
==== Front
Eco Environ Health
Eco Environ Health
Eco-Environment & Health
2772-9850
Elsevier

S2772-9850(24)00045-0
10.1016/j.eehl.2024.05.004
Original Research Article
Assessing bioactivity of environmental water samples filtered using nanomembrane technology and mammalian cell lines
Morgan Sarah E. ad
DeLouise Lisa A. lisa_delouise@urmc.rochester.edu
abcd∗
a Department of Environmental Medicine, University of Rochester Medical Center, Rochester, NY 14642, USA
b Department of Biomedical Engineering, University of Rochester, Rochester, NY 14627, USA
c Department of Dermatology, University of Rochester Medical Center, Rochester, NY 14642, USA
d Lake Ontario Center for Microplastics and Human Health in a Changing Environment, University of Rochester Medical Center, Rochester, NY 14642, USA
∗ Corresponding author. lisa_delouise@urmc.rochester.edu
29 5 2024
9 2024
29 5 2024
3 3 347354
23 2 2024
10 5 2024
21 5 2024
© 2024 The Author(s)
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
This project reports on the use of a novel nanomembrane filtering technology to isolate and analyze the bioactivity of microplastic (MP)-containing debris from Lake Ontario water samples. Environmental MPs are a complex mixture of polymers and sorbed chemicals that are persistent and can exhibit a wide range of toxic effects. Since human exposure to MPs is unavoidable, it is necessary to characterize their bioactivity to assess potential health risks. This work seeks to quantify MP presence in the nearshore waters of Lake Ontario and begin to characterize the bioactivity of the filtrate containing MPs. We utilized silicon nitride (SiN) nanomembrane technology to isolate debris sized between 8 and 20 μm from lake water samples collected at various times and locations. MPs were identified with Nile red staining. Cell-based assays were conducted directly on the filtered debris to test for cell viability, aryl hydrocarbon receptor (AhR) activity, and interleukin 6 (IL-6) levels as a measure of proinflammatory response. All samples contained MPs. None of the isolated debris impacted cell viability. However, AhR activity and IL-6 levels varied over time. Additionally, no associations were observed between the amount of plastic and bioactivity. Observed differences in activity are likely due to variations in the physiochemical properties of debris between samples. Our results highlight the need for increased sampling to fully characterize the bioactivity of MPs in human cells and to elucidate the role that sample physiochemical and spatiotemporal properties play in this activity.

Graphical abstract

Image 1

Highlights

• Filtrate from Lake Ontario contains microplastics sized between 8 and 20 μm.

• Filtrate containing microplastics has variable bioactivity in immortalized human keratinocyte (HaCaT) cells.

• Sampling time and location impact filtrate and the included microplastics.

• SiN nanomembranes and vacuum filtration present a novel technique for isolating small microplastics.

Keywords:

Microplastics
SiN nanomembranes
Lake Ontario
Aryl hydrocarbon receptor (AhR) activity
IL-6 levels
Bioactivity
==== Body
pmc1 Introduction

Plastic pollution is a major problem worldwide, with over ten million tons entering the environment annually [1]. A large portion of this plastic pollution can be characterized as microplastics (MPs) or nanoplastics (NPs). MPs are small pieces of plastic that range in size from 1 μm to 5 mm [2,3]. There are two types of MPs: primary and secondary. Primary MPs are particles that are manufactured at the micro-size level and in pre-production pellets and microbeads in consumer products [4]. On the other hand, secondary MPs are produced by the degradation of larger plastic debris in the environment [5,6]. Once these particles fall below 1 μm in size, they are classified as NPs [7,8]. Although plastics degrade in the environment, they are estimated to persist for hundreds of years [9] and thus comprise a ubiquitous environmental contaminant. MPs have been detected on all seven continents [10] and in various geographical features including oceans [11], lakes [12], rivers [13], soil [14], and air [15]. Proximity to urban areas has been positively linked with an increased presence of plastic particles [16]. Researchers have detected both spatial and temporal variations within samples from the same water source [17,18]. Temporal trends varied as some studies found no clear trend [19], whereas others observed an increase in MPs over time [20]. Investigating temporal trends through a seasonal lens may not provide sufficient information for establishing conclusive trends. Depending on the study, MP contamination is higher in either the wet [21] or the dry [22] seasons. MPs are widespread but are differentially impacted by various environmental factors.

In addition to being a ubiquitous environmental contaminant, MPs have been found in various foodstuffs, including honey [23], sugar [23], bottled water [24,25], and milk [26]. Given the persistent nature and ubiquity of plastic, human exposure to MPs is inevitable. Therefore, it is important to characterize the risks to human health associated with this exposure. This is especially challenging, given that a key characteristic of environmental MPs is their high degree of heterogeneity due to a wide range of polymers, adsorbed pollutants, morphologies, and dimensions present [27]. The sorbed chemicals include both those added during production to alter the plastic's properties to suit their post-production use [28] and contaminants picked up after plastic enters the environment [29]. Contaminants that have been identified in MP samples include plasticizers such as bisphenol A (BPA), an endocrine-disrupting chemical (EDC), polychlorinated biphenyls [30,31], aryl hydrocarbon receptor (AhR) agonists [32], and heavy metals [33]. Due to the widespread presence of MPs and the vast pool of AhR and EDC ligands, it is highly likely that most environmental MPs contain these chemical classes. The AhR is a promiscuous receptor that is considered to be a xenobiotic sensor [34]. It is of particular interest since exogenous activation is associated with disruptions to homeostasis and immunity and with increased cancer susceptibility [35]. Additionally, MPs may act similarly to other particulate matter exposures to induce inflammation. One broad measure of inflammation in cells is interleukin 6 (IL-6) levels [36]. Previous studies have investigated these endpoints in pure MPs [[37], [38], [39]]. Therefore, AhR activity and IL-6 concentration are putative measures of bioactivity worth studying in environmental samples.

This study seeks to investigate the bioactivity of environmental debris sourced from the nearshore waters of Lake Ontario (LO). LO is the largest body of water in Western New York. The LO water basin serves approximately 9 million people between the United States and Canada [40]. Residents rely on the lake for freshwater access, including drinking water, fishing, and recreational activities such as swimming and boating. Different areas of the LO shoreline are used for different activities depending on factors such as the availability of sandy beachfront access, surface current, and pollutant levels related to proximity to rivers and tributaries. Potential seasonal trends are of interest in relation to LO, as there is an increase in water entering the lake following snow thawing in the spring that could alter the MPs in the lake. Additionally, lake usage by residents is season-dependent; therefore, understanding whether exposure risk and/or hazard varies between seasons would be important for the development of guidelines for minimizing exposure impact. One common limitation with environmental samples is that it is difficult to isolate particles at the lower end of the MP size range [41] and nearly impossible to isolate NPs [42]. To overcome this limitation, we utilized commercial nanomembrane filters to isolate particles sized between 8 and 20 μm. The nanomembranes have additional benefits, including being biologically inert, optically clear, plastic-free, resistant to clogging, and having a faster flow rate than other filters [43,44]. In addition to the novelty of the size range, smaller MPs were targeted because laboratory studies have shown smaller particles are more toxic [45,46].

In this study, water samples were collected from four lakeshore locations at up to four timepoints across multiple years. Environmental debris was isolated using nanomembrane filters, and plastic presence was confirmed via Nile red staining. Bioactivity was measured using assays for AhR activity and IL-6 levels. Our results suggest that the innovative method for isolating smaller MPs from water samples described has applications for future samples while highlighting the need for a better understanding of the impacts of sample physical/chemical, spatial, and temporal properties on MP bioactivity and the risk these particles pose to human health. Developing methods to sample plastics in the nanoparticle size range is also critical for assessing human health impacts.

2 Methods

2.1 Sample collection

Up to five 1-L water samples were collected from the nearshore waters of LO (approximately 6 inches beneath the surface of water, approximately 2 feet deep) from up to four locations (Fig. 1). Samples were collected from Ontario Beach Park (OBP) (location 2) on November 10, 2021, July 19, 2022, February 17, 2023, and April 21, 2023. On April 21, 2023, samples were also collected from Hamlin Beach (HB) (location 1), Durand Eastman Beach (DEB) (location 3), and Webster Park (WP) (location 4). Locations were selected due to the variety of uses of the beaches and surrounding land. Visitors are attracted to Hamlin Beach State Park for its clear water and sandy beaches [47]. Similarly, OBP has a beach and water access, but the water is less attractive due to the proximity of the mouth of the polluted Genesee River, so there is less swimming, but the proximity to the city of Rochester translates to plenty of boating [48]. DEB is similarly near the city, but due to strong currents, there are limited swimming hours [49]. Lastly, WP has a rocky beach and pier, so activities there mainly include fishing [50]. Clean 1-L glass bottles were rinsed three times in lake water before being filled and capped underwater. The bactericide sodium azide (0.05%) was added to the samples to prevent growth. Prior to collection, bottles were scrubbed with soap, rinsed three times with tap water, rinsed three times with deionized (DI) water, and sprayed with 70% ethanol.Fig. 1 Map of sampling locations. Water samples were collected from Lake Ontario at (1) Hamlin Beach State Park (HB), (2) Ontario Beach Park (OBP), (3) Durand Eastman Beach (DEB), and (4) Webster Park (WP). Created with Proxi.

Fig. 1

2.2 Filtering

Water samples were filtered using previously established protocols [43]. Briefly, water samples were thoroughly mixed to suspend any precipitated particulates before an aliquot was measured. The aliquot was first passed through a 20-μm metal sieve (Artesian Systems) before being filtered through a 400-nm-thick SiN Nanomembrane filter (SiMPore MSSN400-3L-8.0) in a vacuum filtration set-up (Fig. 2). Each filter was comprised of 3 windows (0.7 × 3 mm). In each window, there were 730 slits. Each slit measured 8 × 50 μm. This set-up allows for the isolation of an infrequently studied size-fraction (8–20 μm). The design of these membranes paired with applied vacuum allowed large volumes of the collected water samples to be isolated efficiently, with 250 mL taking roughly 15 min to filter. Furthermore, these membranes are optically clear and made of non-plastic biocompatible materials, allowing assays to be performed directly on them. All aliquots filtered were of 250-mL volume. Select filters were stained with Nile red [51] and trypan blue for plastics and cellulosic materials, respectively. After each stain, these filters were rinsed with 2.25 mL. There was no additional processing for filters used for cell culture experiments. Blank filters were run through the aforementioned method as process controls. Prior to filtering, all glassware was scrubbed with soap, rinsed three times with tap water, rinsed three times with DI water, and sprayed with 70% ethanol.Fig. 2 Experimental set-up. (a) Lake water was collected in 1-L glass bottles and treated with sodium azide before filtering through a 20-μm sieve and an 8-μm SiN nanomembrane. (b) HaCaT cells were exposed to debris for 24 h before cellular components were allocated for various endpoints. Created with Biorender.com under institutional site license, #UU26FRGOHN. HaCaT, immortalized human keratinocyte.

Fig. 2

2.3 Imaging

Bright field and fluorescence images of the filtered debris were captured using an epifluorescence microscope. Nile red and trypan blue fluorescence were imaged using the Texas Red (ex. 586 nm/em. 647 nm) and 4′,6-diamidino-2-phenylindole (DAPI, ex. 377 nm/em. 447 nm) filters, respectively. Nile red–positive area was quantified using ImageJ. Briefly, each filter (3 slots per filter) was marked as a region of interest (ROI) using the bright field image. ROIs were then applied to the Nile red image; each ROI was threshold-adjusted, and the percent of pixels above the threshold (percent Nile red) was measured.

2.4 Cell culture

Immortalized human keratinocytes (HaCaTs) were cultured in DMEM supplemented with 10% fetal bovine serum (FBS), and 1% Penicillin Streptomycin and incubated in a humidified incubator at 37 °C with 5% CO2. Cells were seeded at 1.2 × 105 cells/well on top of SiN nanomembranes placed in 48 well plates. Following 24 h of exposure, supernatant was collected, cell viability was measured in plate, and RNA was isolated.

2.5 Cell viability

After 24 h of exposure, cell culture media was replaced with a 10% presto blue (Invitrogen P50200) solution in the media. Plates were incubated for 30 min at 37 °C before fluorescence was measured (ex. 560 nm/em. 590 nm). Fluorescence values were background-corrected before the percentage of live cells was calculated relative to the media controls. Background control was reagents without cells, and media controls were cells exposed to media only. Blank filters were used to confirm that the results were not due to the SiN nanomembranes.

2.6 AhR activation via real-time reverse transcription quantitative polymerase chain reaction (RT-qPCR)

RNA was isolated using the E.Z.N.A Total RNA Kit I (Omega Bio-Tek R6834) according to the manufacturer's instructions before its concentration was measured using the Nanodrop method. RNA was converted to complementary DNA (cDNA) using the iScript cDNA Synthesis Kit (Bio-Rad 1708890). The resulting cDNA was used to determine the messenger RNA (mRNA) expression of CYP1A1 and CYP1B1 relative to the hypoxanthine phosphoribosyltransferase (HPRT) housekeeping gene and media controls using the PowerUp SYBR Green Master Mix (Applied Biosystems A25742) and the ΔΔCt method. HPRT was used as the housekeeping gene, and media controls were cells exposed to media only. Blank filters were used to confirm that the results were not due to the SiN nanomembranes. Primer sequences for RT-qPCR are listed in Table 1.Table 1 Primer sequences [52].

Table 1Gene	Forward sequence	Reverse sequence	
HPRT	5′-TGCTGAGGATTTGGAAAGGG-3′	5′-ACAGAGGGCTACAATGTGATG-3′	
CYP1A1	5′-TAGACACTGATCTGGCTGCAG-3′	5′-GGGAAGGCTCCATCAGCATC-3′	
CYP1B1	5′-CATGCGCTTCTCCAGCTTTGT-3′	5′-GGCCACTTCACTGGGTCATGA-3′	

2.7 IL-6 Enzyme-linked Immunosorbent Assay (ELISA)

IL-6 concentration was measured in supernatant collected from HaCaTs exposed to filtered debris for 24 h using a human IL-6 uncoated ELISA Kit (Invitrogen 88-7066) in accordance with the manufacturer's instructions. Briefly, on day 1, an ELISA plate (Corning 9018) was coated with a capture antibody and incubated overnight. On day 2, the plate was washed, blocked with diluted ELISA/ELISPOT for 1 h at room temperature, washed again, and 100 μL of IL-6 standard or sample supernatant was added before overnight incubation at 2 °C. The following day (day 3), the plate was washed, incubated with detection antibody for 1 h at room temperature, washed, incubated with streptavidin-horseradish peroxidase (HRP) for 30 min at room temperature, washed, and incubated with tetramethylbenzidine (TMB) for 15 min at room temperature. The reaction was stopped with 0.5 M HCl, and absorbance was measured at 450 nm and 570 nm. Absorbance at 570 nm was used as background correction, and IL-6 sample concentrations were calculated relative to media controls. Media controls were cells exposed to media only. Blank filters were used to confirm that the results were not due to the SiN nanomembranes.

2.8 Statistics

All statistical analyses were performed using GraphPad Prism 10. One-way analyses of variance (ANOVAs) and linear regression tests were performed as needed. Tukey's multiple comparison test was used to determine the significance between exposures following a significant (p < 0.05) ANOVA result. An asterisk (∗) was used to indicate that the two exposures were significantly different. Exposures were tested with n = 3, and each experimental replicate was from a separate 1-L sample collected from LO.

3 Results

3.1 MPs are identified in the filtrate of Lake Ontario samples year-round

Due to the ubiquity of MPs, it is likely they are present in LO, and it is important to assess plastic levels before determining the impacts of exposure to these particles. To accomplish this, debris from LO water samples between 8 and 20 μm were filtered onto SiN nanomembranes and stained with Nile red and trypan blue. Nile red stains lipophilic particles [51], whereas trypan blue stains cellulosic materials. Composite and Nile red channel images of representative filters are shown in Fig. 3a. All samples filtered contained Nile red–positive particles. The percent Nile red–positive area in the filter windows (3 per filter) was measured using threshold analysis in ImageJ (Fig. 3b and c). Nile red–positive area did not significantly change over time in samples from OBP (Fig. 3b) or between sampling locations in April 2023 (Fig. 3c). However, there was some variation in the amount of debris collected per filter between timepoints and sampling locations (Fig. 3b and c). All samples contained detectable levels of Nile red–positive plastics, although the amount of plastic present in the 8–20 μm range did not vary between samples.Fig. 3 Lake Ontario debris contains plastic particles. (a) Representative bright field and fluorescence microscopy images of debris isolated from Lake Ontario water samples stained with Nile red (red-top, black-bottom) and trypan blue (blue) and quantification of the percent positive Nile red area per filter to show (b) temporal and (c) spatial trends; n = 1–3.

Fig. 3

3.2 Filtered debris has no impact on cell viability

Toxicity testing often investigates impacts on mortality at the organism level or viability at the cellular level. A PrestoBlue assay was selected as a proxy for cell viability as it measures alterations in metabolic activity through differences in oxidizable nicotinamide adenine dinucleotide (NADH). HaCaTs were cultured directly on the filtered debris for 24 h before cell viability was assessed. Samples taken at various times from OBP (Fig. 4a) and from various beaches in April 2023 (Fig. 4b) did not exhibit differences in metabolic activity. Overall, none of the filtered samples significantly altered cell viability.Fig. 4 HaCaTs exposed to filtered debris exhibit no alterations in cell viability. HaCaT cells were exposed to Lake Ontario debris for 24 h. Metabolic activity data are organized to show (a) temporal and (b) spatial trends using samples collected from Ontario Beach Park and in April 2023, respectively. Analyzed by one-way ANOVA followed by Tukey's multiple comparisons test; n = 3. ANOVA, analysis of variance; HaCaT, immortalized human keratinocyte.

Fig. 4

3.3 Debris AhR activity changes over time but not between sampling locations

The AhR has a role in homeostasis, immunity, and cancer, and therefore, exposures that alter AhR activity have implications for human health. CYP1B1 and CYP1A1 mRNA expression were measured via RT-qPCR as a proxy for AhR activation [53] using RNA isolated from HaCaTs cultured on the filtered debris. CYP1B1 expression (Fig. 5) showed statistically significant variation between samples. CYP1A1 expression showed similar trends (Fig. S1). The AhR activity of the filtered debris varied over time (p = 0.0104; Fig. 5a). No spatial variations in AhR activity were observed (Fig. 5b). When AhR activity was plotted as a function of Nile red–positive area, no association was observed (p = 0.1960, r2 = 0.01021; Fig. 5c). AhR activity decreased over time but did not change with sampling location.Fig. 5 Exposure to select filtered debris activates the AhR in HaCaT cells. AhR activity was measured through CYP1B1 mRNA levels in HaCaT cells exposed to Lake Ontario debris for 24 h. AhR activity is organized to show (a) temporal and (b) spatial trends using samples collected from Ontario Beach Park and in April 2023, respectively. (c) AhR activity was analyzed as a function of the Nile red–positive area. Analyzed by one-way ANOVA followed by Tukey's multiple comparisons test and/or linear regression; n = 3; ∗∗∗p < 0.001, ∗∗p < 0.01. AhR, aryl hydrocarbon receptor.

Fig. 5

3.4 Debris IL-6 levels change over time and between sampling locations

Particle exposure often results in inflammation, so MP exposure may induce inflammation as well. IL-6 is a common measure of innate inflammation. The supernatant from HaCaTs exposed to filtered debris was used for an IL-6 ELISA. Elevated IL-6 levels were detected at different times at OBP between November 2021 and November 2023 (p = 0.0065; Fig. 6c). Additionally, IL-6 levels were significantly increased relative to blank filters in samples exposed to debris from Hamlin Beach in April 2023 (p = 0.0008; Fig. 6a) and OBP in November 2023 (p = 0.0471; Fig. 6b). No significant spatial variations were observed between samples collected in April 2023 (Fig. 6b). IL-6 levels were found to have no association with Nile red–positive area (p = 0.3888, r2 = 0.03395; Fig. 6c). IL-6 levels varied with time and location sampled.Fig. 6 Exposure to select filtered debris alters IL-6 concentration in HaCaT cells. IL-6 levels were measured by ELISA in supernatant from HaCaT cells exposed to Lake Ontario debris for 24 h. IL-6 levels are organized to show (a) temporal and (b) spatial trends using samples collected from Ontario Beach Park and in April 2023. (c) IL-6 levels were analyzed as a function of the Nile red–positive area. Analyzed by one-way ANOVA followed by Tukey's multiple comparisons test and/or linear regression; n = 3; ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Fig. 6

4 Discussion

Plastic particles were present in all samples and ranged from 0.19% to 0.65% of the filter area (Fig. 3). Percent area was selected over particle enumeration due to the high variability in particle size and the presence of particle agglomeration. Due to the quantification method used, it is difficult to compare concentrations between this study and others. Previous studies have found concentrations of 0.8 particles/L [16] and 30.6 pellets/m2 [54]. While these concentrations are likely lower, the sampling methods used differed, and the isolated size fractions were much larger (>125 μm and 63–5,600 μm, respectively). The lack of a trend in plastic amount over time is not wholly unexpected as the area surrounding OBP did not experience any major changes in usage or any spillage events. However, differences between the more urban locations, OBP and DEB, and the more rural beaches, HB and WP, were anticipated based on differences in pollution generated by land and water usage in the surrounding areas. This may be explained by the facts that the two furthest beaches, HB and WP, are only about 30 miles apart, the nearshore waters of LO near Rochester exhibit strong currents that are largely influenced by wind [55,56], and models of the Laurentian Great Lakes, of which LO is one, have shown that lake plastic disperses to the sediment and shore [24,57]. It is also of interest to note that the currents [58] and waves [59] along the shore of LO fluctuate between east and west, and thus contaminants can disperse in the near-shore waters in not only the prevailing easterly direction but also westerly directions. Additionally, the lack of observed differences in plastic amount may be due to variations in the size distribution of the plastics present in the different samples that could not be captured since only one size fraction was isolated. On the other hand, variations in the amount of debris collected between samples may be due to samples being collected at different points in the tide cycle and different topographical features between beaches that varied the amount of sediment suspended in the lake during sampling. Overall, samples taken from LO contain plastics, but plastic concentration was not found to vary between samples, pointing to the need to conduct more frequent sampling.

None of the filtered samples induced significant alterations to HaCaT metabolic activity and are therefore assumed not to be cytotoxic (Fig. 4). This may be due to the isolated size fraction being too large to be internalized by the exposed cells [60]. Despite the lack of observed cytotoxicity, the filtered debris still had the potential to impact other components of cellular homeostasis. Studies using pristine plastic particles have observed alterations to cellular activity without impacts on cell viability [61]. Additionally, the lack of cell death following exposure to filtered debris indicates that changes observed in other measured endpoints are real and not simply an artifact of decreased cell numbers.

AhR activation is of interest as the AhR has roles in cellular homeostasis, immunity, cancer, and development. Since it is a promiscuous receptor with a wide range of ligands, it is frequently used as a xenobiotic sensor. The AhR is a cytosolic receptor that translocates to the nucleus following activation, where select genes are upregulated. Two of the genes that are strongly upregulated following AhR activation in keratinocytes are CYP1B1 and CYP1A1 [62]. The AhR exhibits ligand-specific gene expression [63], so it is not unexpected that significant alterations in expression were observed with CYP1B1 expression (Fig. 5) but not CYP1A1 expression (Fig. S1). Somewhat unexpectedly, AhR activity in the filtered debris depends on the sampling time between November 2021 and November 2023 (Fig. 5a). This may indicate changes to the chemical composition of the pollutants present on the debris and in the water. Similarly, variations in AhR activity between the sampling locations were anticipated due to differences in the rurality of the surrounding locations. Chemical runoff between urban and rural areas does differ [64,65]. Given the wide range of identified AhR ligands, it is plasuible that both types of runoffs could exhibit AhR acitivity in the filtered retentate. However, no AhR activity was observed in April 2023 in samples taken at differnet locations (Fig. 5b), and a decrease in AhR activity was observed in OBP samples taken between November 2021 and November 2023. Clearly, more sampling is needed to determine if AhR activity is a good biomarker of MP presence and/or lake-water health. Another factor that likely has a role in the AhR activity observed is the fact that the AhR is a cytosolic receptor, and since the filtered debris is likely too large for cellular internalization, AhR activation may go undetected. Similarly, the lack of association between plastic amount and AhR activity (Fig. 5c) is also likely due to variations in the chemical composition of the pollutants present on the debris and in the water due to the vast number and variety of chemicals that act as AhR ligands. Future studies that include analysis of the chemical composition of the filtered debris, in addition to increased sampling frequency, would help investigate this. Additionally, changes in mRNA levels are insufficient on their own to demonstrate receptor activation in the long run; however, following AhR activation by 2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD), a common AhR agonist, both CYP1B1 mRNA and protein levels have been shown to increase together [66,67]. To confirm activation, future work will also include an assessment of CYP enzyme activity and protein levels following debris exposure. Exposure to MPs may result in perturbations to AhR signaling through MPs acting as a vector for AhR ligands.

IL-6 is a common marker for general inflammation, which has previously been linked to particle exposure, and MPs are expected to behave similarly to nanoparticles. Although we observed a lack of activity in November 2021, we did observe an increase in IL-6 levels in samples collected at other times, with significantly elevated levels in November 2023 (Fig. 6a). Although not significant, the more rural locations, HB and WP, did have slightly elevated IL-6 levels compared to the more urban locations, OBP and DEB (Fig. 6b). This is likely due to differences in the composition of runoff commonly found between urban and rural locations [24,56], which impact the chemical composition of the pollutants found on the MPs isolated from different locations. Similar to AhR signaling, the lack of association between plastic and IL-6 levels is likely due to differences in the chemical profile of the MPs present (Fig. 6c), although additional sampling is needed to establish an association. The observed alterations in IL-6 levels suggest MP exposure may alter the pro-inflammatory/anti-inflammatory balance.

5 Conclusion

MPs are pervasive and persistent pollutants that humans are inevitably exposed to. Therefore, it is imperative that the risk of exposure to these particles be determined. This study utilized nanomembrane filter technology to investigate the presence and bioactivity of filtered debris containing MPs isolated from the nearshore waters of LO. All samples contained plastics, but the filtered debris did not alter cell viability. Some samples induced significant alterations in AhR activity and/or IL-6 levels. It is important to note that increased CYP1B1 mRNA has been interpreted to indicate AhR activity; however, without protein-level analysis, this cannot be confirmed, and future work should incorporate protein analysis. Temporal trends were observed, but spatial ones were not observed on the specific day the samples were collected. These variations could not be explained by differences in the amount of plastic present, suggesting expanded sampling is needed to fully characterize the impacts of physiochemical and spatiotemporal properties on plastic activity. Importantly, this study demonstrates a novel methodology for isolating and characterizing the bioactivity of debris between 8 and 20 μm, an understudied size fraction, filtered from LO that could be adapted to other waterways. In conclusion, filtered debris from LO contains plastics and is not biologically inert. This preliminary study reports on novel methodology and technology that can be used to analyze additional water samples and highlights the need for increased sampling to determine any seasonal and temporal effects. Ultimately, the results of this study and future applications of this methodology will aid in understanding the hazard and risk MP exposure poses to humans.

CRediT authorship contribution statement

S.M.: methodology, investigation, writing - original draft, writing - review & editing, visualization. L.D.: conceptualization, writing - review & editing, funding acquisition, supervision.

Declaration of competing interest

The authors declare that they have no known conflicts of interest.

Appendix ASupplementary data

The following is the Supplementary data to this article:Multimedia component 1

Multimedia component 1

Acknowledgments

We thank Dr. Samantha Romanick, PhD (University of Rochester) for sharing her knowledge on filtering using SiN nanomembranes and Dr. James McGrath, PhD (University of Rochester) for the use of his laboratory space for filtering experiments. This work was supported by the 10.13039/100000066 National Institute of Environmental Health Sciences (NIEHS R01 ES021492 ), the University of Rochester Toxicology Training Program (NIEHS T32 ES007026 ), and the University of Rochester Environmental Health Sciences Center (NIEHS P30 ES001247 ).

Appendix A Supplementary data to this article can be found online at https://doi.org/10.1016/j.eehl.2024.05.004.
==== Refs
References

1 International Union for Conservation of Nature Marine plastic pollution https://www.iucn.org/resources/issues-brief/marine-plastic-pollution
2 Thompson R.C. Olsen Y. Mitchell R.P. Davis A. Rowland S.J. John A.W.G. McGonigle D. Russell A.E. Lost at sea: where is all the plastic? Science 304 2004 838 10.1126/science.1094559 15131299
3 Moore C.J. Synthetic polymers in the marine environment: a rapidly increasing, long-term threat Environ. Res. 108 2008 131 139 10.1016/j.envres.2008.07.025 18949831
4 Germanov E.S. Marshall A.D. Bejder L. Fossi M.C. Loneragan N.R. Microplastics: no small problem for filter-feeding megafauna Trends Ecol. Evol. 33 2018 227 232 10.1016/j.tree.2018.01.005 29422348
5 Andrady A. Microplastics in the marine environment Mar. Pollut. Bull. 62 2011 1596 1605 10.1016/j.marpolbul.2011.05.030 21742351
6 Song Y.K. Hong S.H. Jang M. Han G.M. Jung S.W. Shim W.J. Combined effects of UV exposure duration and mechanical abrasion on microplastic fragmentation by polymer type Environ. Sci. Technol. 51 2017 4368 4376 10.1021/acs.est.6b06155 28249388
7 Gigault J. Pedrono B. Maxit B. Halle A.T. Marine plastic litter: the unanalyzed nano-fraction Environ. Sci. Nano 3 2016 346 350 10.1039/C6EN00008H
8 Ter Halle A. Jeanneau L. Martignac M. Jardé E. Pedrono B. Brach L. Gigault J. Nanoplastic in the North Atlantic subtropical gyre Environ. Sci. Technol. 51 2017 13689 13697 10.1021/acs.est.7b03667 29161030
9 Turner A. Arnold R. Williams T. Weathering and persistence of plastic in the marine environment: lessons from LEGO Environ. Pollut. 262 2020 114299 10.1016/j.envpol.2020.114299
10 Barnes D.K.A. Galgani F. Thompson R.C. Barlaz M. Accumulation and fragmentation of plastic debris in global environments Philos. Trans. R. Soc. B Biol. Sci. 364 2009 1985 1998 10.1098/rstb.2008.0205
11 Noren F. Small Plastic Particles in Coastal Swedish Waters 2007 KIMO and N-Research Sweden http://www.n-research.se/pdf/Small%20plastic%20particles%20in%20Swedish%20West%20Coast%20Waters.pdf
12 Eriksen M. Mason S. Wilson S. Box C. Zellers A. Edwards W. Farley H. Amato S. Microplastic pollution in the surface waters of the Laurentian Great Lakes Mar. Pollut. Bull. 77 2013 177 182 10.1016/j.marpolbul.2013.10.007 24449922
13 Hurley R. Woodward J. Rothwell J.J. Microplastic contamination of river beds significantly reduced by catchment-wide flooding Nat. Geosci. 11 2018 251 257 10.1038/s41561-018-0080-1
14 Zhang S. Yang X. Gertsen H. Peters P. Salánki T. Geissen V. A simple method for the extraction and identification of light density microplastics from soil Sci. Total Environ. 616–617 2018 1056 1065 10.1016/j.scitotenv.2017.10.213
15 Ageel H.K. Harrad S. Abdallah M.A.-E. Occurrence, human exposure, and risk of microplastics in the indoor environment Environ. Sci. Process. Impacts 24 2022 17 31 10.1039/D1EM00301A 34842877
16 Grbić J. Helm P. Athey S. Rochman C.M. Microplastics entering northwestern Lake Ontario are diverse and linked to urban sources Water Res. 174 2020 115623 10.1016/j.watres.2020.115623
17 Walters L.J. Craig C.A. Dark E. Wayles J. Encomio V. Coldren G. Sailor-Tynes T. Fox D.W. Quantifying spatial and temporal trends of microplastic pollution in surface water and in the eastern oyster Crassostrea virginica for a dynamic Florida Estuary Environments 9 2022 131 10.3390/environments9100131
18 Bakir A. Doran D. Silburn B. Russell J. Archer-Rand S. Barry J. Maes T. Limpenny C. A spatial and temporal assessment of microplastics in seafloor sediments: a case study for the UK Front. Mar. Sci. 9 2023 1093815 https://www.frontiersin.org/articles/10.3389/fmars.2022.1093815
19 Eryaşar A.R. Gedik K. Şahin A. Öztürk R.Ç. Yılmaz F. Characteristics and temporal trends of microplastics in the coastal area in the Southern Black Sea over the past decade Mar. Pollut. Bull. 173 2021 112993 10.1016/j.marpolbul.2021.112993
20 Li J. Huang W. Xu Y. Jin A. Zhang D. Zhang C. Microplastics in sediment cores as indicators of temporal trends in microplastic pollution in Andong salt marsh, Hangzhou Bay, China Reg. Stud. Mar. Sci. 35 2020 101149 10.1016/j.rsma.2020.101149
21 Warrier A.K. Kulkarni B. Amrutha K. Jayaram D. Valsan G. Agarwal P. Seasonal variations in the abundance and distribution of microplastic particles in the surface waters of a Southern Indian Lake Chemosphere 300 2022 134556 10.1016/j.chemosphere.2022.134556
22 Wang G. Lu J. Li W. Ning J. Zhou L. Tong Y. Liu Z. Zhou H. Seasonal variation and risk assessment of microplastics in surface water of the Manas River Basin, China Ecotoxicol. Environ. Saf. 208 2021 111477 10.1016/j.ecoenv.2020.111477
23 Liebezeit G. Liebezeit E. Non-pollen particulates in honey and sugar Food Addit. Contam. A 30 2013 2136 2140 10.1080/19440049.2013.843025
24 Mason S.A. Welch V.G. Neratko J. Synthetic polymer contamination in bottled water Front. Chem. 6 2018 407 10.3389/fchem.2018.00407 30255015
25 Li H. Zhu L. Ma M. Wu H. An L. Yang Z. Occurrence of microplastics in commercially sold bottled water Sci. Total Environ. 867 2023 161553 10.1016/j.scitotenv.2023.161553
26 Kutralam-Muniasamy G. Pérez-Guevara F. Elizalde-Martínez I. Shruti V.C. Branded milks – are they immune from microplastics contamination? Sci. Total Environ. 714 2020 136823 10.1016/j.scitotenv.2020.136823
27 Lambert S. Scherer C. Wagner M. Ecotoxicity testing of microplastics: considering the heterogeneity of physicochemical properties Integr. Environ. Assess. Manag. 13 2017 470 475 10.1002/ieam.1901 28440923
28 Hansen E. Nilsson N. Lithner D. Lassen C. Hazardous Substances in Plastic Materials 2013 COWI and the Danish Technological Institute
29 Wang F. Wong C.S. Chen D. Lu X. Wang F. Zeng E.Y. Interaction of toxic chemicals with microplastics: a critical review Water Res. 139 2018 208 219 10.1016/j.watres.2018.04.003 29653356
30 Mato Y. Isobe T. Takada H. Kanehiro H. Ohtake C. Kaminuma T. Plastic resin pellets as a transport medium for toxic chemicals in the marine environment Environ. Sci. Technol. 35 2001 318 324 10.1021/es0010498 11347604
31 Rochman C.M. Hoh E. Hentschel B.T. Kaye S. Long-term field measurement of sorption of organic contaminants to five types of plastic pellets: implications for plastic marine debris Environ. Sci. Technol. 47 2013 1646 1654 10.1021/es303700s 23270427
32 Rummel C.D. Escher B.I. Sandblom O. Plassmann M.M. Arp H.P.H. MacLeod M. Jahnke A. Effects of leachates from UV-weathered microplastic in cell-based bioassays Environ. Sci. Technol. 53 2019 9214 9223 10.1021/acs.est.9b02400 31257880
33 Khalid N. Aqeel M. Noman A. Khan S.M. Akhter N. Interactions and effects of microplastics with heavy metals in aquatic and terrestrial environments Environ. Pollut. 290 2021 118104 10.1016/j.envpol.2021.118104
34 Kim H.-M. Kim J.-W. Choi Y. Chun H.-S. Im I. Han Y.-M. Song C.-W. Yoon S. Xeno-sensing activity of the aryl hydrocarbon receptor in human pluripotent stem cell-derived hepatocyte-like cells Sci. Rep. 6 2016 21684 10.1038/srep21684
35 Kawajiri K. Fujii-Kuriyama Y. The aryl hydrocarbon receptor: a multifunctional chemical sensor for host defense and homeostatic maintenance Exp. Anim. 66 2017 75 89 10.1538/expanim.16-0092 27980293
36 Tanaka T. Narazaki M. Kishimoto T. IL-6 in inflammation, immunity, and disease Cold Spring Harb. Perspect. Biol. 6 2014 a016295 10.1101/cshperspect.a016295 25190079
37 Fournier E. Leveque M. Ruiz P. Ratel J. Durif C. Chalancon S. Amiard F. Edely M. Microplastics: what happens in the human digestive tract? First evidences in adults using in vitro gut models J. Hazard. Mater. 442 2023 130010 10.1016/j.jhazmat.2022.130010
38 Choi D. Bang J. Kim T. Oh Y. Hwang Y. Hong J. In vitro chemical and physical toxicities of polystyrene microfragments in human-derived cells J. Hazard. Mater. 400 2020 123308 10.1016/j.jhazmat.2020.123308
39 Gautam R. Jo J. Acharya M. Maharjan A. Lee D. P.B. K.c.Kim C. Kim K. Evaluation of potential toxicity of polyethylene microplastics on human derived cell lines Sci. Total Environ. 838 2022 156089 10.1016/j.scitotenv.2022.156089
40 Lake Ontario Waterkeeper Lake Ontario http://www.waterkeeper.ca/lake-ontario
41 Pinto da Costa J. Reis V. Paço A. Costa M. Duarte A.C. Rocha-Santos T. Micro(nano)plastics–Analytical challenges towards risk evaluation TrAC – Trends Anal. Chem. 111 2019 173 184 10.1016/j.trac.2018.12.013
42 Jakubowicz I. Enebro J. Yarahmadi N. Challenges in the search for nanoplastics in the environment—a critical review from the polymer science perspective Polym. Test. 93 2021 106953 10.1016/j.polymertesting.2020.106953
43 Madejski G.R. Ahmad S.D. Musgrave J. Flax J. Madejski J.G. Rowley D.A. DeLouise L.A. Berger A.J. Silicon nanomembrane filtration and imaging for the evaluation of microplastic entrainment along a municipal water delivery route Sustainability 12 2020 10655 10.3390/su122410655
44 DesOrmeaux J.P.S. Winans J.D. Wayson S.E. Gaborski T.R. Khire T.S. Striemer C.C. McGrath J.L. Nanoporous silicon nitride membranes fabricated from porous nanocrystalline silicon templates Nanoscale 6 2014 10798 10805 10.1039/C4NR03070B 25105590
45 Lee K.-W. Shim W.J. Kwon O.Y. Kang J.-H. Size-dependent effects of micro polystyrene particles in the marine copepod Tigriopus japonicus Environ. Sci. Technol. 47 2013 11278 11283 10.1021/es401932b 23988225
46 Jeong C.-B. Won E.-J. Kang H.-M. Lee M.-C. Hwang D.-S. Hwang U.-K. Zhou B. Souissi S. Microplastic size-dependent toxicity, oxidative stress induction, and p-JNK and p-p38 activation in the Monogonont Rotifer (Brachionus koreanus) Environ. Sci. Technol. 50 2016 8849 8857 10.1021/acs.est.6b01441 27438693
47 New York State Hamlin Beach State Park https://parks.ny.gov/parks/hamlinbeach/details.aspx
48 Monroe County NY–Ontario Beach Park https://www.monroecounty.gov/parks-ontariobeach
49 City of Rochester Durand Eastman Beach https://www.cityofrochester.gov/durandbeach/
50 Monroe County NY–Webster Park https://www.monroecounty.gov/parks-webster
51 Erni-Cassola G. Gibson M.I. Thompson R.C. Christie-Oleza J.A. Lost, but found with Nile Red: a novel method for detecting and quantifying small microplastics (1 mm to 20 μm) in environmental samples Environ. Sci. Technol. 51 2017 13641 13648 10.1021/acs.est.7b04512 29112813
52 Phelan-Dickinson S.J. Palmer B.C. Chen Y. DeLouise L.A. The UVR filter octinoxate modulates aryl hydrocarbon receptor signaling in keratinocytes via inhibition of CYP1A1 and CYP1B1 Toxicol. Sci. Off. J. Soc. Toxicol. 177 2020 188 201 10.1093/toxsci/kfaa091
53 Qin C. Aslamkhan A.G. Pearson K. Tanis K.Q. Podtelezhnikov A. Frank E. Pacchione S. Pippert T. AhR activation in pharmaceutical development: applying liver gene expression biomarker thresholds to identify doses associated with tumorigenic risks in rats Toxicol. Sci. 171 2019 46 55 10.1093/toxsci/kfz125 31127949
54 Corcoran P.L. Belontz S.L. Ryan K. Walzak M.J. Factors controlling the distribution of microplastic particles in benthic sediment of the Thames River, Canada Environ. Sci. Technol. 54 2020 818 825 10.1021/acs.est.9b04896 31884780
55 NOAA-Great Lakes Environmental Research Laboratory Great Lakes Currents Map https://www.glerl.noaa.gov/res/glcfs/currents/
56 Corcoran P.L. Norris T. Ceccanese T. Walzak M.J. Helm P.A. Marvin C.H. Hidden plastics of Lake Ontario, Canada and their potential preservation in the sediment record Environ. Pollut. 204 2015 17 25 10.1016/j.envpol.2015.04.009 25898233
57 Hoffman M.J. Hittinger E. Inventory and transport of plastic debris in the Laurentian Great Lakes Mar. Pollut. Bull. 115 2017 273 281 10.1016/j.marpolbul.2016.11.061 27988025
58 Rao Y.R. Murthy C.R. Nearshore currents and turbulent exchange processes during upwelling and downwelling events in Lake Ontario J. Geophys. Res. Oceans 106 2001 2667 2678 10.1029/2000JC900149
59 Demirbilek Z. Lin L. Hayter E. O'Connell C. Mohr M. Chader S. Forgette C. Modeling of waves Hydrodynamics and Sediment Transport for Protection of Wetlands at Braddock Bay 2015 US Army Corps of Engineers New York https://apps.dtic.mil/sti/tr/pdf/ADA614136.pdf
60 Mazumdar S. Chitkara D. Mittal A. Exploration and insights into the cellular internalization and intracellular fate of amphiphilic polymeric nanocarriers Acta Pharm. Sin. B 11 2021 903 924 10.1016/j.apsb.2021.02.019 33996406
61 Wu B. Wu X. Liu S. Wang Z. Chen L. Size-dependent effects of polystyrene microplastics on cytotoxicity and efflux pump inhibition in human Caco-2 cells Chemosphere 221 2019 333 341 10.1016/j.chemosphere.2019.01.056 30641374
62 Swanson H.I. Cytochrome P450 expression in human keratinocytes: an aryl hydrocarbon receptor perspective Chem. Biol. Interact. 149 2004 69 79 10.1016/j.cbi.2004.08.006 15501429
63 Safe S. Han H. Goldsby J. Mohankumar K. Chapkin R.S. Aryl hydrocarbon receptor (AhR) ligands as selective AhR modulators: genomic studies Curr. Opin. Toxicol. 11–12 2018 10 20 10.1016/j.cotox.2018.11.005
64 Mallin M.A. Johnson V.L. Ensign S.H. Comparative impacts of stormwater runoff on water quality of an urban, a suburban, and a rural stream Environ. Monit. Assess. 159 2009 475 491 10.1007/s10661-008-0644-4 19067207
65 Al-Mashaqbeh O. Jiries A. El-HajAli Z. Stormwater runoff quality generated from an urban and a rural area in the Amman-Zarqa basin WIT Trans. Ecol. Environ. 182 2014 379 390 10.2495/WP140331
66 Jacob A. Hartz A.M. Potin S. Coumoul X. Yousif S. Scherrmann J.-M. Bauer B. Declèves X. Aryl hydrocarbon receptor-dependent upregulation of Cyp1b1 by TCDD and diesel exhaust particles in rat brain microvessels Fluids Barriers CNS 8 2011 23 10.1186/2045-8118-8-23 21867498
67 Procházková J. Kozubík A. Machala M. Vondráček J. Differential effects of indirubin and 2,3,7,8-tetrachlorodibenzo-p-dioxin on the aryl hydrocarbon receptor (AhR) signalling in liver progenitor cells Toxicology 279 2011 146 154 10.1016/j.tox.2010.10.003 20951181
