# Experimental Evidence from the Field that Naturally Weathered Microplastics Accumulate Cyanobacterial Toxins in Eutrophic Lakes

Eden Hataley, René Shahmohamadloo, Xavier Ortiz Almirall, Anna Harrison, Chelsea Rochman, Shan Zou, Diane Orihel

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## To cite this version:

Eden Hataley, René Shahmohamadloo, Xavier Ortiz Almirall, Anna Harrison, Chelsea Rochman, et al.. Experimental Evidence from the Field that Naturally Weathered Microplastics Accumulate Cyanobacterial Toxins in Eutrophic Lakes. Environmental Toxicology and Chemistry, 2022, 41 (12), pp.3017-3028. 10.1002/etc.5485. hal-04917798

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Eden Hataley ORCID iD: 0000-0002-4347-4201

Rene Shahmohamadloo ORCID iD: 0000-0002-5373-9808

Diane Orihel ORCID iD: 0000-0002-6933-3650

Hataley et al.

Microplastic pollution and harmful algal blooms

Experimental Evidence from the Field that Naturally Weathered Microplastics

Accumulate Cyanobacterial Toxins in Eutrophic Lakes

Eden K. Hataley,ª0000-0002-4347-4201 René S. Shahmohamadloo,ᵇ.c0000-0002-5373-9808 Xavier Ortiz Almirall,ª,d,e0000-0002-5079-9506 Anna L. Harrison,f0000-0002-9145-4555 Chelsea M. Rochman,g Shan Zou,h0000-0002-2480-6821 and Diane M. Orihela,i*0000-0002-6933-3650

ª School of Environmental Studies, Queen's University, Kingston, ON, K7L 3N6, Canada

ᵇ School of Biological Sciences, Washington State University, Vancouver, WA, 98686, United States

ᶜ Department of Integrative Biology, University of Guelph, Guelph, ON, N1G 2W1, Canada

ᵈ Ontario Ministry of the Environment, Conservation and Parks, Etobicoke, ON, M9P 3V6, Canada

ᵉ IQS School of Engineering, Universitat Ramon Llull, Via Augusta, 390, 08017, Barcelona

This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version and the Version of Record. Please cite this article as doi: 10.1002/etc.5485.
f Géosciences Environnement Toulouse, UMR 5563, Centre National de la Recherche Scientifique (CNRS), Toulouse, France, 31400

g Department of Ecology and Evolutionary Biology, University of Toronto, St. George Campus, Toronto, ON, Canada, M5S 3B2

h Metrology Research Centre, National Research Council Canada, Ottawa, ON, Canada, K1A 0R6

i Department of Biology, Queen's University, Kingston, ON, Canada K7L 3N6

*Correspondence diane.orihel@queensu.ca

7/5/22; 8/11/22; 9/16/22

**Abstract:** Freshwater ecosystems with recurring harmful algal blooms can also be polluted with plastics. As such, the two environmental problems may interact. To test if microplastics influence the partitioning of microcystins in freshwater lakes, we examined the sorption of four microcystin congeners to different polymers of commercially available plastics (low-density polyethylene, polyethylene terephthalate, polyvinyl chloride, and polypropylene). We conducted three experiments: a batch sorption experiment in the laboratory with pristine microplastics of four different polymers, a second batch sorption experiment in the laboratory to compare pristine and naturally weathered microplastics of a single polymer, and a 2-month sorption experiment in the field with three different polymers experiencing natural weathering in a eutrophic lake. This series of experiments led to a surprising result: microcystins sorbed poorly to all polymers tested under laboratory conditions (less than 0.01% of the initial amount added), irrespective of weathering, yet, in the field experiment, all polymers accumulated microcystins under ambient conditions in a eutrophic lake (range: 0-84.1 ng/g).
Furthermore, the sorption capacity for microcystins differed among polymers in the laboratory experiment yet were largely the same in the field. We also found that the affinity for plastic varied among microcystin congeners, namely more polar congeners demonstrated a greater affinity for plastic than less polar ones. This study improves our understanding of the role of polymer and congener type in microplastic-microcystin sorption, and moreover, provides novel evidence from the field, showing that naturally weathered microplastics in freshwater lakes can accumulate microcystins. Consequently, we caution that microplastics may alter the persistence, transport, and bioavailability of microcystins in freshwaters, which could have implications for human and wildlife health.

**Keywords:** absorption; adsorption; algal toxins; biofilm; freshwater toxicology; microplastics

This article includes online-only Supporting Information.

*Address correspondence to diane.orihel@queensu.ca

Published online XXXX 2022 in Wiley Online Library (www.wileyonlinelibrary.com).

DOI: 10.1002/etc.xxxx

## INTRODUCTION

Plastic pollution is increasingly common in freshwater ecosystems with pervasive harmful algal blooms (HABs) (Earn et al., 2021; Egessa et al., 2020; Zhang et al., 2020).

The interplay between these globally important environmental issues (i.e., plastic pollution and eutrophication) is critical to understand as plastics, particularly microplastics (<5 mm), could potentially exacerbate the problem of freshwater HABs.

For example, microplastics may act as a dispersal vector for bloom-forming

This article is protected by copyright. All rights reserved.
cyanobacteria (Masó et al., 2003; Naik et al., 2019; Oberbeckmann et al., 2014; Yokota et al., 2017; Zettler et al., 2013), expanding the spatial and temporal extent of HABs.

Furthermore, microplastics may influence the growth of toxin-producing cyanobacteria or the production and release of toxins by these cyanobacteria (Sánchez-Fortún et al., 2021; Wan et al., 2021; Wu et al., 2021; Zhou et al., 2021). An emerging concern and focus of the present work is whether microplastics can influence the environmental fate of waterborne algal toxins released by cyanobacteria. Microplastics are known to act as sinks and vectors for environmental contaminants (Bellasi et al., 2020; Caruso, 2019; Hartmann et al., 2017), and have the potential to change a number of environmental behaviours of algal toxins (e.g., partitioning, transport, persistence, and bioavailability).

If this is indeed the case, plastic pollution in freshwater ecosystems could possibly alter the exposure of humans and wildlife to these harmful toxins.

One mechanism by which microplastics can influence the fate of waterborne contaminants is through the process of sorption. This interaction has been documented for a wide range of environmental contaminants, including polycyclic aromatic hydrocarbons (Frias et al., 2010; Rios et al., 2007; Rochman et al., 2013), polychlorinated biphenyls ((Endo et al., 2005; Frias et al., 2010; Mato et al., 2001; Rios et al., 2007; Rochman et al., 2013), metals (Ashton et al., 2010; Richard et al., 2019; Rochman et al., 2014), and pharmaceuticals (Magadini et al., 2020). Recent work demonstrates microplastics can sorb microcystins (Moura et al., 2022; Pestana et al., 2021; Qu et al., 2019), a potent hepatotoxin produced by freshwater HABs that can have deleterious health effects on humans (Carmichael, 2001) and wildlife (Gene et al., 2019; Shahmohamadloo et al., 2021).
Microcystins are a family of over 300 monocyclic heptapeptides (Bouaïcha et al., 2019). The pervasive success of microcystins in eutrophic freshwaters can be attributed to various factors including their structural diversity and physicochemical properties (Ortiz et al., 2017; Puddick et al., 2014; Zastepa et al., 2014). Microcystins vary in polarity depending on their amino acid composition (e.g., number of arginine moieties conferring polarity), which in turn dictates their sorption potential to surfaces including sediments (Harada & Tsuji, 1998; Miller et al., 2001; X. Wu et al., 2011; Zastepa et al., 2014, 2017) as well as glass and plastics (Altaner et al., 2017; Hyenstrand et al., 2001). Cation-bridging, hydrogen-bonding, and electrostatic interactions are some of the sorption mechanisms responsible for the interaction between microcystins and hydrophobic phases in water (Altaner et al., 2017; Newcombe et al., 2003; X. Wu et al., 2011), mechanisms which are also responsible for the sorption of waterborne hydrophilic contaminants, such as antibiotics, to microplastics (Li et al., 2018; Liu et al., 2019; Zhang et al., 2018). As freshwater HABs are projected to increase in duration under a warming climate (Chapra et al., 2017), the frequency and occurrence of microcystins in global freshwaters may likewise increase, affording more opportunity for interaction with microplastics.

While recent work has demonstrated microcystin congeners quickly adsorb to pristine microplastics in artificial freshwater laboratory assays (He et al., 2022; Moura et al., 2022; Pestana et al., 2021), to our knowledge no study has resolved if weathered microplastics affect the environmental fate of microcystins in surface waters of natural ecosystems. Once plastics enter the aquatic environment, their properties immediately begin to change as a result of weathering processes (Arp et al., 2021), which include
abiotic and biotic weathering, as well as mechanical degradation and fragmentation.

Weathering processes can significantly change the physicochemical properties of plastics (e.g., specific surface area, crystallinity, and oxygen content of surface functional groups), and thereby influence their sorption behaviour and other interactions with co-pollutants (Duan et al., 2021; P. Liu et al., 2020). Biofilm formation may likewise influence the kinetics of uptake and release of contaminants by microplastics (Rummel et al., 2017). Consequently, it is prudent to resolve if natural weathering of microplastics modifies their sorption capacity for microcystins in freshwater ecosystems afflicted by HABs.

In this study, we assessed the sorption of four structurally diverse and recurring microcystins {microcystin-arginine-arginine (MC-RR), microcystin-tyrosine-arginine (MC-YR), microcystin-leucine-arginine (MC-LR), and microcystin-leucine-alanine (MC- LA)} to four types of commercially available plastics {low-density polyethylene (LDPE), polyethylene terephthalate (PET), polyvinyl chloride (PVC), and polypropylene (PP)}.

Because microplastic-contaminant sorption is influenced by the physicochemical properties of both the microplastic and the contaminant as well as the conditions of the natural environment (Hartmann et al., 2017), we conducted two batch sorption laboratory experiments, one with unweathered microplastics of four different polymers (Exp. 1) and one with a single polymer exposed to different weathering conditions (Exp. 2), and a sorption field experiment with three different polymers in a eutrophic lake (Exp. 3). This research was done to test the following hypotheses: sorption capacity for microcystins differs among polymer types ($H_1$), affinity for plastic varies among microcystins ($H_2$),
and weathered microplastics have a greater sorption capacity for microcystins compared
to unweathered microplastics (H3).

MATERIALS AND METHODS

Laboratory batch sorption experiment with unweathered microplastics (Exp. 1)

The first experiment was conducted at the Ontario Ministry of the Environment,
Conservation and Parks (MECP; Etobicoke, Ontario, Canada) and consisted of five
experimental groups (6 replicates/group): four treatments of different polymers (LDPE,
PET, PVC, and PP) and a negative control (no added microplastics). Thirty 1-L
deactivated borosilicate clear glass bottles were filled with 500 mL of ultrapure water
(conductivity < 2 mS/cm at 25°C, total organic carbon < 2 ppm) and 500 microplastic
particles, measured gravimetrically. Each bottle was spiked with 2.5 µg each of MC-RR,
MC-YR, MC-LR, and MC-LA (all ≥ 95% pure; Enzo Life Sciences, Farmingdale, New
York, USA) to achieve a final concentration of 20 µg/L total microcystins, which is
representative of the World Health Organization's guideline value for maximum
acceptable levels of total microcystins (expressed as MC-LR equivalent) in recreational
waters (WHO, 2020). Bottles were rotated continuously at 20°C throughout the
experiment for 48 h. An exposure period of 48 h was chosen because (1) a pilot study
showed the microplastic-microcystin equilibrium was reached within 48 h (data not
shown) and (2) many other laboratory-based microplastic-contaminant sorption studies
employ a similar exposure period (Guo et al., 2018; Liu et al., 2019; Zhang et al., 2018).
At 0, 1, 2, 4, 8, 16, 32, and 48 h, 5 mL of solution and 50 microplastics were collected
from each bottle for microcystin analysis. Solution subsamples were stored in
polypropylene tubes at -40 °C and microplastic subsamples were stored in polypropylene tubes with 10 mL of methanol at room temperature (20°C).

*Laboratory batch sorption experiment with weathered microplastics (Exp. 2)*

The second experiment, also performed at the MECP laboratory, consisted of five
experimental groups (6 replicates/group): three treatments of one polymer under different
weathering conditions (weathered LDPE in filtered lake water, unweathered LDPE in
filtered lake water, and unweathered LDPE in ultrapure water), a negative filtered lake
water control, and a negative ultrapure water control. Filtered lake water (nylon mesh,
100 µm) obtained from Buck Lake was used to increase the environmental relevance of
the experiment. LDPE microplastics were chosen because they demonstrated sorption
capacity for all microcystin congeners tested in Exp. 1. Microplastics were weathered
from 1 July to 15 September 2019 in Buck Lake (44°32′51.4″ N 76°26′09.8″ W, Ontario,
Canada), a small (surface area: 755 ha, volume: 25.9×10⁶ m³) and deep (depth: mean 11.9
m, maximum 40.9 m) mesotrophic lake that is not prone to cyanobacterial HABs and
therefore, microcystin-contaminated microplastics or lake water samples were unlikely to
interfere with the experiment. We confirmed no microcystins in weathered microplastics
and lake water samples. Microplastics were enclosed in a wooden frame wrapped in a
black fiberglass mesh screen (1 mm²), which was suspended from stainless steel wire
fixed to two floating docks and submerged approximately 0.1 m below the surface of the
water (Figure S1, Supporting Information). Microplastics were retrieved from Buck Lake
36 h prior to the experiment and transported in lake water at ambient temperature. This
experiment followed the same procedure as Exp. 1 (i.e., microplastics were exposed to 20
µg/L total microcystins consisting of MC-RR, MC-YR, MC-LR, and MC-LA), except
subsamples were collected at 0, 1, 2, 4, 8, 16, 32, 64, and 128 h to ensure a state of
sorption equilibrium was reached as the pilot study referenced above was conducted with
unweathered microplastics only.

Field-based sorption experiment (Exp. 3)

The third experiment was conducted in Dog Lake (44°23′43.9′′ N 76°21′50.0′′ W, Ontario, Canada), a small (surface area: 964 ha, volume: 55.4×10⁶ m³) eutrophic lake.

This lake has a deep northern basin (depth: maximum 50 m) and a shallow southern basin (depth: <3 m), where HABs are pervasive during warmer months. In the past decade, total microcystins in surface water concentrations of this lake have ranged from <0.05 to >7 µg/L (MECP, personal communication, August 26, 2022). This experiment was conducted at two sites in the littoral zone of the southern basin where HABs are known to recur.

Experiment 3 consisted of four experimental groups (3 replicates/group): three treatments of different polymers (LDPE, PET, and PVC) and a control (water with no microplastics). Following the same experimental set-up as Exp. 2, 18 wooden frames (3 frames × 3 polymer × 2 sites) were deployed in Dog Lake on 21 August 2019 and 250 microplastics were collected from each wooden frame weekly for microcystin analysis between 29 August and 24 October 2019 (excluding 25 September and 2 and 16 October due to adverse weather). Therefore, the microplastics used in Exp. 3 were weathered for a similar period of time (9 weeks) compared to the microplastics used in Exp. 2 (10 weeks). Upon collection, microplastics were rinsed with reverse-osmosis water and gently dried with a Kimwipe™ to remove lake water droplets. Subsurface (0.5 m) water samples were collected in 1-L amber glass bottles in triplicate at the time of microplastic
collection and stored on ice for return to the laboratory. Water samples were
homogenized by manual agitation, and a 40 mL aliquot of whole water was collected for
microcystin analysis. Microplastic and water samples were stored in polypropylene tubes
at -20 °C. Bottles were sterilized between sampling events.

Plastic polymers and characterization

All experiments used recycled LDPE, PET, PVC, and PP (Kal-Polymers, Mississauga, Ontario, Canada). These plastics were made from recaptured post-consumer waste to create pre-production plastic pellets for reusable materials and likely contained a complex blend of chemical additives (see Brosché et al., 2021 for a recent global study). Polymers had different morphologies (Table 1) and identifications were confirmed by Fourier transform infrared (FT-IR) spectrometry (Agilent Cary 630, Santa Clara, California, USA) (Figure 1A, Table S1). Surface areas of each polymer was determined by the Brunauer-Emmett-Teller (BET) method from ultra-high purity nitrogen gas adsorption isotherms at 77 K using an ASAP 2000 surface area and porosity analyzer (Micrometrics, Norcross, Georgia, USA) (Table 1). FT-IR and BET analyses were performed at the Metrology Research Centre, National Research Council Canada (Ottawa, Ontario, Canada). Surface morphology of each polymer was observed by scanning electron microscopy (SEM) (FEI Quanta 250, Hillsboro, Oregon, USA) at Queen's University (Kingston, Ontario, Canada).

Microcystin analysis

Microcystins sorbed to microplastics were extracted in triplicate with 10 mL of methanol. Microcystins are highly soluble in methanol (Altaner et al., 2017), which is why this solvent was chosen to both store and extract the microplastic samples. Samples
were sonicated for 15 min and left to sit at room temperature (20 °C) for 30 min, after
which extracts were transferred to new polypropylene tubes. Extracts were evaporated to
near dryness (<0.5 mL) under a stream of nitrogen gas at 50 °C. Microcystins were
resuspended in a 4.5 mL mixture of ultrapure water/methanol (75:25, V:V). Final
solutions were filtered through a hydrophilic polypropylene membrane (0.45 µm), and 2
mL was collected for microcystin analysis. Extracts from microplastic samples collected
from Dog Lake were diluted 1:4 in a mixture of ultrapure water/methanol (75:25, V:V)
after filtration. Water samples were filtered through a hydrophilic polypropylene
membrane (0.45 µm), and 2 mL was collected for microcystin analysis. Water samples
from Dog Lake required three freeze/thaw cycles (at -40 and 50 °C, respectively) before
filtration to ensure recovery of cell-bound microcystins.

Concentrations of MC-RR, MC-YR, MC-LR, and MC-LA in microplastic extract and water were determined by on-line solid-phase extraction coupled to liquid chromatography-quadrupole time-of-flight mass spectrometry (Water Xevo G2-XS, Milford, Massachusetts, USA) (Ortiz et al., 2017). Nodularin (≥ 95% pure; Enzo Life Sciences, Farmingdale, New York, USA), a cyanobacterial toxin with a monocyclic pentapeptide structure like that of microcystin not commonly present in fresh water, was used as an internal standard. In Exp. 1 and 2, changes in aqueous microcystin concentration were undetectable. Microplastic extract and water from Exp. 3 were also tested for microcystin-homotyrosine-arginine (MC-HtyR), microcystin-homoisoleucine-arginine (MC-HilR), microcystin-tryptophan-arginine (MC-WR), microcystin-leucine-tryptophan (MC-LW), microcystin-leucine-phenylalanine (MC-LF), microcystin-leucine-tyrosine (MC-LY), microcystin-Dha⁷-leucine-arginine (MC-Dha⁷-LR), and microcystin-
Dha⁷-arginine-arginine (MC-Dha⁷-RR). In Exp. 1 and 2, total microcystins (Σ₄MCs) is defined as the sum of the concentrations of MC-RR, MC-YR, MC-LR, and MC-LA. In Exp. 3, total microcystins (Σ₁₂MCs) is defined as the sum of the concentrations of MC-LR, MC-YR, MC-RR, MC-HtyR, MC-HilR, MC-WR, MC-LW, MC-LA, MC-LF, MC-LY, MC-Dha⁷-LR, and MC-Dha⁷-RR. To account for the different morphology of polymers (Table 1), microcystin concentrations were standardized in two different ways: ng microcystin per gram of polymer (ng/g) and ng microcystin per surface area of polymer (ng/m²).

## Statistical Analyses

Data analyses were performed using Sigma Plot 14.5 (SYSTAT Software, Chicago, Illinois, USA). For Exp. 3, differences among polymer types were tested by One-Way Analysis of Variance (ANOVA) on each sampling date. Post hoc Bonferroni tests were used to distinguish significantly different mean values (α = 0.05). Normality and homogeneity of variance were verified by Shapiro-Wilk and Brown-Forsythe tests, respectively. If assumptions were not met, data were analyzed by Kruskal-Wallis One-Way ANOVA on Ranks. Post hoc Dunn's tests were used to distinguish significant differences among experimental groups (α = 0.05).

## RESULTS

### Laboratory batch sorption experiment with unweathered microplastics (Exp. 1)

Microcystins sorbed poorly, or not at all, to four polymers of microplastics. Concentrations of Σ₄MCs (n = 8 sampling events) sorbed to microplastics ranged from 1.1 to 4.1 ng/g for LDPE, 0.2 to 1.0 ng/g for PET, 0.0 to 0.3 ng/g for PP, and 0.0 to 0.2 ng/g for PVC (Figure 2; see Figure S2 for concentrations standardized to surface area).
Maximum concentrations of Σ₄MCs sorbed to microplastics reached within 1 hr, and generally declined thereafter, while microcystins in water samples remained at the initial concentration of 20 µg/L (Table S2). The final mass of Σ₄MCs sorbed to LDPE, PET, PP, and PVC after 48 hr was approximately 0.008%, 0.005%, 0.0005%, and 0.00005%, respectively, of the initial amount added.

Microcystins with higher polarity (Table 2) demonstrated a greater affinity for microplastics than those with lower polarity (Figure 3; see Figure S3 for concentrations standardized to surface area). Affinity of microcystins for all microplastics followed the general trend: MC-RR > MC-YR > MC-LR > MC-LA. Sorption of MC-RR, MC-YR, MC-LR, and MC-LA by LDPE and PET increased within 1 h, after which MC-RR, MC-YR, and MC-LR appeared to reach equilibrium, whereas MC-LA desorbed. Sorption of all microcystins by PP and PVC was largely negligible, barring the sorption of MC-RR by PP, which marginally increased within 1 h, after which MC-RR appeared to reach equilibrium.

*Laboratory batch sorption experiment with weathered microplastics (Exp. 2)*

In the 128-hr sorption experiment with LDPE only, microcystins sorbed poorly to microplastics under laboratory conditions irrespective of weathering. Concentrations of Σ₄MCs (n = 9 sampling events) sorbed to microplastics ranged from 0.0 to 3.0 ng/g for weathered LDPE in filtered lake water, 0.2 to 4.6 ng/g for unweathered LDPE in filtered lake water, and 0.0 to 2.6 ng/g for unweathered LDPE in ultrapure water (Figure 4; see Figure S4 for concentrations standardized to surface area). The final mass of Σ₄MCs sorbed to weathered LDPE in filtered lake water, unweathered LDPE in filtered lake

Accepted Article
water, and unweathered LDPE in ultrapure water after 128 h was approximately 0.006%, 0.005%, and 0.003%, respectively, of the initial amount added.

Microcystins with higher polarity (Table 2) demonstrated a greater affinity for weathered LDPE microplastics than less polar ones (Figure S1; see Figure S5 for concentrations standardized to surface area). Sorption of MC-RR, MC-YR, and MC-LR by weathered LDPE in lake water, unweathered LDPE in lake water, and unweathered LDPE in ultrapure water increased from 1 to 4 h, after which point sorption capacity of weathered LDPE in lake water and unweathered LDPE in ultrapure water reached equilibrium. Unweathered LDPE in lake water, however, continued to increase. Sorption of MC-LA followed no apparent trend.

### Field-based sorption experiment (Exp. 3)

Surface water concentrations of Σ₁₂MCs in Dog Lake during the field experiment (n = 8 sampling events) ranged from 0.1 to 3.6 µg/L, with maximum concentrations observed in early October (Figure S7). On average, site 2 had concentrations of Σ₁₂MCs three times greater than site 1. MC-LA was the dominant congener, accounting for >88% and >53% of Σ₁₂MCs measured in surface waters of sites 1 and 2, respectively. MC-LR was detected at both sites, but MC-RR and MC-YR were detected at site 2 only.

In the field experiment, microcystins were detected in 100% of PET, 92% of LDPE, and 86% of PVC samples. Concentrations (ng/g) of sorbed Σ₁₂MCs on microplastics ranged from 0.2 to 84.1 for PET, 0.0 to 76.5 for LDPE, and 0.0 to 66.1 for PVC (Figure 5; see Figure S8 for concentrations standardized to surface area). On average, site 2 had concentrations of accumulated Σ₁₂MCs three times greater than site 1. The mean concentration of accumulated Σ₁₂MCs for PET was 14.3 ± 20.0 ng/g (47.7 ±
66.6 ng/m²), for LDPE was 18.7 ± 24.1 ng/g (9.4 ± 12.1 ng/m²), and for PVC was 14.4 ± 18.0 ng/g (4.2 ± 5.4 ng/m²). There was a significant difference in sorption capacity among polymer types at site 2 on only two sampling dates: LDPE microplastics accumulated significantly more microcystins than PET microcystins on both 10 October (F(2,6) = 7.182, p = 0.027) and 24 October (F(2,5) = 15.414, p = 0.009).

All microcystins measured in surface waters of Dog Lake were also measured on microplastic samples collected at the same time (Figure S9; see Figure S10 for concentrations standardized to surface area). Accumulated concentrations of microcystins on all polymers followed the trend: MC-LA > MC-LR > MC-RR > MC-YR. Of the microplastic samples collected at site 1 and 2 (n = 18 per site), MC-LA was detected in 86 and 96 % and MC-LR was detected in 29 and 94 %, respectively, and site 2 additionally detected MC-YR and MC-RR in 20 and 63% of samples, respectively.

DISCUSSION

Sorption capacity for microcystins differs among polymer types (H₁)

The present work demonstrates that the sorption capacity for microcystins differed among polymers in the laboratory experiment (Exp. 1) yet were largely the same in the field (Exp. 3). Under laboratory conditions, LDPE and PET consistently sorbed more microcystins than PP and PVC (Figure 2, Figure S2), and therefore, we accept the hypothesis that sorption capacity for microcystins differs among polymer types (H₁).

We anticipated differences in sorption capacity among the microplastics tested (Table 1) because sorption behaviour is partly driven by the distinct physicochemical properties of each polymer type. Crystallinity, the proportion of crystalline to amorphous regions in a polymer, may explain some of the responses seen in Exp. 1. Sorption occurs
predominately in amorphous regions, where greater sorption is typical in flexible rubbery
polymers because rigid glassy polymers are more condensed and cross-linked, thus
having lower permeability (Alimi et al., 2018). Some previous studies on waterborne
contaminants support this rule (Wang & Wang, 2018), but others do not (Li et al., 2018).
Given LDPE and PP are considered rubbery, and PET and PVC are considered glassy
(Alimi et al., 2018), crystallinity may therefore explain the enhanced sorption capacity of
LDPE and the reduced sorption capacity of PVC but neglects to explain the responses of
PP and PET.

Pestana et al. (2021) also found that some glassy polymers (polystyrene (PS) and PVC) had a greater sorption capacity for microcystins compared to their rubbery counterpart (PE). This exception to the rule is likely explained by the presence of other more dominant drivers of the interaction, such as some microplastics having a rougher surface, resulting in more sites for adsorption (Pestana et al., 2021). In our case, the poor response of PP may be better explained by the microplastic’s low specific surface area (Table 1), which can be an important factor in governing the sorption of hydrophilic contaminants to microplastics (Godoy et al., 2020; Liu et al., 2020). Enhanced sorption capacity of PET may also result from the polarity of the polymer, something seen by Moura et al. (2022) between PET and the more hydrophilic congener microcystin-tryptophan-arginine.

In contrast, sorption capacity was similar among the polymer types tested when
microplastics were exposed to microcystins in a eutrophic lake. Although LDPE
accumulated more microcystins on average in the field compared to PET and PVC, the
difference was considerably smaller than the response under laboratory conditions, and
the greatest accumulation within each sampling period alternated between each polymer type (6x LDPE, 3x PET, and 3x PVC). This discrepancy in findings between Exp. 1 and Exp. 3 is interesting, albeit somewhat expected, as the factors governing the interaction between microplastics and microcystins will differ between these two environments (see section 4.3 for further discussion).

### Affinity for plastic varies among microcystins (H₂)

Experiments 1 and 2 showed that affinity for plastic varies among microcystin congeners when exposed under laboratory conditions. Specifically, more polar microcystins (i.e., congeners with one or two arginine residues—denoted by “R” in the naming scheme—such as MC-RR, MC-YR, and MC-LR) demonstrated a greater affinity for plastic than less polar congeners (i.e., congeners with no arginine residues such as MC-LA) (Figure 3, Figure S3; Figure S1, Figure S5), and therefore, we accept the hypothesis that affinity for plastic varies among microcystins (H₂).

The chemical structure of the four congeners studied differed only in the amino acid residues present in the variable positions X² and Z⁴, which are known to govern differences in sorption behaviour among microcystins (Altaner et al., 2017). For instance, surface electrostatic interactions are a dominant mechanism in microplastic-contaminant sorption (Elizalde-Velázquez et al., 2020; Godoy et al., 2020; Guo et al., 2018) and likely explain the trend in affinity for plastic among the congeners tested. Microplastics commonly carry a net negative surface charge and are likely to attract positively charged species and repel negatively charged species (Godoy et al., 2019). Microcystins also primarily exist in anionic form but can contain both negatively and positively charged groups. For example, at neutral pH, MC-RR has two negatively charged groups and two
positively charged groups, MC-YR and MC-LR each have two negatively charged groups and a single positively charged group, and MC-LA has two negatively charged groups (Newcombe et al., 2003; see net charge at pH 7 in Table 2). Consequently, simultaneous surface electrostatic attraction and repulsion likely influence the microplastic-microcystin interaction; an idea supported by MC-RR's strong affinity for plastic compared to MC-LA's poor affinity for plastic. The pH of our laboratory experiments was 7.0 while the pH of our field experiment ranged between 7.2 and 9.2, indicating that for the duration of our experiments, MC-RR was net neutral, MC-YR and MC-LR carried a -1 charge, and MC-LA carried a -2 charge.

Other studies examining the microplastic-microcystin interaction observed the opposite trend, namely, as congener hydrophobicity increased, affinity for plastic also increased (Moura et al., 2022; Pestana et al., 2021). This suggests hydrophobic interactions are a leading force driving the sorption capacity of plastics for microcystins. However, given that microcystins are predominantly hydrophilic molecules, others have noted that hydrophobic interactions are unlikely to play a significant role in their sorption to hydrophobic phases in water and instead have suggested mechanisms including cation-bridging, hydrogen-bonding, and surface electrostatic interactions are more likely (Wu et al., 2011). The discrepancies in affinity for plastic among congeners between our study and others (Moura et al., 2022; Pestana et al., 2021) further bolster the notion that the interaction between microplastics and microcystins is a multifactor process that is context-dependent.

*Weathered microplastics have a greater sorption capacity for microcystins (H₃)*
The present work further demonstrates that although there is no apparent
difference in sorption capacity for microcystins between unweathered and weathered
LDPE when exposed under laboratory conditions (Exp. 2; Figure 4, Figure S4), polymers
accumulated more microcystins in the field. Therefore, we accept the hypothesis that
weathered microplastics have a greater sorption capacity for microcystins compared to
unweathered microplastics (H3). The latter result is likely due to biofilm formation on the
microplastics' surface when left to weather under ambient conditions in a eutrophic lake.

Biofilms can increase microplastic surface polarity, hydrophilicity, and specific
surface area, contributing to greater sorption capacity, especially for polar and
hydrophilic contaminants (Rummel et al., 2017) like microcystins. Biofilm formation
across all polymer types could explain why PVC demonstrated sorption capacity for
microcystins in the field but not in the laboratory. Further, the accumulation of metals
and pharmaceuticals on plastic can be mediated by a biofilm (Magadini et al., 2020;
Rochman et al., 2014). Thus, the sorption capacity for microcystins among different
polymers may be comparable in the field but noticeably different under laboratory
conditions because the biofilm, whose composition can be similar among polymer types
within a given location (Ye & Andrady, 1991; Zettler et al., 2013), is the dominant layer
for sorption. This hypothesis has recently been tested for one polymer type (PS) and a
single microcystin congener (MC-LR) under laboratory conditions (He et al., 2022). In
their study, these authors found PS had enhanced sorption capacity for MC-LR after
biofilm formation due to an alteration of the microplastic's surface properties,
specifically the addition of oxygen-containing functional groups increasing polar
interactions (He et al., 2022). Together, these findings have important implications for
studies that assess the interaction between microplastics and microcystin in the laboratory using pristine microplastics. As such, we recommend that future studies include the influence of biofilm formation on sorption capacity.

Additionally, the biofilm itself may contain toxin-producing cyanobacteria. A growing body of evidence demonstrates that microplastics interact with primary producers, including bloom-forming algae such as dinoflagellates and cyanobacteria (Kettner et al., 2019; Masó et al., 2003; Oberbeckmann et al., 2014; Yokota et al., 2017; Zettler et al., 2013). For example, Oberbeckmann et al. (2014) identified cyanobacteria as a dominant constituent of the plastisphere on PET drinking bottles submerged in the North Sea and PE, PP, and PS microplastics collected from the same area. Similarly, in a laboratory-based experiment, Yokota et al. (2017) exposed two microcystin-producing freshwater cyanobacteria, *Microcystis aeruginosa* and *Dolichospermum flos-aquae*, to microplastics and found colonial and filamentous cyanobacteria to be a dominant part of the plastisphere. For our study, it is probable that the microplastic-associated microcystins detected in the field included extracellular (i.e., waterborne) toxins sorbed from the ambient environment and intracellular (i.e., cell-bound) toxins from cyanobacteria colonizing the microplastic's surface. The latter additional, more indirect microplastic-microcystin interaction mechanism may also explain why we measured greater accumulated concentrations in the field (Exp. 3) than in the laboratory (Exp. 1 and 2) despite a considerably lower aqueous microcystin concentration in Dog Lake (Figure S7).
CONCLUSION

A comprehensive understanding of the combined effects of microplastic pollution and HABs in freshwater ecosystems is vital for assessing environmental risk. Recent studies on nanoplastics and microcystins offer insights into these potential synergies. For example, amino-modified PS nanoparticles can promote microcystin synthesis and extracellular release from *Microcystis aeruginosa* (Feng et al., 2020); nanoplastics can weaken inhibitory effects of hydrogen peroxide, a control measure for HABs (Guo et al., 2021); and prolonged exposure to PS nanoparticles can enhance MC-LR toxicity in the nematode *Caenorhabditis elegans* (Qu et al., 2019). On the latter concern regarding adverse impacts to aquatic life, organisms at every level of the food web have been shown to ingest or interact with microplastics (Bucci et al., 2020), and biofilm formation can increase their likelihood of doing so (Rummel et al., 2017), potentially increasing the possibility of microcystins causing a toxic insult. Still, the role of microplastic ingestion as a dominant pathway for contaminant transfer to biota is contested. Some researchers argue that the fraction of a given contaminant associated with microplastics is negligible compared with other natural media such as water, sediment, and food (Bakir et al., 2016; Koelmans et al., 2016). However, given that this idea is often discussed in reference to legacy contaminants, which waterborne microcystins are not, whether microplastics act as a vector for exposure to microcystins in biota should be investigated further.

Our findings also indicate that microplastics may influence the persistence and
facilitate the transport of microcystins in freshwater ecosystems. As stated previously,
unlike legacy contaminants, the concentration of waterborne microcystins is in constant
flux. Microcystins typically persist in the water for weeks to months after a bloom event,
contingent on natural detoxification routes such as dilution, sorption to sediment, thermal decomposition, photolysis, and biological degradation (Chorus and Welker, 2021).

Microplastics may aid in microcystin removal from the water column via accumulation, as was seen by He et al. (2022). However, microplastics may also make microcystins more persistent in surface waters if accumulated microcystins are more stable than waterborne microcystins or if microplastics carry microcystins to new areas within a waterbody with no or low aqueous concentrations. Again, these scenarios merit further investigation to determine the risk posed by both microplastics and microcystins when they are present in the same freshwater ecosystem.

*Supporting Information*—The Supporting information are available on the Wiley Online Library at DOI: 10.1002/etc.xxxx.

*Acknowledgment*—We thank K. and T. Schneider and the residents of Hideaway Lane for providing access to Buck Lake and Dog Lake, respectively. We also thank H. Vanderlip and A. Zborowski for their assistance in the lab and A. Tian, M. Dias, K. Brouwer, and J. Reynolds for their assistance in the field. This research was funded by the Natural Sciences and Engineering Council of Canada (Discovery Grant RGPIN-2020-06956 to D. M. Orihel) and in-kind contributions from the Ontario Ministry of the Environment, Conservation and Parks. E. K. Hataley was supported by the Natural Sciences and Engineering Council of Canada (Collaborative Research and Training Experience Program Grant 401208952 to R. S. Brown, Queen's University, D. M. Orihel, and others). Kal-Polymers (Mississauga, Canada) provided samples of recycled plastic polymers for this study.

*Disclaimer*—The authors declare no conflicts of interest.
*Data availability*—Data, associated metadata, and calculation tools are available from the
corresponding author (diane.orihel@queensu.ca).

*Author contributions statement*—Eden K. Hataley: Conceptualization; Methodology; Resources; Investigation; Formal analysis; Writing – original draft; Writing – review and editing; Visualization. René S. Shahmohamadloo: Investigation; Writing – original draft; Writing – review and editing. Xavier O. Almirall: Conceptualization; Methodology; Resources; Investigation; Formal analysis; Writing – original draft; Writing – review and editing; Supervision. Anna L. Harrison: Conceptualization; Formal analysis; Writing – review and editing. Chelsea M. Rochman: Conceptualization; Formal analysis; Writing – review and editing. Shan Zou: Investigation; Formal analysis; Writing – review and editing. Diane M. Orihel: Conceptualization; Methodology; Formal analysis; Writing – original draft; Writing – review and editing; Supervision; Project administration; Funding acquisition.

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Accepted Article
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FIGURE CAPTIONS

<figure><img src="image_5.png" /><figcaption>**Figure 1.** Fourier transform infrared spectra (A) of low-density polyethylene (LDPE), polyethylene terephthalate (PET), polyvinyl chloride (PVC), and polypropylene (PP). Characteristic peaks were identified and match respective polymer types (Table S1). Scanning electron microscopy images at 400x and 1600x magnification of LDPE (B and C), PET (D and E), PVC (F and G), and PP (H and I).</figcaption></figure>
<figure><img src="image_6.png" /></figure>
**Figure 2.** Sorption kinetics of total microcystins (Σ4MCs; the sum of microcystin-arginine-arginine (MC-RR), microcystin-tyrosine-arginine (MC-YR), microcystin-leucine-arginine (MC-LR), and microcystin-leucine-alanine (MC-LA)) by polyethylene terephthalate (PET), low-density polyethylene (LDPE), polyvinyl chloride (PVC), and polypropylene (PP) at t = 0, 1, 2, 4, 8, 16, 32, and 48 h. Each marker represents the mean sorbed concentration (ng/g) ± standard deviation (n = 6). Data from Experiment 1 (see Materials and Methods Section 2.1).
<figure><img src="image_7.png" /></figure>
**Figure 3.** Sorption kinetics of microcystin-arginine-arginine (MC-RR) (A), microcystin-tyrosine-arginine (MC-YR) (B), microcystin-leucine-arginine (MC-LR) (C), and microcystin-leucine-alanine (MC-LA) (D) by polyethylene terephthalate (PET), low-density polyethylene (LDPE), polyvinyl chloride (PVC), and polypropylene (PP) at t = 0, 1, 2, 4, 8, 16, 32, and 48 h. Each marker represents the mean sorbed concentration (ng/g) ± standard deviation (n = 6). Vertical axes differ among panels. Data from Experiment 1 (Materials and Methods Section 2.1).
<figure><img src="image_8.png" /></figure>

Weathered LDPE (LW)

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

Unweathered LDPE (LW)

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

Unweathered LDPE (UW)

<figure><img src="image_11.png" /></figure>
**Figure 4.** Sorption kinetics of total microcystins (Σ4MCs; the sum of microcystin-arginine-arginine (MC-RR), microcystin-tyrosine-arginine (MC-YR), microcystin-leucine-arginine (MC-LR), and microcystin-leucine-alanine (MC-LA)) by weathered low-density polyethylene (LDPE) in filtered lake water (LW), unweathered LDPE in filtered lake water (LW), and unweathered LDPE in ultrapure water (UW) at t = 0, 1, 2, 4, 8, 16, 32, 64, and 128 h. Each marker represents the mean sorbed concentration (ng/g) ± standard deviation (n = 6). Data from Experiment 2 (Materials and Methods Section 2.2).
<figure><img src="image_12.png" /></figure>
**Figure 5.** Temporal variation in concentration of total microcystins (Σ12MCs; the sum of MC-LR, MC-YR, MC-RR, MC-HtyR, MC-HilR, MC-WR, MC-LW, MC-LA, MC-LF, MC-LY, MC-Dha7-LR, and MC-Dha7-RR) accumulated by each polymer type (polyethylene terephthalate (PET), low-density polyethylene (LDPE), and polyvinyl chloride (PVC)) from 29 August to 24 October 2019 at sites 1 (A) and 2 (B) in Dog Lake, Ontario, Canada. Each bar represents the mean accumulated concentration (ng/g) ± standard deviation (n = 3). Vertical axes differ among panels. Lowercase letters represent significant differences across polymer types within locations at each sampling timepoint. Data from Experiment 3 (Materials and Methods Section 2.3).

## Tables

<table><caption>**Table 1.** Physical characterization of polymers used for Experiments 1 to 3. Weight and surface area shown as mean ± standard deviation (n = 3).</caption><thead><tr><th rowspan="2">Polymer<sup>a</sup></th><th rowspan="2">Shape</th><th rowspan="2">Largest Dimension<br>(mm)</th><th rowspan="2">Weight<br>(mg)</th><th>Specific Surface</th></tr><tr><th>Area<sup>b</sup><br>(m<sup>2</sup> g<sup>-1</sup>)</th></tr></thead><tbody><tr><td>Low-density polyethylene<br>(LDPE)</td><td>Disc-<br>shaped</td><td>3-5</td><td>18.7 ±<br>1.4</td><td>2.0 ± 1.1</td></tr><tr><td>Polyethylene terephthalate<br>(PET)</td><td>Fragmented</td><td>5</td><td>54.7 ±<br>7.4</td><td>0.3 ± 0.1</td></tr><tr><td>Polyvinyl chloride (PVC)</td><td>Cylindrical</td><td>3</td><td>18.5 ±<br>0.3</td><td>3.4 ± 0.3</td></tr><tr><td>Polypropylene (PP)</td><td>Cylindrical</td><td>3</td><td>25.1 ±<br>0.1</td><td>0.3 ± 0.1</td></tr></tbody></table>
ᵃpolymer as confirmed by Fourier transform infrared spectroscopy (see Figure 1); ᵇspecific surface area as measured by the Brunauer-Emmett-Teller (BET) method.

<table><caption>**Table 2.** Physical and chemical properties of microcystins used for Experiments 1 to 2.</caption><thead><tr><th>Congener</th><th>Name</th><th>Molecular Weight<sup>a</sup> (Da)</th><th>Net Charge, pH 7<sup>a</sup></th><th>log(D<sub>ow</sub>), pH 7<sup>b</sup></th><th>Hydrophobicity</th></tr></thead><tbody><tr><td>MC-RR</td><td>microcystin-arginine-arginine</td><td>1037</td><td>0</td><td>-0.72 ± 0.05</td><td>Least</td></tr><tr><td>MC-YR</td><td>microcystin-tyrosine-arginine</td><td>1044</td><td>-1</td><td>--</td><td>↓</td></tr><tr><td>MC-LR</td><td>microcystin-leucine-arginine</td><td>994</td><td>-1</td><td>-1.2 ± 0.2</td><td>Most</td></tr><tr><td>MC-LA</td><td>microcystin-leucine-alanine</td><td>909</td><td>-2</td><td>0.16 ± 0.05</td><td>Most</td></tr></tbody></table>

## References

Newcombe, G., Cook, D., Brooke, S., Ho, L., & Slyman, N. (2003). Treatment options for microcystin toxins: Similarities and differences between variants. *Environmental Technology*, 24(3), 299–308. https://doi.org/10.1080/09593330309385562

McCord, J., Lang, J. R., Hill, D., Strynar, M., Chernoff, N. (2018). pH dependent octanol-water partitioning coefficients of microcystin congeners. *Journal of Water Health*, 16(3), 340-345. https://doi.org/10.2166%2Fwh.2018.257

<footnote>ᵃNewcombe et al., 2003; ᵇMcCord et al., 2018.</footnote>