Foaming at the mouth: Ingestion of floral foam microplastics by aquatic animals

Journal Pre-proof Foaming at the mouth: Ingestion of floral foam microplastics by aquatic animals

Charlene Trestrail, Milanga Walpitagama, Claire Hedges, Adam Truskewycz, Ana Miranda, Donald Wlodkowic, Jeff Shimeta, Dayanthi Nugegoda PII:

S0048-9697(19)35821-8

DOI:

https://doi.org/10.1016/j.scitotenv.2019.135826

Reference:

STOTEN 135826

To appear in:

Science of the Total Environment

Received date:

4 October 2019

Revised date:

20 November 2019

Accepted date:

26 November 2019

Please cite this article as: C. Trestrail, M. Walpitagama, C. Hedges, et al., Foaming at the mouth: Ingestion of floral foam microplastics by aquatic animals, Science of the Total Environment (2019), https://doi.org/10.1016/j.scitotenv.2019.135826

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© 2019 Published by Elsevier.

Journal Pre-proof Foaming at the mouth: ingestion of floral foam microplastics by aquatic animals Charlene Trestrail1*, Milanga Walpitagama2, Claire Hedges1, Adam Truskewycz3, Ana Miranda1, Donald Wlodkowic2, Jeff Shimeta1, Dayanthi Nugegoda1

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Centre for Environmental Sustainability and Remediation, School of Science, RMIT University,

Bundoora, Victoria, Australia.

Phenomics Laboratory, School of Science, RMIT University, Melbourne, VIC 3083, Australia

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Advanced Manufacturing and Fabrication, School of Engineering, RMIT University, VIC 3000,

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Australia *

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Correspondence to: Charlene Trestrail, Ecotoxicology Laboratory, School of Science, RMIT

University, Plenty Road, PO Box 71, Bundoora, VIC 3083, Australia, E-mail:

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[email protected]

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Journal Pre-proof Abstract Phenol-formaldehyde plastics are used globally as floral foam and generate microplastics that can enter the environment. This study is the first to describe how aquatic animals interact with this type of microplastic, and the resultant physiological responses. We analysed “regular foam” microplastics generated from petroleum-derived phenol-formaldehyde plastic, and “biofoam” microplastics generated from plant-derived phenol-formaldehyde plastic. Regular foam and biofoam microplastics

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showed similar FTIR spectra. Both types of microplastics were consumed by all six invertebrate

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species tested: the freshwater gastropod Physa acuta, the marine gastropod Bembicium nanum, the

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marine bivalve Mytilus galloprovincialis, adults and neonates of the freshwater crustacean Daphnia magna, the marine amphipod Allorchestes compressa, and nauplii of the marine crustacean Artemia

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sp. For all species, the occurrence of ingestion was similar for regular foam and biofoam

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microplastics. Biofoam microplastics leached more than twice as much phenolic compounds than regular foam microplastics. The leachates from regular foam and biofoam microplastics showed the

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same acute toxicity to Artemia nauplii (24-h LC50 =27.4 mg mL-1 and 22.8 mg mL-1, respectively) and

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D. magna (48-h LC50 =17.8 mg mL-1 and 15.3 mg mL-1, respectively). However, biofoam microplastic

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leachate was twice as toxic to embryos of the zebrafish, Danio rerio, compared with leachate from regular foam microplastic (96-h LC50 = 43.8 mg mL-1 vs 27.1 mg mL-1). Using M. galloprovincialis, we show that regular foam microplastic leachate and the physical presence of the microplastics exerted separate and cumulative effects on catalase (CAT) activity, glutathione-s-transferase (GST) activity and lipid peroxidation. Microplastic ingestion did not affect the activity of acetylcholinesterase (AChE). Taken together, these results show that phenol-formaldehyde microplastics can interact with a range of aquatic animals, and affect sublethal endpoints by leaching toxic compounds, and through the physical presence of the microplastics themselves.

Keywords: floral foam; phenol-formaldehyde resin; ingestion; leachate; acute toxicity; oxidative stress. 2

Journal Pre-proof Abbreviations: AChE, acetylcholinesterase; CAT, catalase; GST, glutathione-s-transferase; ROS, reactive oxygen species; SEM, scanning electron microscopy; TBARS, thiobarbituric acid reactive substances.

1. Introduction Since plastic was first mass produced in the 1950s, plastic demand has increased at a phenomenal rate and global production currently exceeds 320 million tonnes (PlasticsEurope, 2016; Thompson et

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al., 2009; Worm et al., 2017). The biggest proportion of plastics is manufactured for single-use applications (Geyer et al., 2017; Hopewell et al., 2009), which generate large amounts of plastic

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waste that can escape into the environment (Jambeck et al., 2015). Through the actions of

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mechanical stress, heat and UV exposure (Andrady, 2011), this plastic waste can fragment into microplastics measuring < 5 mm (Arthur et al., 2009; Koelmans et al., 2015; Rios Mendoza et al.,

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2018). Additionally, some plastic particles are manufactured in the microplastic size range for use in

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domestic products, and so are already classed as microplastics when they escape into the

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environment (Cheung and Fok, 2017).

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Microplastics can significantly affect organism health. Studies have observed microplastic consumption in several invertebrate taxa and across various feeding modes (Allen et al., 2017a; Avio et al., 2015; Cole et al., 2013; Hämer et al., 2014; Sussarellu et al., 2016; Van Cauwenberghe et al., 2015; Vroom et al., 2017; Welden & Cowie, 2016), leading to a range of sublethal responses including suppressed somatic growth rates and diminished reproductive output (reviewed by Trestrail et al., 2019). Under particular conditions, microplastic ingestion can result in death (Jemec et al., 2016; Ziajahromi et al., 2018). These responses are concerning because they have the potential to affect animal populations that maintain crucial ecosystem services (Mulder et al., 1999). Most research on biological responses has focussed on polyethylene and polypropylene microplastics; the biological responses to several other families of plastic are entirely unstudied (de

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Journal Pre-proof Sá et al., 2018). Phenol-formaldehyde resins are a type of plastic for which no biological response information exists, even though this plastic type has been commercially manufactured longer than any other family of synthetic polymers (de Sá et al., 2018; Pathak et al., 2014). This knowledge gap has now become pertinent because phenol-formaldehyde resins are at the centre of a viral social media trend (described below) that promotes microplastic-generating activities which can increase the amount of phenol-formaldehyde microplastics entering the environment.

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Phenol-formaldehyde resin is used to manufacture blocks of floral foam, which has been used

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globally in the floristry industry since the 1950s as a method to support the stems of flowers in floral

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arrangements (Cornick, 2010; Smithers, 1956). The open cells of the foam are friable and easily rupture (Del Saz-Orozco et al., 2015; Mougel et al., 2019), so floral foam readily generates secondary

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microplastics (as defined by Cole et al., 2011). In addition to floral foam’s widespread use and

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increasing global demand in the floristry industry (Cornick, 2010), floral foam has become popular to a more general consumer base: videos of people using their fingers to crush blocks of floral foam

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have spread virally across social media platforms such as YouTube and Instagram (Gallagher, 2019;

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Smith and Snider, 2019; Tansill-Suddath, 2018). This practice generates innumerable microplastics.

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Floral foam users can release phenol-formaldehyde microplastics into the environment through several disposal routes. Foam microplastics disposed of via domestic wastewater systems can be discharged into rivers and oceans from waste-water treatment facilities (Lebreton et al., 2017; Mason et al., 2016; Murphy et al., 2016), and be dispersed across large distances through wind action (Chubarenko et al., 2016). Phenol-formaldehyde microplastics can also contaminate terrestrial ecosystems when water with suspended microplastics is applied directly to garden soil, if phenol-formaldehyde microplastics are in biosolid sludge applied to arable land as fertiliser (Mahon et al., 2017; Rochman et al., 2015), and when floral foam is composted (Weithmann et al., 2018). Even phenol-formaldehyde microplastics that are disposed of via landfill can escape into the surrounding environment in landfill leachate (He et al., 2019). The mobility of microplastics through

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Journal Pre-proof the environment increases the likelihood that animals will encounter and interact with phenolformaldehyde microplastics. The limited understanding of the biological responses to phenol-formaldehyde resins and microplastics has sparked industry and community concern (Feldmann, 2019; Latona, 2018; Straub, 2009). It is well known that phenol-formaldehyde plastic is synthesised from toxic, petroleum-based phenol and formaldehyde monomers (SI Fig 1) which can leach from plastics into the surrounding

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environment (Lithner et al., 2011; Tingley et al., 2017; Zjawiony, 1992). Yet, to the best of our

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knowledge, the only research into biological responses to phenol-formaldehyde plastic ingestion is

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limited to basic toxicity tests conducted on rats (discussed in Gardziella et al., 2000, Chapter 8, p. 516). This data has limited applicability to other animals. Even less is known about biological

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responses to bio-based phenol-formaldehyde plastics manufactured from plant-derived phenolic

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monomers, which have recently become commercially available (Li et al., 2017; Wang et al., 2018). This study addresses this poor understanding of how phenol-formaldehyde microplastics affect

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animals by exploring the biological responses to phenol-formaldehyde microplastics generated from

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two types of phenol-formaldehyde foams. We firstly characterised the physical and chemical nature

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of the foams and determined the potential for microplastics generated from these foams to be ingested by a range of aquatic animals with different feeding modes. We then determined whether these microplastics could leach chemicals that were acutely toxic to aquatic animals. Finally, we evaluated whether microplastic leachate and the physical effects of the microplastics cause different sublethal cellular responses by measuring the activities of the enzymes catalase (CAT), glutathione-Stransferase (GST) and acetylcholinesterase (AChE), and the concentration of malondialdehyde as a biomarker of lipid peroxidation. 2. Materials and methods

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Journal Pre-proof We used microplastics generated from two types of phenol-formaldehyde floral foam manufactured by Smithers-Oasis (USA): Oasis® Ideal Floral Foam (“regular foam”) and Oasis® Enhanced Biofoam (“biofoam”). Manufacturing specifications confirm that regular foam is a phenol-formaldehyde plastic (Oasis Products, 2013), however, corresponding specifications for biofoam are not publicly available (Koch & Co, 2019; Oasis Floral Products, 2019).

2.1. Characterisation of floral foam microplastics

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A range of techniques were used to assess the physical and chemical characteristics of the foams and foam microplastics. To visualise the physical, three-dimensional structure of the foams, we

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observed foam samples using scanning electron microscopy (SEM). Light microscopy was used to

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observe the shape and thickness of microplastics generated from the foams. To address the knowledge gap regarding the chemical specifications of the foams, we used Attenuated Total

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Reflection Fourier-Transform Infrared (ATR-FTIR) to determine the chemical fingerprint of both

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foams, and identified these against known plastic spectra in a database.

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The physical structure of the foam was characterised using a scanning electron microscope (Phillips XL30) at an operating voltage of 25 kV and a spot size of 5. Particles were sputter-coated with gold

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prior to imaging. Using a mortar and pestle, foams were crushed to generate secondary microplastics, which were examined using light microscopy (DM500, Leica microsystems). The chemical nature of the foam microplastics was characterised directly using ATR-FTIR, a technique that has previously been used to characterise regular and plant-based phenolformaldehyde foams (Hu et al., 2012; Li et al., 2017; Tseng and Kuo, 2002). Following the procedure of Li et al. (2017), foam samples were crushed and pressed in a macro-ATR accessory (GladiATR, PIKE Technologies) coupled with an FTIR spectrometer equipped with a deuterated lanthanum α alaninedoped triglycine sulphate detector (Cary 670, Agilent). Spectra were recorded at room temperature in the frequency range 400–4000 cm-1 at a resolution of 4 cm-1 using 64 co-addition scans. To reduce

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Journal Pre-proof noise and optimise database searching, spectra were processed with in-program Optimized Corrections before being compared to polymer-related spectral databases (listed in SI Table 1) using the KnowItAll software (Bio-Rad Laboratories Inc., USA). We report the top five matches with their respective Hit Quality Index (HQI), a scale between 0 – 100 which indicates how well the database spectrum matches the test spectrum.

2.2. Ingestion of floral foam microplastics

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Currently, no data exists about whether phenol-formaldehyde microplastics are ingested by animals; therefore, we conducted ingestion tests. We selected organisms from a range of feeding modes,

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because feeding mode can influence microplastic ingestion (Setälä et al., 2016). Since phenol-

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formaldehyde microplastics can contaminate both ecosystems when they are disposed of via domestic wastewater systems (see Introduction), we tested both freshwater and marine animals in

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the ingestion tests.

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We assessed ingestion in the freshwater gastropod Physa acuta and the freshwater pelagic

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crustacean Daphnia magna, as well as the following marine invertebrates: the bivalve Mytilus

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galloprovincialis, the gastropod Bembicium nanum, the amphipod Allorchestes compressa, and nauplii of the pelagic crustacean Artemia sp. All species except B. nanum have been reported to eat microplastics made from polymers other than phenol-formaldehyde (Avio et al., 2015; Chua et al., 2014; Ogonowski et al., 2016; Scherer et al., 2017; Wang et al., 2019). Organisms were either collected from locations in Victoria, Australia, or purchased from biological specimen suppliers (SI Table 2). Regular foam and biofoam microplastics were generated by freshly crushing foam and were sized using a laser particle sizer (Mastersizer 3000, Malvern) according to Napper et al. (2015). The volume-weighted mean diameter ± SD was taken from an average of three measurements and

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Journal Pre-proof determined as 170.4 ± 147.5 µm for regular foam microplastics, and 155.9 ± 56.4 µm for biofoam microplastics. Ten individuals of a similar size (SI Table 2) were chosen for each species and starved for 24 h to clear their digestive tracts. Individuals of the same species were then exposed together in a glass petri dish of appropriate media (see SI for details) containing 1 mg mL-1 freshly crushed microplastics from regular foam or biofoam. No aeration was used during the exposure, which allowed

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microplastics to settle to the bottom of the vessel and become accessible to the grazing

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invertebrates. Exposure lasted for 2 h, except for D. magna which was exposed for 24 h to account

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for the fact that this species is typically not a bottom feeder (Ebert, 2005). D. magna birthed offspring during the exposure; 10 of these neonates were randomly selected to assess whether

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microplastic ingestion had occurred in this age class.

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To ascertain whether microplastic ingestion had occurred, we observed D. magna adults and neonates, and Artemia nauplii, under a microscope (DM2500, Leica microsystems) to determine the

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presence of microplastics in the digestive tract of live individuals. Organisms that had been starved

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for 24 h but not exposed to microplastics were observed as controls. The digestive tracts of the

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remaining species were not visible using microscopy. These animals were placed individually in separate beakers, and the faeces produced within 24 h of exposure ending were collected. Squash preparations of faecal pellets were analysed using microscopy for the presence of microplastics; only microplastics embedded within the faecal pellets, rather than microplastics attached to the surface, were considered as evidence of ingestion.

2.3. Toxicity of floral foam microplastic leachate Several types of plastic leach toxic chemicals into the surrounding environment (Li et al., 2016). Phenol-formaldehyde plastics are also known to leach chemicals, such as unpolymerized monomers, into the surrounding atmosphere (Zjawiony, 1992). We tested whether phenol-formaldehyde

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Journal Pre-proof microplastics leach monomers into aquatic environments, and then determined the toxicity of microplastic leachates using standard toxicity tests on three model aquatic organisms.

2.3.1. Leachate preparation Freshly crushed foam microplastics were mixed with sterile distilled water (18 MΩ, Milli-Q) to create a slurry with the concentration 50 mg microplastics mL-1. Microplastic slurries were shaken in polystyrene tubes at 250 rpm for 24 h at 25 °C (as per Lithner et al., 2012, 2009), then filtered to

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0.45 µm through a syringe filter (Millex®, Merck) to remove microplastics and obtain a clarified leachate stock. Distilled water subjected to the same extraction process was used as a negative

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control. The pH, salinity and dissolved oxygen concentration of leachates was recorded (HQ30d

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2.3.2. Microplastic leachate toxicity tests

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meter, Hach).

Microplastic leachate stock was used in toxicity tests within 3 hours of filtration. Both leachates

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were extremely acidic after the leaching process was completed (SI Table 3) and the pH needed to

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be adjusted prior to conducting toxicity tests in order to meet the specifications of the test protocols

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for each animal. Immediately prior to use, the leachate stocks were adjusted to the required pH using 50% NaOH (< 1% v/v added) and diluted to the test concentrations (equivalent to 0 mg mL-1, 10 mg mL-1, 20 mg mL-1, 30 mg mL-1, 40 mg mL-1, 50 mg mL-1) in the appropriate medium for each species. All leachate toxicity tests were conducted under static conditions in darkness at 28 ± 0.5 °C. To test the toxicity of leachate to invertebrates, we conducted toxicity tests with Artemia nauplii and D. magna neonates (see SI Table 2 for sources). These are standard invertebrates used for toxicity testing, and a large body of information exists regarding their responses to contaminants (Libralato et al., 2016; Persoone et al., 2009). Artemia cysts were hatched in sterile artificial seawater (37.5 mS cm-1, Ocean Fish, Prodac), and 30 h old instar II-III nauplii were used in a standard 24 h toxicity test

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Journal Pre-proof (Microbiotests, 2014). For the D. magna toxicity test, ephippia were hatched in media according to the manufacturer’s instructions, and 24 h old neonates were used in 48 h toxicity tests (OECD, 2004). To assess the effects of leachates on aquatic vertebrates, we conducted Fish Embryo Acute Toxicity tests using the zebrafish, Danio rerio (Tübingen strain, see SI for animal husbandry details), according to OECD test methods (OECD, 2013) with the modifications of Walpitagama et al. (2019). Fertilised eggs were identified by the development of a blastula, and those with no obvious defects in

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morphology or in the chorion were selected. At 5 hours post-fertilisation, 15 embryos were

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randomly allocated to each treatment and incubated individually in the wells of polystyrene 96-

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multiwell plates containing 200 µl of leachate diluted in E3 solution. Hatching rate and mortality were visually determined after 96 h exposure. Mortality was identified when one of the following

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conditions was met: coagulation of embryos, lack of somite formation, non-detachment of the tail,

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or lack of heartbeat (OECD, 2013).

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2.3.3. Phenolic compound quantification

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Phenolic compounds are known to leach out of phenol-formaldehyde resins (Zjawiony, 1992) and

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these can cause toxicity to aquatic organisms. To assess whether phenolic compounds could have contributed to the toxicity of the tests in 2.3.2, we analysed the phenolic content of leachates from regular foam and biofoam microplastics and determined if phenolic concentration was stable over 96-h, which covers the duration of the leachate toxicity tests. Using the microplate method of Attard (2013), 10 µl of leachate was combined with 100 µl Folin-Ciocalteu reagent (1:10 dilution), and 80 µl 1M NaCO3 was added. The tests were incubated at room temperature in the dark for 30 min, and absorbance was read at 630 nm in a microplate reader (POLARstar, Omega). Sample blanks containing distilled water instead of Folin-Ciocalteu reagent were used to correct for background absorbance. Total phenolic content was determined using a standard curve of gallic acid, and the results were expressed as gallic acid equivalents (µg mL-1).

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Journal Pre-proof 2.4. Toxicity of floral foam microplastics: separating the effects of leachate and physical presence To determine whether the effects of microplastic leachate could be differentiated from the physical effects of the microplastics, we conducted an experiment using M. galloprovincialis and regular foam microplastics. Since microplastic ingestion is rarely lethal (as noted by Galloway et al., 2017), we selected four sublethal endpoints commonly used in microplastic studies (Prokid et al., 2019) to

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assess the effects of microplastic leachate and physical presence. Mussels were selected as the test

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organism because their substantial biomass enables the quantification of multiple sublethal

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endpoints, thus providing a comprehensive assessment of biological responses. Further, Mytilus have previously been used to assess sublethal responses to other types of microplastics (Avio et al.,

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2015; Paul-Pont et al., 2016).

2.4.1. Animal collection

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Mussels were collected in September 2018 from Newport, Australia (37° 50' 48'' S, 144° 53' 55''

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E), and acclimated for four days in a communal tank of aerated natural seawater at ambient

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sea surface temperature (12 °C) with a 16 h light cycle. 2.4.2. Experimental treatments We used four treatments to determine whether phenol-formaldehyde microplastic leachate and the physical presence of the microplastics caused different responses in mussels: (1) a ‘leachate only’ treatment, prepared using the method in section 2.3.1 and stored at 4 °C for use throughout the experiment; (2) a ‘microplastics only’ treatment free of potential leachates (see 2.4.2.1 for method); (3) a ‘microplastics + leachate’ treatment, made from freshly ground microplastics containing potential leachates; and (4) a seawater control. All treatments used natural seawater filtered to 0.22

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Journal Pre-proof µm to prevent waterborne algae and bacteria from influencing microplastic behaviour in the water column (Galloway et al., 2017).

2.4.2.1. Removal of microplastics leachate To create the ‘microplastics only’ treatment, microplastics were cleaned of potential leachates by soaking 500 mg of freshly crushed regular foam microplastics in 1 L distilled water (18 MΩ, Milli-Q) in a glass bottle. The bottle was shaken at 250 rpm at 25 °C for 1 h and the microplastics were retrieved

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by vacuum filtration (0.22 µm, PES Express Membrane, Millipore). This process was repeated three times with distilled water, once with 50% methanol, and a final time with distilled water to remove

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residual methanol. The clean microplastics were dried to a constant weight at 40 °C.

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2.4.2.2. Treatment exposure

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For each treatment, individual mussels were placed in separate beakers (n = 6) containing 80 mL natural seawater filtered to 0.22 µm. For the ‘leachate only’ treatment, leachate stock was dosed in

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each beaker at a concentration equivalent to leachate from 1 mg microplastics mL-1. For the

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‘microplastics only’ and ‘microplastics + leachate’ treatments, the relevant microplastics were dosed

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at 1 mg microplastics mL-1. Beakers were provided with gentle aeration and water was renewed daily. Mussels were not fed during the experiment. After three days’ exposure, mussels were dissected on ice and homogenised in 50 mM potassium phosphate buffer (pH 7.6, 4 °C) containing EDTA-free SigmaFAST™ protease inhibitor. Homogenates were centrifuged for 15 min at 4000 x g at 4 °C, and the resultant supernatant was stored at -80 °C until used for biochemical analyses.

2.4.3 Biomarker analyses We assessed sublethal biological responses to treatments by measuring the activities of enzymes and the amount of lipid oxidative damage in freshly thawed supernatant. Unless stated otherwise, all biochemical analyses were conducted in triplicate at room temperature, and absorbance was

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Journal Pre-proof read using a microplate reader (POLARstar, Omega). The results of all biochemical analyses were normalised to protein content, which was determined according to Bradford (1976) using bovine serum albumin as a standard.

CAT activity (EC 1.11.1.6) We analysed CAT activity because this enzyme provides the first line of defence against oxidative damage by degrading toxic H2O2, the concentrations of which can rapidly increase when organisms

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are exposed to stressors, including microplastics (Magara et al., 2018; Tang et al., 2018). CAT activity was determined using a modified method of Hadwan & Abed (2016). Briefly, 25 µl of supernatant

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was added to 250 µl of 30 mM H2O2 (Aebi, 1990). The reaction proceeded at room temperature for 4

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min before termination by the addition of 1 mL 32.4 mM ammonium molybdate. The absorbance of the resulting chromophore was read at 374 nm. Specific CAT activity was calculated using the

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equation for a first-order reaction (Hadwan and Abed, 2016), and results are presented as kU min-1

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mg-1 protein.

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GST activity (EC 2.5.1.18)

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GST catalyses the degradation of xenobiotics and is a sensitive biomarker for phenolic compounds (Vidal-Liñán et al., 2015). GST activity was measured according to the methods of Habig et al. (1974) with adaptations for use in a microplate by Frasco & Guilhermino (2002). Briefly, 200 µl of reaction mixture (1.5 mM 1-chloro-2,4-dinitrobenzene and 1.5 mM L-glutathione reduced in 0.1 M potassium phosphate buffer, pH 6.5) was added to 25 µl of supernatant, and the increase in absorbance at 340 nm was measured for 5 min. Homogenising buffer was used as a blank. The amount of substrate hydrolysed was determined using ε = 9.6 mM-1 cm-1. Results are presented as nmol substrate hydrolysed min-1 mg-1 protein.

AChE activity (EC 3.1.1.7)

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Journal Pre-proof AChE activity is a well-established biomarker of neurotoxicity and has been used to identify the effects of microplastics (Prokid et al., 2019). We measured AChE activity by adapting the standard Ellman et al. (1961) method to a microplate. In a 96-multiwell plate, 12.5 µl of supernatant was combined with 137.5 µl reaction mixture (0.9 mM 5,5′-dithiobis(2-nitrobenzoic acid) and 68.2 µM acetylthiocholine iodide in 0.1 M phosphate buffer, pH 7.2) and incubated at 25 °C for 10 min. The production of thiols was monitored at 405 nm for 5 min. Temperature was strictly controlled in order to confidently use ε = 14.05 x 103 M-1 cm-1 (Eyer et al., 2003). AChE activity is presented as

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nmol substrate hydrolysed min-1 mg-1 protein.

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Lipid peroxidation

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Periods of oxidative stress are characterised by increased oxidative damage to lipids, which can be quantified by measuring the lipid peroxidation product, malondialdehyde (Vlahogianni et al., 2007).

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We quantified malondialdehyde using the thiobarbituric acid reactive substances (TBARS) method of

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Parrilla-Taylor et al. (2013). In an Eppendorf tube, 90 µl of supernatant was incubated at 37 °C for 15 min. Then, 180 µl of stopping solution (12.5% trichloroacetic acid, 1% thiobarbituric acid in 1 M HCl)

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was added and the sample was incubated at 90 °C for 10 min. The samples were cooled to room

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temperature in a water bath and centrifuged at 10 000 x g for 10 min to remove precipitate. The absorbance of the supernatant was measured at 352 nm against a blank that did not contain any sample. TBARS were quantified using ε = 155 mM-1 cm-1, and the results are presented as nmol TBARS mg-1 protein.

2.5. Statistical analyses All statistical analyses were conducted using SPSS Statistics (v25, IBM). The mean size of regular foam and biofoam microplastics used for ingestion studies were compared using an independent samples t-test, and differences in foam ingestion across species were assessed using two-tailed Fischer’s exact tests. For leachate toxicity tests, LC50 concentrations were calculated using a probit

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Journal Pre-proof regression and results were presented as mg microplastic mL-1. A repeated measures ANOVA was conducted to determine if the phenolic content differed between leachates and over time; Mauchly’s test of sphericity indicated that the assumption of sphericity was violated (χ 2(9) = 21.332, p = 0.029), so a Greenhouse-Geisser correction (ε = 0.299) was applied. Biomarker data were analysed with a one-way multivariate analysis of variance and a post hoc Tukey’s test was applied to

Results

3.1.

Characterisation of foam microplastics

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2.

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evaluate statistical difference. Significance was set at  = 0.05.

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Both the regular foam and the biofoam consisted of an open-celled honeycomb structure: thick

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resinous struts formed the framework of the foam, and membranous films formed the cell walls (Fig

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1). In the regular foam, thick resin connected the cell walls to the surrounding struts at several attachment points (Fig 1). The cell walls of biofoam, however, were attached to the surrounding

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struts with thin resin, and at fewer attachment points. This difference in morphology suggests that

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the biofoam is more friable than regular foam. Both the thick resinous struts and the membranous

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walls were present as secondary microplastics when the foams were crushed (Fig 2). The regular foam and biofoam displayed similar ATR-FTIR spectra (Fig 3). Both foams showed peaks consistent with phenol-formaldehyde spectra reported in the literature. Specifically, regular foam and biofoam had typical hydroxyl group absorption between 3300 – 3400 cm-1 and C-H stretching at 2900 cm-1. We attribute the clear peaks observed in both spectra at 1600 cm-1, 1500 cm-1, and 1200 cm-1 to phenolic bands (Li et al., 2017). Compared with regular foam, biofoam showed slightly more absorption at 1042 cm-1, which could be attributed to additional sulfonate groups present (Hu et al., 2012). The spectra of both foams strongly resembled (HQI scores > 94) the spectra of alkalicondensed phenol-formaldehyde resins (Table 1). For regular foam and biofoam, the highest-ranked spectral match (HQI > 97 and > 96, respectively) was with the compound poly(2,6methylenephenol), and the next four highest-ranked matches for both foams were with phenol15

Journal Pre-proof formaldehyde resins. The biofoam spectrum did not strongly match with the spectra of any plantderived plastics available in the spectral libraries.

3.2.

Ingestion of floral foam microplastics

The mean size of regular foam microplastics (170.4 ± 147.5 µm) was not significantly different from that of the biofoam microplastics (155.9 ± 56.4 µm, t(5) = 0.185, p = 0.861). All the invertebrate species tested demonstrated the capacity to directly ingest phenol-formaldehyde foam microplastics

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(Table 2). The occurrence of ingestion did not significantly differ between regular foam and biofoam microplastics (p > 0.05 for all species, two-tailed Fischer’s exact test). In Artemia nauplii and D.

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magna adults and neonates, foam microplastics were seen densely compacted along the length of

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the digestive tract (Fig 4, SI Fig 2). Whilst individual microplastics could not be determined in the digestive tract, clumps of microplastics were observed moving within the gut lumen of D. magna

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through peristaltic movements (SI Video 1). Microplastic-laden faecal pellets were egested by the

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gastropods, bivalve and amphipod species tested (Fig 4). Resin-rich microplastics, derived from the struts of the floral foam, were easily detected in faecal pellets, and squash preparations of the faecal

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pellets revealed that membranous microplastics generated from the foam cell walls were also

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present in faeces. Due to the variable sizes and shape of the microplastics prior to ingestion, we were unable to ascertain whether passing through the digestive system degraded the microplastics.

3.3.

Toxicity of floral foam leachate

LC50 values generated from leachate exposure are presented in Table 3. The regular foam leachate and the biofoam leachate showed the same level of toxicity to Artemia nauplii, generating respective LC50 values of 27.4 mg microplastics mL-1 and 22.8 mg microplastics mL-1 after 24 h exposure. We observed that a precipitate had formed by the end of the Artemia exposure experiment. For D. magna neonates, leachate from regular foam microplastics was slightly more toxic at 24 h (LC50 = 36.2 mg microplastics mL-1) than biofoam microplastics leachate (LC50 = 26.9 mg microplastics mL-1);

16

Journal Pre-proof however, there was no difference in toxicity after 48 h. For D. rerio embryos, biofoam leachate was almost twice as toxic as regular foam leachate (LC50 = 27.1 mg microplastics mL-1 and 43.8 mg microplastics mL-1, respectively) after 96 h exposure. Leachate from biofoam microplastics contained significantly more phenolic compounds than leachate generated from regular foam microplastics (F(1,4) = 1846.361, p <0.01; Fig 5). The concentration of phenolic compounds in the leachates did not change significantly over 96 h

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(F(1.197, 4.786) = 5.943, p < 0.05), indicating that treatments remained consistent for the duration

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of all the leachate toxicity tests conducted. There were no significant interactions between time and

Toxicity of floral foam microplastics: separating the effects of leachate and physical

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3.4.

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leachate type (F(1.197, 4.786) = 2.294, p > 0.05).

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presence

No mortality was observed during the experiment. CAT activity showed a non-significant declining

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trend, with activity progressively declining in the leachate only treatment, then the microplastics

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only treatment, and finally showing the lowest activity in the microplastics + leachate treatment (Fig

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6). GST activity showed the opposite, non-significant increasing trend (Fig 6). AChE activity changed according to treatment and was significantly reduced in the leachate only treatment compared to the microplastics + leachate treatment. TBARS concentration decreased in all treatments compared to the control and was significantly lower in the microplastics-only treatment and the microplastics + leachate treatment.

4.

Discussion

4.1 Characteristics of floral foam microplastics Both regular foam and biofoam possessed the open-celled, honeycomb structure typical of phenolic foams (Cornick, 2010). The SEM images suggest that the cell walls of biofoam were thinner and more

17

Journal Pre-proof friable than the cell walls of regular foam, a phenomenon that has been observed in other phenolic foams manufactured from plant-sourced monomers (Del Saz-Orozco et al., 2015). Biofoam could, therefore, generate more secondary microplastics than regular foam. Quantifying this is difficult as the number of microplastics generated will vary based on the duration and pressure of foam crushing. ATR-FTIR spectroscopy confirmed both regular foam and biofoam as phenol-formaldehyde plastics.

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The similarity of the foams’ spectra supports earlier findings that the source of the phenol

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monomers used to manufacture foams has limited impact of the molecular structure of the resulting

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phenol-formaldehyde foams (Li et al., 2017). This molecular similarity has biological significance because it means that microplastics from both types of foam will have limited biodegradability

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(Gusse et al., 2006; Kaplan et al., 1979), and will persist in the environment where they can interact

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with biota.

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4.2 Ingestion of floral foam microplastics

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This study is the first report of phenol-formaldehyde microplastics ingestion in any organism. We

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demonstrated that floral foam readily generates microplastics in a size-range that is consumed by grazing, filter-feeding and shredding invertebrates, and that ingestion of these microplastics occurred in both freshwater and marine organisms. These results demonstrate that phenolformaldehyde microplastics can interact with a diverse range of species, and can therefore pose the same threats to animals as more commonly studied types of microplastics, such as polyethylene and polystyrene. All the species we tested consumed regular and biofoam microplastics, even though no food was present during the exposure period. Ingestion of other types of microplastics has previously been reported for all the organisms we tested except B. nanum (Avio et al., 2015; Chua et al., 2014; Ogonowski et al., 2016; Scherer et al., 2017; Wang et al., 2019). Why animals ingest microplastics of

18

Journal Pre-proof any polymer type remains undetermined. In addition to particle size, chemoreception likely influences post-capture particle consumption of some invertebrates (Allen et al., 2017; Cole et al., 2013; Rosa et al., 2013). Since we saw no difference in ingestion between regular foam microplastics and biofoam microplastics, we conclude that the ingestion of these microplastics was likely driven by the physical presence of the microplastics. The presence of chemicals which would differ between the two foam types, such as unpolymerized monomers, did not appear to influence ingestion.

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In addition to physical presence, the ingestion of phenol-formaldehyde microplastics may also be

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influenced by some aspect of the molecular structure of the phenol-formaldehyde polymers. Phenol-

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formaldehyde plastics consist of a chain of phenol molecules interspersed with carbon molecules (SI Fig 1). Some organisms may detect the molecular signature of the polymer and interpret this as an

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indication of a nutritious food source. Alternatively, such as signal may be generated by one

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component of the phenol-formaldehyde molecular signature. If this is the case, it is unlikely that the phenol component of the polymer is driving ingestion, since phenolic compounds can reduce food

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consumption (Steinberg, 1988). The interspersed carbon molecules, however, may be interpreted by

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animals as an organic coating, like carbohydrates, thereby signalling nutritional organic content and

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leading to ingestion (Mahon & Dauer, 2005; Taghon, 1982). 4.3 Effects of phenol-formaldehyde microplastic leachate The leaching of phenolic compounds from petroleum-based phenol-formaldehyde plastics is well documented (O’Brien and Olofsson, 1980; Zjawiony, 1992). However, ours is the first report of phenolic compounds leaching from plant-derived phenol-formaldehyde plastics. Like phenol, residual formaldehyde monomers can also leach out of phenol-formaldehyde plastics, but this has only been reported to occur in freshly polymerised resins (Pasch and Schrod, 2004; Poljanšek and Krajnc, 2005). Residual formaldehyde degrades over time because it evaporates below room temperature, and because it is oxidised by atmospheric oxygen (National Center for Biotechnology Information, n.d.). Considering the time lag between the manufacture of the foams and its use in our 19

Journal Pre-proof experiment, and the nature of our leaching protocol, we think it unlikely that appreciable concentrations of formaldehyde were present in the leachates. Despite their different phenolic contents, the leachates caused the same level of toxicity to Artemia nauplii, which was substantially lower than the toxicity of plastic leachates to other planktonic marine crustaceans (Bejgarn et al., 2015). We attribute this discrepancy to the low solubility of phenols in seawater (Noubigh et al., 2007), a phenomenon which also explains the precipitate

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formation during the experiment. For D. magna, 48 h LC50 values for both leachates were

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substantially more toxic than the leachates generated by several other families of plastic, which

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reached as high as 235 mg mL-1 (Lithner et al., 2012, 2009). Whilst biofoam leachate was slightly more toxic than regular foam leachate to D. magna, this difference was striking in D. rerio embryos

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as biofoam leachate was almost twice as toxic as regular foam leachate. This is likely caused by the

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variable sensitivity of D. rerio to different phenolic compounds (Martins et al., 2007); and variability of the chemical leaching from each of the foam formulation.

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4.4 Microplastic ingestion: separating the effects of leachate and physical presence The magnitude of change in the measured biomarker responses typically increased according to the

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order: leachate only < microplastics only < microplastics + leachate treatment. This trend indicates that this order of treatments posed increasingly severe oxidative challenges to M. galloprovincialis, and that microplastic leachate and the physical presence of microplastics exert separate and cumulative effects on organisms. Although biomarkers are commonly employed in microplastic studies (Prokid et al., 2019), they have yielded conflicting intra-study results (as in the case of O’Donovan et al., 2018 and Ribeiro et al., 2017) and, as in our study, statistically insignificant changes (Pittura et al., 2018; Ribeiro et al., 2017; Rodríguez-Seijo et al., 2018a, 2018b). Whilst biomarker responses may be influenced by microplastic treatments, some response variability can also be attributed to characteristics of the organism and to experimental conditions, such as age, seasonality, feeding regime and toxicant exposure times (Ivanina et al., 2008; O’Donovan et al., 20

Journal Pre-proof 2018; Verlecar et al., 2008). These factors can influence biomarker responses so drastically as to obscure the effects of experimental treatments (González-Fernández et al., 2017). Given this, it is possible that modifying exposure time or selecting for a different age class of mussels could have yielded statistically significant results in our experiment. Statistical insignificance aside, the trends we observed in biomarker responses point to potential oxidative mechanisms underlying sublethal responses. The declining CAT activity across the

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treatments supports other non-significant CAT reductions reported for M. galloprovincialis following

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microplastic ingestion (Avio et al., 2015). This response may be widespread among taxa, as plastic

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leachates have also reduced CAT activity in fish embryos (Walpitagama et al., 2019). Although CAT is upregulated to meet oxidative challenges, suppressed CAT activity can occur when environmental

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stressors generate particularly high intracellular H2O2 concentrations (Walpitagama et al., 2019).

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Since CAT is involved in acute responses to oxidative challenges, extended exposure times may

2018; Pittura et al., 2018).

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result in the variable CAT responses reported for Mytilus exposed to microplastics (Magara et al.,

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As with CAT, the activity of GST is also typically upregulated in response to waterborne toxicants

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because they conjugate xenobiotics for detoxification and elimination from the body (Fernández et al., 2012). The non-significant increase in GST activity in the leachate treatment compared to that of the control suggests that at least one organic compound in the microplastic leachate was absorbed across the gill epithelium and required detoxification. Surprisingly, the microplastic-only treatment also elicited higher GST activity than the control group; our cleaning process may have been insufficient to remove all leachable compounds from the phenol-formaldehyde microplastics, or the physical presence of the phenol-formaldehyde polymer in the digestive tract triggered the upregulation of GSTs. In addition to conjugating xenobiotics, GSTs catalyse the destruction of lipid peroxides, the precursor to malondialdehyde (Katsuhara et al., 2005; Singh et al., 2001), which is the biomarker we quantified 21

Journal Pre-proof as a proxy for lipid peroxidation. This is the likely cause of the inverse correlation between GST activity and TBARS concentration observed across our treatments. The same correlation has been reported in M. gallloprovincialis exposed to other waterborne toxicants (Pinto et al., 2019). Since we cannot be certain that our measures of TBARS accurately reflects the amount of lipid peroxidation occurring during the experiment, we cannot determine whether our treatments caused oxidative stress, the state of redox imbalance favouring oxidative radicals (Sies, 1986, 1985). Such confounding factors can be circumvented in future studies by utilising other biomarkers of oxidative damage,

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such as DNA damage and protein carbonyls (as used by Magni et al., 2018), as indicators of oxidative

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stress.

We found that ingesting phenol-formaldehyde microplastics alone did not affect AChE activity,

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which contrasts with reports of microplastic-induced AChE suppression in other invertebrates

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(Ribeiro et al., 2017; Yu et al., 2018). AChE activity decreased slightly in the leachate only treatment and increased marginally in the macroplastic + leachate treatment. This resulted in significantly

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different AChE activity between these treatment groups, although neither were statistically different

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from the control. These results indicate that AChE is only somewhat affected by some component of

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the leachate, and this response is altered when microplastics are physically present in the digestive system. Bivalves restrict filtering activity in the presence of waterborne pollutants (Kraak et al., 1994; Ostroumov and Widdows, 2006; Rodrigues et al., 2013), and M. galloprovincialis potentially restricted filtering activity in response to a component of the leachate treatment, hence reducing the amount of AChE synthesised during the exposure period. The trends in biomarker responses we observed imply that phenol-formaldehyde microplastics affected antioxidant enzyme activity and lipid peroxidation levels through both leachate and the physical presence of the microplastics in the digestive tract. The physical presence of foam microplastics could generate reactive oxygen species production (ROS), which would influence these antioxidant biomarkers. This response has been observed in invertebrates that have ingestion

22

Journal Pre-proof polystyrene (Jeong et al., 2016; Paul-Pont et al., 2016). Increased ROS can also result from higher metabolic rates, which are known to increase in some invertebrates following microplastic consumption (Green et al., 2016; Rodríguez-Seijo et al., 2018; Van Cauwenberghe et al., 2015). Increased metabolic rates could be driven by additional energy required to repair the damage ingested microplastics cause to intestinal structures (Lei et al., 2018; Pedà et al., 2016; von Moos et al., 2012), or to fuel immune defences against pathogens which have invaded the body across

Conclusions

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5.

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sections of damaged gut lumen (Rowley and Powell, 2007).

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We demonstrated that phenol formaldehyde microplastics are eaten by aquatic organisms with

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filter-feeding, grazing and shredding feeding modes. These microplastics leach chemicals into the surrounding water and are more toxic to aquatic invertebrates than leachates from other plastic

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families. We also demonstrated that phenol formaldehyde microplastic leachate, and the physical

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presence of the microplastics themselves, exert separate and cumulative sublethal effects on a

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marine bivalve by influencing antioxidant enzymes.

23

Journal Pre-proof Conflict of interest The authors declare no conflicts of interest. Funding This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors. Charlene Trestrail receives an Australian Government Research Training

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Program Scholarship through RMIT University.

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Acknowledgements

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We thank Rita Feldmann for alerting us to the potential environmental impacts of floral foam, and

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for providing the foam we used in this study. We acknowledge the facilities and the scientific and technical assistance from the Australian Microscopy & Microanalysis Research Facility at the RMIT

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Microscopy & Microanalysis Facility, and the National Measurement Institute, Australia. We thank

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David Haines from Agilent Technologies for assisting us with the ATR-FTIR analysis.

24

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sediment-dwelling invertebrates. Environ. Pollut. 236, 425–431.

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Journal Pre-proof Table 1. Top HQI-ranked matches of foam ATR-FTIR spectra against compounds in polymer-related spectral libraries in the KnowItAll database. Available compound descriptions from the respective libraries are provided; dashes indicate no description was available. HQI

Compound name

Compound description

Library (manufacturer)

Regular foam

97.63

Poly(2,6methylenephenol)

-

Polymers, Hummel Industrial (Wiley)

95.26

Alkali-condensed phenolformaldehyde resin

-

Polymers, Hummel Industrial (Wiley)

95.24

Supraplast-Harz 5

Alkali-condensed phenolformaldehyde resin

Polymers, Hummel

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GP-217

Phenolic resin

Poly(2,6methylenephenol)

95.15

Resi-Lam GP-117

95.08

-

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96.56

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Biofoam

Phenolic resin

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94.96

Durite AL-5390

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94.97

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Foam

Supraplast-Harz 5

Resocinol phenolformaldehyde resin

(Bio-Rad) Polymers & Monomers, Comprehensive (Bio-Rad) Polymers & Monomers, Comprehensive (Bio-Rad) Polymers, Hummel Industrial (Wiley) Polymers & Monomers, Comprehensive (Bio-Rad)

Alkali-condensed phenol-formaldehyde resin

Polymers, Hummel (Bio-Rad)

95.03

Alkali-condensed phenolformaldehyde resin

-

Polymers, Hummel Industrial (Wiley)

94.94

GP-217

Phenolic resin

Polymers & Monomers, Comprehensive (Bio-Rad)

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Journal Pre-proof Table 2. Percentage of tested aquatic invertebrates that ingested phenol-formaldehyde microplastics generated from regular foam and biofoam. All differences in ingestion were non-significant (p > 0.05). Taxonomy

Habitat

Daphnia magna (adults)

Crustacea (Branchiopoda)

Freshwater 100

100

Daphnia magna (neonates)

Crustacea (Branchiopoda)

Freshwater 90

100

Physa acuta

Mollusca (Gastropoda)

Freshwater 100

Bembicium nanum

Mollusca (Gastropoda)

Marine

Mytilus galloprovincialis

Mollusca

Marine

Artemia sp (nauplii)

Crustacea (Branchiopoda)

Allorchestes compressa

Crustacea

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100

100

90

100

90

40

70

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90

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Marine

Marine

100

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n = 10

(Bivalvia)

Ingestion of biofoam microplasticsa (%)

(Malacostraca)

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a

Ingestion of regular foam micropalsticsa (%)

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Organism

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Journal Pre-proof Table 3. Lethal concentrations (LC50) of leachates from regular foam microplastics and biofoam microplastic to aquatic organisms. LC50 values are presented as mg microplastics mL-1, brackets represent 95% confidence intervals.

Exposure time (h)

Artemia sp.

24

27.4 (21.9 – 35.2)

D. magna

24

36.2 (32.3 – 40.1)

D. magna

48

17.8 (14.2 – 21.7)

15.3 (10.0 – 20.7)

D. rerio

96

27.1 (24.2 – 29.8)

22.8 (18.0 – 29.2)

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26.9 (22.9 – 31.1)

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43.8 (40.9 – 46.8)

Biofoam microplastic leachate

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Organism

Regular foam microplastic leachate

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LC50 (mg mL-1)

44

Journal Pre-proof

Fig 1. SEM images of regular foam (A & B) and biofoam (C & D) showing the network of open cells. Both foams were comprised of resin-rich struts and thin membranes covering the open cells. The cell walls of regular foam have thicker, more numerous attachments to the surrounding struts (panel B, red arrows) compared with the cell walls of biofoam, which have thinner and fewer attachment points (panel D, yellow arrows). Fig 2. Secondary microplastics from regular foam (A) and biofoam (B) formed from the resin-rich struts (black arrows) and the cell walls (red arrows) of the foams. Scale bar is 200 µm. Fig 3. ATR-FTIR spectra of regular foam (top) and biofoam (bottom). Biofoam spectrum is offset on the y-axis to aid interpretation.

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Fig 4. Green, regular foam microplastics were ingested by freshwater and marine invertebrates with a range of different feeding modes. Control adult D. magna (A) had empty digestive tracts, whilst adult and neonate D. magna exposed to foam microplastics (B & C, respectively) showed green microplastics throughout their digestive tracts. Similarly, control Artemia nauplii (D) had empty digestive tracts, whilst Artemia nauplii exposed to foam microplastics (E) showed boli of green microplastics throughout the digestive tract. Microplastics were observed embedded in the faecal pellets of the gastropods P. acuta (F) and B. nanum (G), the bivalve M. galloprovincialis (H), and in squash preparations of faeces from the amphipod A. compressa (I). Scale bar is 250 µm.

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Fig 5. Concentration of phenolic compounds in leachate generated from regular foam microplastics (open circles) and biofoam microplastics (closed circles) over 96 h. Data are mean ± S.E (n = 3). Fig 6. Specific enzymatic activity of CAT (A), GST (B), AChE (C) and concentrations of TBARS (D). * indicates significant differences, p < 0.05. Data are mean ± SE (n=6).

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Journal Pre-proof Declaration of interests

☒ The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

☐The authors declare the following financial interests/personal relationships which may be

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considered as potential competing interests:

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Journal Pre-proof Highlights Phenol-formaldehyde microplastics are eaten by marine and freshwater invertebrates.

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These microplastics leach phenolic compounds that are toxic to aquatic animals.

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Animals respond to both the leachate and physical effects of these microplastics.

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Ingesting phenol-formaldehyde microplastics alters antioxidant enzyme biomarkers.

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