated with plastic degradation or transformation (Ekanayaka et al. 2022; Munir et al. 2018), raising the hypothesis that environmental conditions could influence the activity or competitiveness of these taxa. However, because biodegradation processes were not assessed in this study, these interpretations remain speculative, and future research incorporating functional or metabolic assays will be necessary to determine whether these spatial patterns translate into ecological functions.
Notably, the genus Aspergillus (Aspergillaceae), detected in flooded zones of this study, has been identified by the World Health Organization as a priority group of emerging concerns due to its pathogenic potential and relevance to human health. Although the detection of Aspergillus spp. does not imply pathogenic activity, its occurrence highlights the relevance of considering plastisphere-associated fungi in future risk assessments. Recent analyses highlight its virulence and antimicrobial resistance capacity (Andrade et al. 2025), underscoring the need for continued monitoring. While the present study did not evaluate pathogenicity or resistance genes, our findings emphasize the importance of considering potential health implications, particularly given the limited understanding of fungal pathogenicity within plastisphere environments (Ormsby et al. 2023).
Overall, our results provide strong support for hypothesis H2, as the flooding regime emerged as the main factor structuring plastisphere communities. Diversity indices, NMDS ordination, and core microbiome analyses consistently indicated differentiation in bacterial and fungal community composition among dry, intermediate, and flooded zones. These differences were driven primarily by shifts in the relative abundance of shared taxa rather than by taxonomic turnover, suggesting an environmental filtering effect along the flooding gradient.
Limitations and perspectives
Salt marshes located within estuarine environments are of high ecological importance. However, in the extreme south of South America, these habitats and adjacent regions have been identified as hotspots for plastic contamination (Pinheiro et al. 2022, 2021; Ramos et al. 2021). This issue is further exacerbated by the formation of the plastisphere on plastic debris, which can alter the ecological dynamics of aquatic ecosystems by acting as a vector for emerging contaminants (Richard et al. 2019; Silva et al. 2023b), pathogenic species, and antibiotic resistance genes (Wu et al. 2024), as well as by disrupting biogeochemical cycles (Chen et al. 2020). To our knowledge, this is the first study to assess the influence of different plastic characteristics (size, color, and polymer type) on plastisphere formation in the Patos Lagoon Estuary, the largest shallow coastal lagoon in the world. Nevertheless, some limitations of the present study should be acknowledged:
- Environmental variables were not measured during the experiments, which prevents identifying which specific abiotic factors (e.g., salinity, temperature, nutrient availability, and hydrodynamics) drive the spatial patterns observed across flooding zones. These environmental parameters vary seasonally, and the three field experiments were conducted in different periods of the year. Therefore, seasonal variability may have influenced biofilm development and community composition, potentially contributing to some of the observed differences among treatments. This limitation should be considered when interpreting the results, and future studies should aim to couple experimental manipulations with continuous measurements of environmental variables across seasons.
- Only three polymer types and colors were assessed, without considering degradation stage or biodegradability. A broader matrix of polymer conditions could clarify how physicochemical properties shape plastisphere structure.
- Metabarcoding used pooled samples, which limited taxonomic resolution and prevented comparisons across polymer characteristics. Future studies should prioritize individual sample sequencing to assess community variability and functional profiles.
- These limitations are particularly relevant for the interpretation of community-level analyses, which were therefore treated as descriptive and exploratory.
Despite these constraints, our findings establish a basis for understanding plastisphere dynamics in salt marshes at the southernmost limit of the Southwestern Atlantic. Future studies combining environmental measurements, temporal monitoring, functional or metabolic assays, and expanded molecular coverage (e.g., metagenomics and resisted analysis) will be essential to determine how environmental gradients and polymer properties jointly regulate plastisphere composition, stability, and potential ecological or health implications.
Conclusion
This study is the first to evaluate the combined influence of flooding zones and polymer characteristics on plastisphere formation in a salt marsh within the Patos Lagoon Estuary, located in the extreme south of Brazil. Our findings provide novel insights into plastisphere composition over a 21-day period and how plastic contamination may impact the ecological dynamics of this estuarine ecosystem. The results demonstrate that plastisphere development is strongly influenced by the flooding regime (dry, intermediate, and flooded zones), as evidenced by variations in plastisphere weight, bacterial density, photosynthetic pigment concentrations, macroorganism colonization, and microbial community composition derived from metabarcoding analyses. The flooded zone exhibited the highest values for these endpoints; however, species diversity was lower in this zone compared to the intermediate zone, which showed greater diversity. Additionally, we detected a high frequency of Moraxellaceae (bacteria) and the fungal families Phaeosphaeriaceae and Aspergillaceae across the different zones, all of which include taxa with pathogenic potential. Polymer properties also influenced plastisphere formation. For instance, diatoms showed a preference for black and red polymers, and both diatoms and chlorophytes exhibited higher colonization on EVA, while small invertebrates were more frequently associated with red polymers. Nevertheless, further studies employing metabarcoding approaches are needed to confirm and expand upon these findings.
Supplementary Information
Below is the link to the electronic supplementary material.Supplementary file1 (DOCX 822 kb)
The reference list from the paper itself. Each links out to its DOI / PubMed record.
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