Microbeads – A Science Summary

July 2015

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Table of Contents


Microbeads are synthetic polymer particles that, at the time of their manufacture, are greater than 0.1 µm and less than or equal to 5 mm in size, which can vary in chemical composition, size, shape, density, and function. Microbeads are manufactured for specific purposes, including for use in personal care products (such as scrubs, bath products, facial cleaners, toothpastes). They may also be used in other consumer uses including cleaning products and printer toners and in industrial products such as abrasive media (e.g., plastic blasting), industry (e.g., oil and gas exploration, textile printing, and automotive molding), other plastic products (anti-slip, anti-blocking applications) and medical applications.

Microbeads from ‘down the drain’ products will likely be released into the aquatic environment after wastewater treatment. Studies have shown that microplastics, including microbeads, are present in the environment and that they can reside in the environment for a long time. Microbeads have been shown to elicit both short and long-term effects in laboratory organisms.

Proposed conclusion

Based on the available information, it is recommended that microbeads be considered toxic under subsection 64(a) of the Act.  This would enable appropriate preventative measures to be taken to reduce the release of microbeads into the environment. As a precautionary next step, the Government of Canada is proposing to add microbeads to the List of Toxic Substances under the Canadian Environmental Protection Act, 1999 (CEPA 1999).

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1. Introduction

Plastic use continues to increase globally at a significant rate. Global plastic production has increased by 620% since 1975 and was estimated to be 288 million metric tonnes in 2012 (Jambeck et al., 2015).  Due to long residence times in the environment (Andrady, 2011) and poor waste management practices, the environmental burden from plastic litter continues to increase globally (See Figure 1 below), posing environmental, economic and aesthetic issues with complex challenges and impacts (UNEP-IETC, 2012; Jeftic et al., 2009).

Plastic waste entering water and marine ecosystems can come from various sources, the majority of which originate from land-based activities (GESAMP, 2015). Shoreline recreational activities, inadequate waste management and sewer infrastructure, additives in products, and uncontrolled releases from industrial and commercial activities have been cited as major causes of plastic pollution in the marine environment worldwide.  These various sources can generate different types of plastics in the environment, from plastic bags and bottles to microplastics and microbeads.

Figure 1 (See long description below)

Figure 1: Projected releases of plastics (expressed in terms of cumulative amounts) into the marine environment globally resulting from mismanaged plastic waste (high, 40%; mid, 20%; low, 15%) (Jambeck et al., 2015)

Long description for figure 1

A graph displaying data that shows the release of plastics into the marine environment is expected to increase globally due to the mismanagement of plastic waste.

Characterizing plastics and their potential effects on the environment is complex. Microbeads are manufactured with different sizes and shapes and are not comprised of a single chemical composition but a variety of compositions, the most common of which are polyethylene, polyethylene terephthalate, polypropylene, polyamide, polyesters, polystyrene, and polyvinyl chloride. They may contain residual chemicals from manufacture and pollutants adsorbed during different life-cycle stages (e.g., plasticizers, co-contaminants, etc.) (GESAMP, 2015; Browne et al., 2011; Eriksen et al., 2014).  Once in the environment, plastics remain there for many years; for example, polyethylene and polypropylene added to the Bay of Bengal (a marine environment) to measure microbe-mediated biodegradation underwent less than 3% degradation after 6 months  (Andrady, 2011). However, over time breakdown of plastics in the environment occurs through a variety of physical and chemical processes, such as weathering (e.g., wind and water erosion), hydrolysis with water, biodegradation,  and photodegradation resulting in larger particles breaking down into smaller particles (Eriksen et al., 2014; Andrady, 2011).   

1.1 Definitions

Smaller particles of plastics are broadly referred to as microplastics. The term was originally used to differentiate between larger plastics (macro) and those which can only be visualized with a microscope (micro). There is no agreed upon definition as to what constitutes a microplastic. Researchers have used definitions that are largely based on the sampling method used to characterize the microplastic they are investigating. For example, some researchers have used 500 µm and 67 µm sieves as the upper and lower limit for microplastics sampling, while others use less than 5 mm to 333 µm as the upper and lower limits based on the neuston nets used for their sampling (Andrady, 2011). More recently, a report from the United Nations’ Joint Group of Experts on the Scientific Aspects of Marine Environmental Protection (GESAMP, 2015) recommended a less than 5 mm to 1 nm definition (1 nanometer is one billionth of a meter). In addition, regulatory definitions for microbeads also vary; for example, the State of Illinois in the United States of America (USA) only provides an upper bound limit of less than 5 mm  (See State of Illinois Public Act 98-0638), while the Canadian province of Ontario is considering microbeads with an upper bound size limit of less than 1 mm together with targeted uses in their proposed legislation (Bill 75, Microbead Elimination and Monitoring Act, 2015).

Microplastics are organized according to their source, i.e. whether they are manufactured on the micrometer size or are the result of breakdown processes discussed above (such as weathering, photodegradation, etc.) (GESAMP, 2015).

For the purposes of this summary:

The less than or equal to 5 mm cut-off is based on the upper bound limit used in research and by other jurisdictions and is indicative of expert opinion from a workshop on marine debris held in 2008 for secondary microplastics (Arthur et al., 2009). The lower bound of 0.1 µm was intentionally selected to remove nanoscale materials (those within 1-100nm). This cut-off was used: (1) to focus on industrially relevant microbeads; and (2) to differentiate between effects and properties unique to substances on the nanometer scale.

This Science Summary Report focuses on microbeads and provides recommendations on microbeads only. For the purposes of this Report, information identified up to June 2015 was considered for inclusion in this Science Summary. Due to physical-chemical similarities, when studies specific to microbeads were not available, information on secondary microplastics was used as surrogate information.

A review of scientific literature did not identify studies that indicated concerns for human health related to the presence of microbeads in personal care products.  It is expected that microbeads present in personal care products applied to the skin are not absorbed by the body but rather rinsed off or leave the body when epidermal cells are sloughed off, and ultimately released to the environment (UNEP, 2015; Leslie, 2014; SNY, 2014). Although potential effects to human health through consumption of seafood containing microbeads has been flagged by some members of the public as a concern (UNEP, 2015) the limited information on this source of exposure does not indicate a basis for review of potential risk to human health from exposure to microbeads. Accordingly, the scope of this review is limited solely to environmental impacts.

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2. Substance Identity, Properties and Uses

2.1 Substance Identity

Microbeads are synthetic polymer particles manufactured for a specific purpose and application in the size range of greater than 0.1 µm – less than or equal to 5 mm.  They can be composed of a variety of synthetic polymers depending on the required functionality. Table 1 lists the function of typical polymeric particulates found in personal care and cosmetic products (Leslie, 2014). In the case of microbeads, the most common polymers used are polyethylene, poly(methyl methacrylate), polytetrafluoroethylene, polypropylene, Nylon, and polyethylene terephthalate (Norwegian Environment Agency, 2014). Typical polymer forming reactions used to synthesize microbeads are based on the desired particle size (Jinhua & Guangyuan, 2014) and include emulsion polymerization (Chern, 2006; Asua, 2004), suspension polymerization (Brooks et al., 2010; Dowding & Vincent, 2000), and dispersion polymerization (He et al., 2011; DeSimone et al., 1994; Li & Armes, 2010). In addition, microbeads also contain residual chemicals as a result of their synthesis, such as unreacted monomers/reactants, petroleum-based chemicals, etc. These residual chemicals are different than environmental pollutants which adsorb onto the particle during its various life-cycle stages, especially within the aquatic environment (Mato et al., 2001; Teuten et al., 2007).

Table 1: Polymer compositions and corresponding functional properties for typical particulates found in personal care and cosmetic products (PCCP) (Leslie, 2014). Polyethylene, poly(methyl methacrylate), polytetrafluoroethylene, polypropylene, nylon, and polyethylene terephthalate are most typically associated with microbeads (Norwegian Environment Agency, 2014)
Polymer NameFunctions in PCCP Formulations
Nylon-12 (polyamide-12)Bulking, viscosity controlling, opacifying (e.g. wrinkle creams)
Nylon-6Bulking agent, viscosity controlling
Poly(butylene terephthalate)Film formation, viscosity controlling
Poly(ethylene isoterephthalate)Bulking agent
Poly(ethylene terephthalate)Adhesive, film formation, hair fixative; viscosity controlling, aesthetic agent, (e.g. glitters in bubble bath, makeup)
Poly (methyl methylacrylate)Sorbent for delivery of active ingredients
Poly(pentaerythrityl terephthalate)Film formation
Poly(propylene terephthalate)Emulsion stabilising, skin conditioning
PolyethyleneAbrasive, film forming, viscosity controlling, binder for powders
PolypropyleneBulking agent, viscosity increasing agent
PolystyreneFilm formation
Polytetrafluoroethylene (Teflon)Bulking agent, slip modifier, binding agent, skin conditioner
PolyurethaneFilm formation (e.g. facial masks, sunscreen, mascara)
PolyacrylateViscosity controlling
Acrylates copolymerBinder, hair fixative, film formation, suspending agent
Allyl stearate/vinyl acetate copolymersFilm formation, hair fixative
Ethylene/propylene/styrene copolymerViscosity controlling
Ethylene/methylacrylate copolymerFilm formation
Ethylene/acrylate copolymerFilm formation in waterproof sunscreen, gellant (e.g. lipstick, stick products, hand creams)
Butylene/ethylene/styrene copolymerViscosity controlling
Styrene acrylates copolymerAesthetic, coloured microspheres (e.g. makeup)
Trimethylsiloxysilicate (silicone resin)Film formation (e.g. colour cosmetics, skin care, suncare)

2.2 Properties

Microbeads can vary in size, shape and density based on the chemical composition and method of synthesis (Napper & Thompson, 2015 in press). As can be seen from Table 2 (Hidalgo-Ruz et al., 2012), polymer particles (which include microbeads) can range in polymer densities from 0.9-2.10 g/cm3 (density of water at 25°C is approximately 1 g/cm3). In addition to polymer densities, the density of the entire particle will also be a function of other chemicals added during its manufacture (e.g., additives, fillers, etc). This variation in densities means that some synthetic polymer particles (including microbeads) will float on water surfaces and others may be present in the water column or settle to the sediments. Once in the environment, this behaviour will change depending on the aggregation/dis-aggregation and agglomeration/dis-agglomeration behaviour as the microbeads interact with environmental media, e.g., humic/fulvic acids. Moreover, synthetic particles (e.g., plastics) may become fouled by organisms and as a consequence, particles that initially floated may eventually sink to the sea bed. For example, substantial quantities of microplastics have been reported in deep sea sediments (Woodall et al., 2014).

Table 2: Examples of plastic of different polymer compositions and relative densities (Hidalgo-Ruz et al., 2012)Footnote[a]
Polymer TypePolymer Density (g/cm-3)
polyamide (nylon)1.02-1.05
polyvinyl alcohol1.19-1.31
poly methylacrylate1.17-1.20
polyethylene terephthalate1.37-1.45


Footnote a

Data from a total of N = 42 studies.

Return to footnote[a]referrer

Due to the desired functionality of microbeads in a variety of personal care products, they can either be chemically and/or physically stable (e.g., when used as abrasives) or unstable (e.g., when designed to breakdown due to physical or chemical trigger to release other chemicals). Stable microbeads are most likely to persist in the environment.

Figure 4 below shows examples of microbeads found in cosmetic products. The aggregation and/or agglomeration of the microbeads is apparent from the micrographs; however its relevance to environmental fate and effects is unknown.

Figure 2 (See long description below)

Figure 2:  (images - top) Shapes of polyethylene microbeads from four different facial cleansers available in New Zealand (A1-A4). Two of the four cleansers contained additional spherical microbeads (shown in A5 and A6, respectively) with unknown chemical composition; (graphs – B1-B4) the size distribution of microbeads in the tested cleansers (adapted from Fendall & Sewell, 2009)

Long description for figure 2

A figure that shows several images of plastic beads found in store-bought facial cleansers from New Zealand. The images were taken by a compound microscope and the plastic particles range in size from four micrometers to twelve hundred micrometers in diameter. Four graphs that display the size distribution of plastic particles from each of four store-bought facial cleansers.

2.3 Uses

Globally, microbeads have been found to have use in personal care products, other consumer applications, and various industrial applications.

Based on information presented in scientific literature considering personal care products, microbeads have been found in scrubs/peelings, shower/bath products, facial cleaners, creams, deodorants, make-up foundations, nail polishes, eye colours, shaving creams, bubble baths, hair colourings, insect repellants, toothpaste, eye shadows, blush powders, hairsprays, liquid makeups, mascaras, baby products, lotions, and sunscreens. Microbeads may also be found in other consumer uses/products including cleaning products and printer toner (Norwegian Environment Agency, 2014). Some products contain substantial quantities of microbeads. For example, Napper and Thompson (2015, in press) quantified microbeads incorporated in personal care products as exfoliants and showed that abundance varied considerably among products (137,000 – 2,800,000 per 150ml bottle). Some products that are used on a daily basis could result in release to household waste water of 94,500 microbead particles per application (Napper & Thompson, 2015 in press).

In 2015, the Canadian Cosmetic, Toiletry, and Fragrance Association (CCTFA) voluntarily surveyed its members and shared summarized information with the Government of Canada. CCTFA information indicates that in Canada, microbeads were reported to be used in personal care product categories of skin care (which include anti-aging creams, moisturizers, cleansers, etc.), bath and body (which include bath/shower gels or soaps, lotions, talcs or balms, nail polishes, etc.), and cosmetic-like products, which include fluoridated toothpastes, acne therapy, etc. While the specific products were not reported, the total annual volumes of microbeads in Canada by individual CCTFA members ranged from 30kg/year to 68,000 kg/year.

Microbeads are also used in industrial products such as abrasive media (e.g., plastic blasting at shipyards, productions facilities such as garment and car parts), industry (e.g., oil and gas exploration, textile printing, and automotive molding), other plastics products (e.g., anti-slip and anti-blocking applications) and medical applications (biotechnology and biomedical research) (Leslie, 2014; Norwegian Environment Agency, 2014).

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3. Environmental Fate

When used in personal care products, microbeads enter the environment primarily through effluent from wastewater treatment plants from ‘down the drain’ release of products.  Secondary routes of entry into the environment include accidental spills and releases related to industrial applications (GESAMP, 2015).

In a recent study by Talvitie and Heinonen (2014), a preliminary investigation on microplastics removal from the Central Wastewater Treatment Plant in St. Petersburg, Russia suggested that although treatment showed a high removal of microplastics from wastewater effluent (greater than 95 %) after secondary treatment, a number of particles do remain in the effluent and enter the aquatic compartment.  The authors filtered purified effluent water with 300, 100, and 20 µm filters and identified fibers and particles as the primary microplastics in the incoming wastewater. These findings are consistent with a recent study in Paris, France by Dris et al., (2015) who found greater than 90% removal of microplastics after wastewater treatment. Specific to microbeads, New York State recently investigated a number of their wastewater treatment plants (WWTPs) and found that microbeads were present in the effluent of 25 of the 34 WWTPs sampled (New York, 2015). Thus, while additional studies are needed for microbead removal and transformations from wastewater treatment processes, microbeads are expected to be removed to a high degree but also will pass through WWTPs and enter the aquatic environment.

Once in the aquatic compartment, the subsequent behaviour of microbeads depends on their physical-chemical properties.  Microbeads, like other particles (Buchs et al., 2013; Dale et al., 2015; Syberg et al., 2015), will either interact with chemicals in the water column (e.g., sorb natural organic matter) and/or settle to the sediment. Figure 3 describes the processes that microbeads undergo after introduction to the aquatic environment. Once exposed, microbeads can undergo physical transformations (from mechanical degradation, weathering, etc.) and adsorb/desorb a variety of local pollutants (e.g., persistent organic pollutants such as polyaromatic hydrocarbons, polychlorinated biphenyls, etc.) from the surrounding environmental medium (Bakin et al., 2014; Teuten et al., 2009). Recent work has shown that microbeads extracted from cosmetic products have similar potential to adsorb persistent organic pollutants as reported for microplastic particles (Napper & Thompson, 2015 in press). In addition, interactions of microbeads with natural organic matter will have a strong impact on where they will finally reside in the water column. Microbeads with low densities, unless perturbed (e.g., by interacting with dissolved organic matter, other particulates, or micro-organisms), will float and be available to pelagic and avian species, while the denser microbeads in the water column are expected to settle over time. The denser microbeads will then undergo transformations (e.g. agglomerating/aggregating, increasing in size and mass after interacting with dissolved chemical species) and become available to aquatic benthic species. Therefore, microbeads are expected to be present in both the water and sediment compartments.

Figure 3 (See long description below)

Figure 3: Environmental fate and behaviour of plastic particles after release­ to the aquatic environment.  While the figure focuses on secondary microplastics (i.e., the breakdown of larger plastic litter), the behaviour applies to microbeads as well. Note that microbeads can float due to the lower relative density and/or interact with dissolved/dispersed chemicals and eventually partition to sediments. Microbeads can come into contact with organisms at any stage (Leslie, 2014)

Long description for figure 3

A diagram showing possible entry routes of microplastics entering a body of water and probable destinations once introduced. The diagram is presented in a flow-chart format, beginning with vectors of microplastic input into the aquatic environment. It is shown that microplastics may be introduced to the aquatic environment through transport via river bodies, wind, or ships. The flow chart then presents biota-related, chemical-related, and physical process-related effects that microplastics will undergo upon entry into the water bodies.

The figure shows that there is often a contaminate-rich microlayer at the surface of general bodies of water, and here floating microplastics are expected to encounter and adsorb any persistent organic pollutants that may be present. Chemicals dissolved in the water will also act to adsorb and concentrate onto plastic that is present. Further degradation by exposure to wave action and ultra-violet light will act to break up these plastics into ever smaller particles, consequently increasing their overall surface area and thus their ability to adsorb additional pollutants. At this stage, microplastics may also interact with algae on the surface or common aquatic life in the water column and uptake of the particles may follow.

Over time, weathering, biofouling, and sinking of the suspended particles is shown to occur. Additionally, epipelagic organisms that ingest microplastics may subsequently be consumed by predators, potentially initiating food-chain transfer of the particles. Chemical additives used in the manufacture of the plastic will proceed to leach into the surrounding environment; which is depicted to either be the water body itself or potentially the G-I tract of an organism that has consumed such microplastic particles.

The flow chart shows that the long term bioavailability of the particles and their contaminants depends on their ultimate fate. Low density particles will likely remain on the surface and continue to degrade, adsorb pollutants and potentially become ingested by aquatic organisms or birds. High density particles will sink to the sediment and the diagram notes that this will provide a low bioavailability to microbes. There would also likely be a re-suspension of chemicals into the surrounding water via the plastic present in the sediment. And there is also potential for the bioaccumulation of any persistent organic pollutants taken up with the ingestion of plastics in the food chain.

There is very little known about the fate of microbeads (and secondary microplastics) in air. It is unknown whether microbeads, like other particulates (Quadros & Marr, 2010; Hennigan et al. 2011) with low relative densities can partition to the air compartment and, if they do, are they able to adsorb airborne pollutants and/or undergo long-range transport and atmospheric transformations (e.g., reactions with hydroxyl radicals) (Dellinger et al., 2001). In addition, fate in soil is also unknown for microbeads (and secondary microplastics). Based on one study at a municipal wastewater plant in Russia, microplastics, although not completely removed from the effluent, are expected to primarily partition to biosludge after wastewater treatment (Talvitie & Heinonen, 2014). Once in biosolids, microplastics have the potential to be present in soils should the biosolids be applied to land.  Once in the soil, microbeads (Darlington et al., 2009) could be mobile (although this is expected to be unlikely) or immobile depending on the soil chemistry and the size of the microbead relative to the soil particulates (Bradford et al., 2002). 

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4. Environmental Presence of Microplastics

In the environment it is extremely difficult to differentiate and discriminate between microbeads and secondary microplastics. As most studies report only total microplastic concentrations, it is not currently possible to quantify the contribution of microbeads versus all other plastic litter. The only study available on microbead contribution to plastic litter is by Gouin and colleagues (2011), who have conservatively estimated that the use of polyethylene microbeads in liquid soap alone resulted in the consumption of 2.4mg of polyethylene microbead per person per day, thereby emitting a total of 263 tonnes per year of polyethylene microbeads in the United States from liquid soap use. Due to the lack of data explicitly regarding microbeads, information on microplastics (which includes both microbeads and secondary microplastics) was used to highlight the presence of microbeads in the environment.

Microplastics have been measured at almost every location on the globe, including waters, sediments, soils (Hall et al., 2015), deep sea sediment deposits (Woodall et al., 2014) and ice cores (Obbard et al., 2014). Figure 4 indicates concentrations of microplastics in sediment and surface waters from different regions. Several authors have suggested there will be a steady increase of marine litter (which includes microplastics) in the environment (Jambeck et al., 2015) over the next few decades (Figure 1). Law and Thompson (2014) have noted that even with the prevention of additional macroplastics into the environment, concentrations of microplastics will continue to rise due to fragmentation of larger plastics into smaller particulates (Law & Thompson, 2014).

Figure 4 (See long description below)

Figure 4: (A) Global distribution of microplastics in sediments from 18 sandy shores from around the world (Browne et al., 2011); and (B) average concentrations of plastics (primarily microplastics) in surface waters (Cózar et al., 2014)

Long description for figure 4

(A) A simplistic black and white map of the world with green circles of various sizes plotted on the locations of 18 different beaches around the globe. The sizes of the circles correspond to the number of microplastic particles found there per two hundred and fifty milliliters of sediment. The information relating the size of the circles to the respective microplastic concentrations that they represent is shown in a legend to left of the map. The legend presents four different circle sizes that reflect ranges of concentrations that are equal in magnitude. It shows that the smallest circles on the map represent the range of 1 to 10 microplastic particles per two hundred and fifty milliliters of sediment. The next three size ranges are 11 to 20, 21 to 30, and 31 to 40 microplastic particles per two hundred and fifty milliliters of sediment respectively. For the purposes of describing the plotted data, four tiers will be used to represent the plastic concentrations from smallest to largest and will be called levels 1, 2, 3 and 4. The shores that were sampled in this study and their associated data are given as follows.
Port Douglas, Australia, level 1;
Busselton Beach, Australia, level 3;
Kyushu, Japan, level 3;
Dubai, United Arab Emirates, level 1;
Vina Del Mar, Chile, level 2;
Punta Arenas, Chile, level 2;
Malapascua Island, Philippines, level 1;
Fara, Portugal, level 4;
Ponta Delgado, Azores, level 3;
Two beaches in Virginia, USA, level 3;
California, USA, level 1;
Plymouth, USA, level 1;
Western Cape, South Africa, level 3;
Pemba, Mozambique, level 3;
Sennon Cove, United Kingdom, level 3;
River Tyne, United Kingdom, level 2;
and an unspecified beach in Oman, level 2..

(B) A black and white map of the world showing concentrations of plastic debris measured in surface waters of the world's oceans. The data that is plotted on the map is done so via coloured circles of which their color and size indicate mass concentrations of plastic in grams per square kilometer. A legend in the top right corner of the image shows six ranges of concentrations and a corresponding coloured circle that represents each concentration. At zero plastic concentrations a small colourless circle with a blue border is used. From 0 to 50 grams per square kilometer a small blue circle is used. From 50 to 200 grams per square kilometer, a small green circle is used. From 200 to 500 grams per square kilometer, a small yellow circle is used. From 500 to 1000 grams per square kilometer, a medium sized red circle is used. And from 1000 to 2500 grams per square kilometer, a large, dark red circle is used. The data shows average concentrations in 442 sites around the world, and was collected via 1,127 total surface net tows.

According to the image relatively low average plastic concentrations of 0 to 50 grams per square kilometer labelled with small blue circles are present near virtually all global shorelines. The data tends to show however that high concentrations of debris appear to gather in large accumulation zones in the middle of the oceans. These areas are marked on the image as patches of light and dark grey, indicating the outer and inner accumulation zones respectively. These zones in this image are produced from a global surface circulation model, and they correspond with the world’s primary oceanic gyres. These gyres include the North and South Atlantic gyres, the North and South Pacific gyres and the Indian Ocean gyre..

4.1 Presence in Canada

Microplastics have been measured in Canadian waters and sediments. Desforges et al. (2014) found microplastic (ca. 70% microfibers and 30% pellets ranging in size from 64.8 μm to 5810 μm) concentrations ranging from 8 to 9200 particles/m3in sub-surface seawaters of the northeastern Pacific Ocean and coastal British Columbia. Microplastics have also been measured, primarily as fibers, in Nova Scotia beach sediment at concentrations of 20-80 microplastics/10g sediment (Mathalon & Hill, 2014).  In another study by Obbard et al. (2014) microplastic concentrations of 30-234 particles/m3 of ice were found from ice samples during two Arctic expeditions in the Beaufort and Chukchi Seas. In their study, Obbard and colleagues (2014) identified microplastics of rayon (54%), polyester (21%), nylon (16%), polypropylene (3%), polystyrene (2%), acrylic (2%), and polyethylene (2%) with sizes ranging from 0.02 mm to 2 mm. Microplastics have also been measured in freshwater systems, for example in the St. Lawrence River microplastic median concentrations were 52 microplastic/m2 (primarily polyethylene with sizes ranging from 0.4 to 2.16 mm) after sampling across 10 freshwater sites (Castañeda et al., 2014). Similarly in Lake Superior, Lake Huron and Lake Erie average abundance of microplastics was found to be 43,157 particles/km2 with sizes ranging from 0.355 mm to greater than 4.75 mm (81% of the microplastics were in the 0.355-0.999 mm fraction) (see Figure 5 below for distribution) (Eriksen et al., 2013). The differences in units used to report microplastics concentrations in the environment are primarily due to the different methods used to sample microplastics (Andrady, 2011) and lack of adequate quality control (e.g., lacking standard reference materials and proficiency testing). These differences limit the comparability of values across different studies.

Plastics, including microplastics, have also been measured on Canadian beaches of Lake Huron (at concentrations of 38 particles/m2) (Zbyszewski & Corcoran, 2011), Lake Erie (ranging from 0.36-1.78 pieces/m2), and Lake St. Clair (ranging from 0.18-8.38 pieces/m2) (Zbyszewski et al., 2014). In their study, the authors mainly found microplastics composed of polyethylene and polypropylene sorted in size fractions of less than 5 mm and greater than 5 mm. Recently, plastic debris (primarily microplastics) have also been measured on beaches of Humber Bay at concentrations of 16.3 pieces/m2fractionated by less than 1cm (55 fragments/48g total mass), 1-5cm (321 fragments/122.90g total mass), and greater than 5cm (29 fragments/47.60g total mass) (Corcoran et al., 2015). Thus, it is evident that microplastics are present across Canada in freshwater and saltwater ecosystems, and as evidenced by Figure 5 below, microbeads are also present together with microplastics (suggesting similar fate and behavior).

Figure 5 (See long description below)

Figure 5: (A) Distribution of microplastics by count from 21 samples collected in the Laurentian Great Lakes; and (B) microbead found at one sampling site (Eriksen et al., 2013)

Long description for figure 5

(A) A map showing the distribution of microplastics in the Great Lakes. This map includes data for Lake Erie, Lake Superior, Lake Michigan and Lake Huron, but not Lake Ontario. From this data set, the highest numbers of microplastics were found in Lake Erie.

(B) Scanning electron microscopy image of a spherical plastic fragment found in one of the Great Lakes.

4.2 Accumulation in the Environment

Microplastics are accumulating in the environment. In a recent paper by Obbard and colleagues (2014) microplastics were measured in frozen ice cores of the Arctic Ocean in 2010. The authors confirmed that these microplastics have accumulated far from population centers and suggested that polar sea ice is becoming a major sink for microplastic contamination and, as the ice melts, these microplastics can be released into the environment.  In addition, a recent study by Corcoran and colleagues (2015) have found that microplastics have been accumulating in sediment cores of Lake Ontario (10.5 pieces/m2) for the past 38 years.  Similarly, in work by Thompson and colleagues (2004), plastic fibers have been measured in archived plankton samples dating as far as the 1960s and trend data indicates a significant increase in plastic fragment abundance over time (primarily fibers of approximately 20μm in diameter), further confirming that the accumulation of microplastics is increasing in places in the environment (Thompson et al., 2004).

Figure 6 (See long description below)

Figure 6: (A) Images of microplastics (at arrows) found in ice cores from the Arctic Ocean (Obbard et al., 2014); and (B) images of microplastics found from sediment cores of Lake Ontario (Corcoran et al., 2015). Scale bars on the left are 1 mm

Long description for figure 6

(A) Images of ice core samples from the Arctic Ocean that were found to contain microplastics.

(B) Three images of millimeter sized plastic particles found in sediment cores of Lake Ontario.

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5. Effects in Organisms from Microbeads

At the time of this report, over 130 publications on fate and effects of microplastics were reviewed. Key studies only on microbeads have been summarized in the Tables 3 and 4 below. The scope of this summary is not to present all of the effects data on microbeads; but rather to demonstrate different types of effects possible from exposure to microbeads in the environment. In addition to measureable effects, there are multiple studies on microbeads which noted no evidence of adverse effects on aquatic organisms after exposure, such as the study by Kaposi and colleagues (2014) where the survival of sea urchin larvae was not impacted after 5 days of microbead exposure. Where information on microbeads specifically was not available surrogate information on microplastics was used.  No studies measuring acute lethality/effective concentrations (LCx/ECx) for microbeads (i.e., traditional endpoints used in chemical toxicity assessments) was found and only one very recent study which calculated chronic LC50 concentrations in daphnia. From the scientific literature, the effects seen are either primarily driven by physical effects (i.e., effects resulting from blockages, external/internal attachment, etc.) and/or due to the presence of residual chemicals (those chemicals which are present when the microbeads are synthesized) and/or adsorbed pollutants (e.g., persistent organic pollutants (POPs), pesticides, etc. which are adsorbed in later life-cycle stages). Where physical effects are the primary driver for effects, no large differences were seen between freshwater and marine organisms. Current research lacks clarity on whether effects observed are from particulate matter of the plastic (e.g., polyethylene) or from residual chemicals from plastic manufacturing (such as unreacted monomers and petroleum-based chemicals) or from adsorbed pollutants.  It is important to note that many studies use high concentrations of microbeads relative to environmental levels of microplastics (measured concentrations of only microbeads are lacking); while some studies expose organisms in the absence of food. The types of effects are summarized below:

Table 3: Summary findings of microbeads effects in freshwater organisms
OrganismMicrobead type and concentrationSummary FindingsSource
  • Lumbriculus variegatus (California blackworm)
  • Daphnia magna (crustacean)
  • Potamopyrgus antipodarum (New Zealand mud snail)
  • Gammarus pulex (amphipod crustacean)
  • Notodromas monacha (amphipod crustacean)
Red, non-floating fluorescent polymethyl methacrylate microbeads Mean particle size: 29.5±26μm
Specimens fed either a 10:1 or 1:1 food:plastic ratio
  • Fluorescently tagged microbeads were measured in the gut of the exposed organisms (ranging from 32.4±3.8% in Notodromas monacha to 93 ± 0.07% in Lumbriculus variegatus), in intestines of Daphnia magna, and in the faeces (up to 96 ± 0.03% in Gammarus pulex) thereby indicating active uptake of microbeads into organisms.
Imhof et al., 2013
  • Daphnia magna
Fluorescently labeled 1μm carboxylated polystyrene microbeads at 2μg/L (nominal) concentration
  • Fluorescently labeled microbeads accumulated in the GI tract (within 60 minutes) reaching a maximum concentration of ca. 700 times greater than media concentration. Particles underwent relatively rapid depuration and particle concentration decreased by over 90% over 240 minutes. In addition, microbeads were also measured in the specimen’s lipid storage droplets, indicating translocation.
Rosenkranz et al., 2009
  • Hyalella azteca
Fluorescently labeled 10-27μm polyethylene microbeads and aged polypropylene marine rope 20-75μm in length and 20μm diameter at concentrations of 100,000 microbeads/mL (acute), 20,000 microbeads/mL (chronic), and 90 microplastic fibers/mL (acute fiber)
  • Acute exposures (10 day) to daphnia Hyalella Azteca  resulted in increased ingestion of microbeads with increasing microbead concentrations. In chronic exposures size selective uptake of microbeads was seen as the organisms grew, suggesting that as the organisms grow larger they may prefer larger particulate matter and ingest fewer microbeads. No translocation from the gut was measured in this study.
  • During acute exposures (10 day), polypropylene microfibers were found to be more toxic than the spherical beads decreasing growth of the organisms.
  • In chronic exposures (42 day) to polyethylene microbeads resulted in growth reductions and reproduction likely due to the reduced food intake when microbeads are present.
  • 10-d LC50s for polyethylene spherical and polypropylene fiber microbeads were 4.64 X 104 microbeads/mL and 71.43 microbeads/mL, respectively.  
Au, 2015
Table 4: Summary findings of microbeads effects in saltwater organisms
OrganismMicrobead type and concentrationSummary FindingsSource
  • Idotea emarginata (isopod)

Fluorescently-tagged polystyrene microbeads (10μm), polystyrene plastic fragments (1-100um) and polyacrylic fibers (20-2,500um).

Isopods were fed 12 or 120 microbeads/mg, 20 or 350 fragments/mg, and 0.3mg fibers per g food.

  • Isopods did not distinguish between food with and without fluorescent microplastics. Microplastics were found primarily in the gut. No aggregation was observed. 3.5 microplastic particles were detected per mg stomach tissue. less than 1 particle/mg tissue detected in midgut gland sample. There was no impact on survival, intermolt duration, and growth in the exposed isopods.
Haimer et al., 2014
  • Acartia spp.
  • Eurytemora affinis
  • Limnocalanus macrurus
  • Bosmina coregoni maritime
  • Evadne nordmannii
  • Marenzelleria spp.
  • Synchaeta spp.
  • Tintinnopsis lobiancoi
  • Neomysis integer
  • Mysis mixta
  • Mysis relicta
10μm fluorescent polystyrene microbeads at 1000, 2000, and 10,000 microspheres/mL
  • Uptake of microbeads was seen in all organisms. Food web experiments with microbead enriched zooplankton fed to mysid shrimp confirmed the presence of microbeads in mysid intestine after 3h incubation.
Setälä et al., 2014
  • Oryzias latipes (Japanese medaka “rice fish”)

Fish were exposed to two treatments of low-density polyethylene (LDPE) microbeads (less than 500μm) at 8ng/mL:

  • A virgin-plastic treatment (LDPE virgin pre-production plastic)
  • A marine-plastic treatment (LDPE deployed in an urban bay for 3 months prior to experiments)
  • Two month exposure from plain and POP-modified (PAHs, PCBs, and PBDEs) microbeads was conducted in fish. Following exposure, an increase in the concentration of PAHs, PCBs, and PBDEs was found in the fish. In addition, stress was measured in the liver of the fish in both plain and POP-modified treatments as determined by glycogen depletion (seen in 46% and 74% of the fish after unmodified and POP-modified microbead exposure, respectively), fatty vacuolation and single cell necrosis.
Rochman et al., 2013
  • Oryzias latipes (Japanese medaka “rice fish”)
less than 0.5 mm low-density  polyethylene (LDPE) microbeads: pre-production (virgin) and modified through urban bay deployment (marine) at 8ng/mL concentration
  • Two month exposure from plain and POP-modified (PAHs, PCBs, and PBDEs) microbeads was conducted in fish. Altered gene expression was observed in male fish exposed to POP-modified microbeads and gene expression was observed in female fish for both unmodified and POP-modified microbeads. Significant down regulation of choriogenin (Chg H) gene expression in males and significant down regulation of vitellogenin (Vtg I), Chg H and estrogen receptor alpha (ERα) was measured. In addition, histological observation revealed abnormal proliferation of germ cells in one male fish exposed to POP-modified microbeads, Down-regulation of Chg H in fish fed the virgin plastic treatment suggests that the plastic particles are capable of inducing an endocrine-disrupting effect.
  • Pomatoschistus microps (Common goby)
420-500 μm polyethylene microbeads (white, black, and red) at 30 particles/treatment.  Each treatment took place in 300mL of artificial sea water.
  • Differences were noted in feeding behaviour when fish from different sources were fed microbeads in combination with natural feed. In addition to being able to differentiate between microbeads and natural feed, predatory performance was significantly reduced of fish in the presence of black and red microbeads, while predatory efficiency was reduced with all microbeads indicating a direct impact of exposure to feeding ability.
Carlos de Sá et al., 2015
  • Holothuria floridana
  • Holothuria grisea
  • Cucumaria frondosa
  • Thyonella gemmata
4 mm PVC pellets (65.0g), 0.25-15 mm pipe shavings (10.0g), and 0.25-1.5 mm nylon fragments (2.0g) mixed with 600mL of sterile silica
  • Some organisms were found to ingest microbeads preferentially over sand particles. In addition, the study may indicate shape-dependant uptake of microbeads through feed.
  • Tripneustes gratilla (sea urchin larvae)
25-32 μm polyethylene microbeads at 1, 10, 100, and 300 spheres/mL
  • The survival of T. gratilla larvae was not significantly impacted by exposure to microspheres after 5 days. However, larvae that were exposed to the highest concentration had smaller body widths. Other characteristics (abnormality, body length, larval asymmetry) were not affected by exposure to microbeads.
  • Arenicola marina (lugworm)
230 μm polyvinylchloride microbeads (5 wt %)
  • Microbeads modified with environmentally relevant pollutants (nonylphenol, phenanthrene, Triclosan, and PBDE-47) transferred these pollutants into the gut tissue of lugworms, although silica (sand) was found to release a higher concentration of these pollutants into gut tissues. Effects of these transferred pollutants resulted in reduced survival, feeding, immunity, and antioxidant capacity. In addition, microbeads without added pollutants led to greater than 30% decrease in the ability to deal with oxidative stress. 
  • Arenicola marina (lugworm)
400-1300 μm polystyrene microbeads/L sediment (0, 1, 10, and 100 g)
  • Lugworm survival was 94% after 28 day exposure period. No microbeads were found in organisms which survived the entire exposure period. The worms ingested the polystyrene microbeads but these did not accumulate in the organism.
  • Mytilus edulis L. (Blue mussel)
greater than 0 – 80 μm High-density polyethylene (HDPE) microbeads
at 2.5 g HDPE/L
  • HDPE microbeads were found on gills and inside the digestive system. Transfer into the mussels was facilitated via microvilli and cilia movement.
  • Exposure to the microbeads induced a significant increase in granulocytoma formation (after 6hours) and a decrease in lysosomal membrane stability (more apparent after 96hours). Mechanism of toxicity suggested is: (1) particle ingestion (within 3 hours of exposure), granulocytoma formation (after 6 hours), and lysosomal destabilization at the cellular and subcellular level.
  • Microbeads were found in the intestine, in the lumina of primary and secondary ducts of the digestive gland, and in endocytotic vacuoles of digestive epithelial cells indicating that microbeads can be internalized into cells.
  • Mytilus galloprovincialis
less than 100 μm polyethylene (PE) and polystyrene (PS) microbeads.
Mussels were exposed separate treatments of virgin and pyrene-adsorbed beads of both plastic types at a nominal concentration of 1.5 g/L.
  • Microbeads with 200-260 ng/g of adsorbed pyrene were fed to mussels accumulated in tissues. No differences due to chemical composition were seen in the results suggesting effects were driven by physical properties of the microbeads.
  • Histological analyses of treated mussels revealed the presence of microparticles in haemolymph, gills and, especially, in digestive glands where numerous aggregates could be observed in the intestinal lumen, epithelium, and tubules. In digestive glands, concentrations of pyrene were up to 13 folds higher than the control suggesting an elevated desorption and bioconcentration process of pollutants absorbed on microbeads.
  • In addition, biomarkers indicated effects at the cellular level and sub-cellular level. Genotoxicity was measured in the mussels with DNA damage higher in organisms exposed to unmodified microbeads and nuclear alteration seen in both pyrene-modified and unmodified microbeads. 
  • Lytechinus variegatus (sea urchin)
Microbeads not described
  • Relatively pure microbeads and microbeads from a beach source (presumably microplastics) both showed adverse effects on embryos of sea urchin, leading to increased anomalous embryonic development. Adverse effects for the pure microbeads were attributed to residual chemicals from the manufacturing process (these beads were sourced from a petrochemical factory), while for the beach sourced microbeads adsorbed pollutants were expected to be the cause of adverse effects.
  • Calanus helgolandicus (copepod)
20 μm polystyrene microbeads (75 microbeads/mL)
  • Prolonged exposure (9 days) to polystyrene microbeads resulted in decreased reproductive output (possibly due to impedance of copepod feeding behaviour), but no significant differences between the control in egg production rates, respiration, or survival.
  • Tigriopus japonicas (copepod)
0.05, 0.5 and 6 μm polystyrene beads tested at concentrations of 0, 6, 13, 31, 63, 187, 250, and 313 μg/mL.
  • After 96 h, the nauplii and adult females survived when exposed to the highest concentration (313 μg/mL) of the three sizes of PS beads
  • In a two generation toxicity test, the nauplii died within approximately one week after being exposed to PS beads at greater than 12.5 μg/mL in the F0 generation and greater than 1.25 μg/mL in the F1 generation, before the metamorphosis into copepodids.
  • In the two-generation test, high concentrations of 0.5-μm PS beads were found to have caused increased toxicity and had impacts on the survival and development of copepods in the F1 generation.
  • Despite the potential for ingested PS beads to be transferred from mother to offspring, fluorescent beads were not observed in eggs while in the ovisac.
  • 0.5- and 6-μm diameter beads were found to induce a decrease in fecundity

5.2 Secondary Microplastics

Since microbeads and secondary microplastics both have similar physical-chemical properties and similar fates (i.e., long residence time), information on presence in different organisms has been summarized below for secondary microplastics and can be used as surrogate for microbeads.

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6. Uncertainties

The science on microbeads is still emerging. The following highlight some of the limitations with the current state-of-the science:

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7. Recommendations

The concentrations of microplastics are expected to increase significantly in the environment due to the expected linear increase in mismanaged macro plastic debris from increased use of plastics over the next decade (Jambeck et al., 2015) and  fragmentation of existing macro plastic debris (Law & Thompson, 2014). According to a recent study by Eriksen and colleagues (2014), there are approximately 5.25 trillion plastic particles weighing 268,940 metric tonnes currently floating at sea. Of this, microplastics are predicted to account for 92.4% of the global particle count.

Microbeads are a contributor of plastic litter in the environment. The continued use of microbeads will result in increased presence in the environment. In laboratory studies, microbeads have shown adverse short-term and long-term effects in aquatic organisms.  Microbeads may reside in the environment for a long time and continuous release of these substances to the environment may result in long term effects on biological diversity and the ecosystems. Based on the available information, it is recommended that microbeads be considered toxic under subsection 64(a) of the Act.  This would enable appropriate preventative measures to be taken to reduce the release of microbeads into the environment. As a precautionary next step, the Government of Canada is proposing to add microbeads to the List of Toxic Substances under the Canadian Environmental Protection Act, 1999.

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8. References

  1. Andrady, A. L. (2011). Microplastics in the marine environment. Marine Pollution Bulletin, 62(8), 1596-1605.
  2. Arthur, C., Baker, J., Bamford, H. (2009). Proceedings of the International Research Workshop on the Occurrence, Effects, and Fate of Microplastic Marine Debris. Department of Commerce, National Oceanic and Atmospheric Administration, Technical Memorandum NOS-OR&R-30.
  3. Asua, J. M. (2004). Emulsion polymerization: from fundamental mechanisms to process developments. Journal of Polymer Science Part A: Polymer Chemistry, 42(5), 1025-1041.
  4. Au, S. Y., Bruce, T. F., Bridges, W. C., Klaine, S. J. (2015). Responses of Hyalella azteca to acute and chronic microplastic. Environmental Toxicology and Chemistry.
  5. Avio, C. G., Gorbi, S., Milan, M., Benedetti, M., Fattorini, D., d'Errico, G., Pauletto, M., Bargelloni, L.  Regoli, F. (2015). Pollutants bioavailability and toxicological risk from microplastics to marine mussels. Environmental Pollution, 198, 211-222.
  6. Bakir, A., Rowland, S. J., Thompson, R. C. (2014). Transport of persistent organic pollutants by microplastics in estuarine conditions. Estuarine, Coastal and Shelf Science, 140, 14-21.
  7. Besseling, E., Foekema, E. M., Van Franeker, J. A., Leopold, M. F., Kühn, S., Bravo Rebolledo, E. L., Heße, E., Mielke, L., IJzer, J., Kamminga, P. Koelmans, A. A. Mar. Pollut. Bull. (2015)
  8. Besseling, E., Wegner, A., Foekema, E. M., van den Heuvel-Greve, M. J., Koelmans, A. A. (2012). Effects of microplastic on fitness and PCB bioaccumulation by the lugworm Arenicola marina (L.). Environmental science & technology, 47(1), 593-600.
  9. Bradford, S. A., Yates, S. R., Bettahar, M., Simunek, J. (2002). Physical factors affecting the transport and fate of colloids in saturated porous media. Water Resources Research, 38(12), 1327.
  10. Brooks, B. (2010). Suspension polymerization processes. Chemical Engineering & Technology, 33(11), 1737-1744.
  11. Browne, M. A., Crump, P., Niven, S. J., Teuten, E., Tonkin, A., Galloway, T., Thompson, R. (2011). Accumulation of microplastic on shorelines worldwide: sources and sinks. Environmental science & technology, 45(21), 9175-9179.
  12. Browne, M. A., Dissanayake, A., Galloway, T., Lowe, D., Thompson, R. (2008). Ingested microplastic plastic translocates to the circulatory system of the mussel, mytilus edulis. Environmental science & technology, 42, 5026-5031.
  13. Browne, M. A., Niven, S. J., Galloway, T. S., Rowland, S. J., Thompson, R. C. (2013). Microplastic moves pollutants and additives to worms, reducing functions linked to health and biodiversity. Current Biology, 23(23), 2388-2392.
  14. Buchs, B., Evangelou, M. W., Winkel, L. H., Lenz, M. (2013). Colloidal properties of nanoparticular biogenic selenium govern environmental fate and bioremediation effectiveness. Environmental science & technology, 47(5), 2401-2407.
  15. Castañeda, R. A., Avlijas, S., Simard, M. A., Ricciardi, A. (2014). Microplastic pollution in St. Lawrence River sediments. Canadian Journal of Fisheries and Aquatic Sciences, 71(12), 1767-1771.
  16. Chern, C. S. (2006). Emulsion polymerization mechanisms and kinetics. Progress in polymer science, 31(5), 443-486.
  17. Colabuono, F. I., Barquete, V., Domingues, B. S., Montone, R. C. (2009). Plastic ingestion by Procellariiformes in southern Brazil. Marine Pollution Bulletin, 58(1), 93-96.
  18. Cole, M., Lindeque, P., Fileman, E., Halsband, C., Galloway, T. S. (2015). The Impact of Polystyrene Microplastics on Feeding, Function and Fecundity in the Marine Copepod Calanus helgolandicus. Environmental science & technology.
  19. Corcoran, P. L., Norris, T., Ceccanese, T., Walzak, M. J., Helm, P. A., Marvin, C. H. (2015). Hidden plastics of Lake Ontario, Canada and their potential preservation in the sediment record. Environmental Pollution, 204, 17-25.
  20. Cózar, A., Echevarría, F., González-Gordillo, J. I., Irigoien, X., Úbeda, B., Hernández-León, S., Palma, A.T. , Navarro, S., García-de-Lomas, J., Ruiz, A., Fernandez-de-Puelles, M.L. Duarte, C. M. (2014). Plastic debris in the open ocean. Proceedings of the National Academy of Sciences, 111(28), 10239-10244.
  21. Dale, A. L., Casman, E. A., Lowry, G. V., Lead, J. R., Viparelli, E., Baalousha, M. (2015). Modeling nanomaterial environmental fate in aquatic systems. Environmental science & technology, 49(5), 2587-2593.
  22. Dantas, D. V., Barletta, M., da Costa, M. F. (2012). The seasonal and spatial patterns of ingestion of polyfilament nylon fragments by estuarine drums (Sciaenidae). Environmental Science and Pollution Research, 19(2), 600-606.
  23. Darlington, T. K., Neigh, A. M., Spencer, M. T., Guyen, O. T., Oldenburg, S. J. (2009). Nanoparticle characteristics affecting environmental fate and transport through soil. Environmental Toxicology and Chemistry, 28(6), 1191-1199.
  24. de Sá, L. C., Luís, L. G., Guilhermino, L. (2015). Effects of microplastics on juveniles of the common goby (Pomatoschistus microps): Confusion with prey, reduction of the predatory performance and efficiency, and possible influence of developmental conditions. Environmental Pollution, 196, 359-362.
  25. Dellinger, B., Pryor, W. A., Cueto, R., Squadrito, G. L., Hegde, V., Deutsch, W. A. (2001). Role of free radicals in the toxicity of airborne fine particulate matter. Chemical research in toxicology, 14(10), 1371-1377.
  26. Desforges, J. P. W., Galbraith, M., Dangerfield, N., Ross, P. S. (2014). Widespread distribution of microplastics in subsurface seawater in the NE Pacific Ocean. Marine pollution bulletin, 79(1), 94-99.
  27. DeSimone, J. M., Maury, E. E., Menceloglu, Y. Z., McClain, J. B., Romack, T. J., Combes, J. R. (1994). Dispersion polymerizations in supercritical carbon dioxide. Science, 265(5170), 356-359.
  28. Dowding, P. J., Vincent, B. (2000). Suspension polymerisation to form polymer beads. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 161(2), 259-269.
  29. Dris, R., Gasperi, J., Tassin, B. (2014, January). Assessing the microplastics in urban effluents and in the Seine River (Paris). In Fate and impacts of microplastics in marine ecosystems.
  30. Eriksen, M., Lebreton, L. C., Carson, H. S., Thiel, M., Moore, C. J., Borerro, J. C., Galgani, F., Ryan, P.G. Reisser, J. (2014). Plastic Pollution in the World's Oceans: More than 5 Trillion Plastic Pieces Weighing over 250,000 Tons Afloat at Sea. PloS one, 9(12), e111913.
  31. Eriksen, M., Mason, S., Wilson, S., Box, C., Zellers, A., Edwards, W., Farley, H. Amato, S. (2013). Microplastic pollution in the surface waters of the Laurentian Great Lakes. Marine pollution bulletin, 77(1), 177-182.
  32. Eriksson, C., Burton, H. (2003). Origins and biological accumulation of small plastic particles in fur seals from Macquarie Island. AMBIO: A Journal of the Human Environment, 32(6), 380-384.
  33. Fendall, L. S., Sewell, M. A. (2009). Contributing to marine pollution by washing your face: Microplastics in facial cleansers. Marine Pollution Bulletin, 58(8), 1225-1228.
  34. Foekema, E. M., De Gruijter, C., Mergia, M. T., van Franeker, J. A., Murk, A. J., Koelmans, A. A. (2013). Plastic in North Sea fish. Environmental science & technology, 47(15), 8818-8824.
  35. Fossi, M. C., Coppola, D., Baini, M., Giannetti, M., Guerranti, C., Marsili, L., Panti, C., de Sabata, E., Clò, S. Marine Environmental Research. 2014, 100, 17-24.
  36. Fry, D. M., Fefer, S. I., Sileo, L. (1987). Ingestion of plastic debris by Laysan Albatrosses and Wedge-tailed Shearwaters in the Hawaiian Islands. Marine Pollution Bulletin, 18(6), 339-343.
  37. GESAMP. (2015). Microplastics in the ocean.
  38. Gouin, T., Roche, N., Lohmann, R., Hodges, G. (2011). A thermodynamic approach for assessing the environmental exposure of chemicals absorbed to microplastic. Environmental Science & Technology, 45(4), 1466-1472.
  39. Graham, E. R., Thompson, J. T. (2009). Deposit-and suspension-feeding sea cucumbers (Echinodermata) ingest plastic fragments. Journal of Experimental Marine Biology and Ecology, 368(1), 22-29.
  40. Hall, N. M., Berry, K. L. E., Rintoul, L., Hoogenboom, M. O. (2015). Microplastic ingestion by scleractinian corals. Marine Biology, 162(3), 725-732.
  41. Haimer, J., Gutow, L., Koihler, A., Saborowski, R. (2014). Fate of Microplastics in the Marine Isopod Idotea emarginata. Environmental science & technology, 48(22), 13451-13458.
  42. He, W. D., Sun, X. L., Wan, W. M., Pan, C. Y. (2011). Multiple morphologies of PAA-b-PSt assemblies throughout RAFT dispersion polymerization of styrene with PAA Macro-CTA. Macromolecules, 44(9), 3358-3365.
  43. Hennigan, C. J., Miracolo, M. A., Engelhart, G. J., May, A. A., Presto, A. A., Lee, T., Sullivan, A. P., McMeeking, G. R., Coe, H., Wold, C. E., Hao, W.-M., Gilman, J. B., Kuster, W. C., de Gouw, J., Schichtel, B. A., Collett Jr., J. L.,  Kreidenweis, S. M, Robinson, A. L. (2011). Chemical and physical transformations of organic aerosol from the photo-oxidation of open biomass burning emissions in an environmental chamber. Atmospheric Chemistry and Physics, 11(15), 7669-7686.
  44. Hidalgo-Ruz, V., Gutow, L., Thompson, R. C., Thiel, M. (2012). Microplastics in the marine environment: a review of the methods used for identification and quantification. Environmental science & technology, 46(6), 3060-3075.
  45. Imhof, H. K., Ivleva, N. P., Schmid, J., Niessner, R., Laforsch, C. (2013). Contamination of beach sediments of a subalpine lake with microplastic particles. Current biology, 23(19), R867-R868.
  46. Jambeck, J. R., Geyer, R., Wilcox, C., Siegler, T. R., Perryman, M., Andrady, A., Narayan, R. Law, K. L. (2015). Plastic waste inputs from land into the ocean. Science, 347(6223), 768-771.
  47. Jeftic, L., Sheavly, S. B., Adler, E. (2009). Marine litter: a global challenge. N. Meith (Ed.). Regional Seas, United Nations Environment Programme.
  48. Jinhua, L., Guangyuan, Z. (2014). Polystyrene Microbeads by Dispersion Polymerization: Effect of Solvent on Particle Morphology. International Journal of Polymer Science, 2014.
  49. Kaposi, K. L., Mos, B., Kelaher, B. P., Dworjanyn, S. A. (2014). Ingestion of microplastic has limited impact on a marine larva. Environmental science & technology, 48(3), 1638-1645.
  50. Law, K. L. Thompson, R. C. (2014). Microplastics in the seas. Science, 345(6193), 144-145.
  51. Lee, K. W., Shim, W. J., Kwon, O. Y., Kang, J. H. (2013). Size-dependent effects of micro polystyrene particles in the marine copepod Tigriopus japonicus. Environmental science & technology, 47(19), 11278-11283.
  52. Leslie, H. A. (2014). Review of Microplastics in Cosmetics. Institute for Environmental Studies [IVM].
  53. Leslie, H. A., van Velzen, M. J. M., Vethaak, A. D. (2013). Microplastic survey of the Dutch environment. Novel data set of microplastics in North Sea sediments, treated wastewater effluents and marine biota.
  54. Li, Y., Armes, S. P. (2010). RAFT synthesis of sterically stabilized methacrylic nanolatexes and vesicles by aqueous dispersion polymerization. Angewandte Chemie, 122(24), 4136-4140.
  55. Lusher, A. L., McHugh, M., Thompson, R. C. (2013). Occurrence of microplastics in the gastrointestinal tract of pelagic and demersal fish from the English Channel. Marine pollution bulletin, 67(1), 94-99.
  56. Machado, F., Lima, E. L., Pinto, J. C. (2007). A review on suspension polymerization processes. Polímeros, 17(2), 166-179.
  57. Mathalon, A., Hill, P. (2014). Microplastic fibers in the intertidal ecosystem surrounding Halifax Harbor, Nova Scotia. Marine pollution bulletin, 81(1), 69-79.
  58. Mato, Y., Isobe, T., Takada, H., Kanehiro, H., Ohtake, C., Kaminuma, T. (2001). Plastic resin pellets as a transport medium for toxic chemicals in the marine environment. Environmental science & technology, 35(2), 318-324.
  59. Murray, F., Cowie, P. R. (2011). Plastic contamination in the decapod crustacean Nephrops norvegicus (Linnaeus, 1758). Marine Pollution Bulletin, 62(6), 1207-1217.
  60. Napper, I. E. Thompson, R. C. (2015). Characterisation, Quantity and Sorptive Properties of Microplastics Extracted From Cosmetics. Marine Pollution Bulletin (in press).
  61. New York, Office of New York State Attorney General Eric T. Schneiderman [New York]. (2015). Unseen Threat: How Microbeads Harm New York Water, Wildlife, Health And Environment. Retrieved from: http://ag.ny.gov/pdfs/Microbeads_Report_5_14_14.pdf
  62. Nobre, C. R., Santana, M. F. M., Maluf, A., Cortez, F. S., Cesar, A., Pereira, C. D. S., Turra, A. (2015). Assessment of microplastic toxicity to embryonic development of the sea urchin Lytechinus variegatus (Echinodermata: Echinoidea). Marine pollution bulletin, 92(1), 99-104.
  63. Norwegian Environment Agency (Miljødirektoratet). (2014). Sources of microplastic-pollution to the marine environment. Sundt, P., Schulze, P-E., Syversen F. Retrieved from http://www.miljodirektoratet.no/Documents/publikasjoner/M321/M321.pdf
  64. Obbard, R. W., Sadri, S., Wong, Y. Q., Khitun, A. A., Baker, I., Thompson, R. C. (2014). Global warming releases microplastic legacy frozen in Arctic Sea ice. Earth's Future, 2(6), 315-320.
  65. Possatto, F. E., Barletta, M., Costa, M. F., do Sul, J. A. I., Dantas, D. V. (2011). Plastic debris ingestion by marine catfish: an unexpected fisheries impact. Marine Pollution Bulletin, 62(5), 1098-1102.
  66. Quadros, M. E., Marr, L. C. (2010). Environmental and human health risks of aerosolized silver nanoparticles. Journal of the Air & Waste Management Association, 60(7), 770-781.
  67. Rochman, C. M., Hoh, E., Kurobe, T., Teh, S. J. (2013). Ingested plastic transfers hazardous chemicals to fish and induces hepatic stress. Scientific reports, 3.
  68. Rochman, C. M., Kurobe, T., Flores, I., Teh, S. J. (2014). Early warning signs of endocrine disruption in adult fish from the ingestion of polyethylene with and without sorbed chemical pollutants from the marine environment. Science of the Total Environment, 493, 656-661.
  69. Rosenkranz, P., Chaudhry, Q., Stone, V., Fernandes, T. F. (2009). A comparison of nanoparticle and fine particle uptake by Daphnia magna. Environmental Toxicology and Chemistry, 28(10), 2142-2149.
  70. Setälä, O., Fleming-Lehtinen, V., Lehtiniemi, M. (2014). Ingestion and transfer of microplastics in the planktonic food web. Environmental pollution, 185, 77-83.
  71. State of New York [SNY] (2014).  Unseen Threat: How Microbeads Harm New York Waters, Wildlife, Health and Environment.  Available from: http://www.ag.ny.gov/press-release/ag-schneiderman-releases-report-outlining-urgent-need-pass-microbeads-ban
  72. Syberg, K., Khan, F. R., Selck, H., Palmqvist, A., Banta, G. T., Daley, J., Sano, L. Duhaime, M. B. (2015). Microplastics: addressing ecological risk through lessons learned. Environmental Toxicology and Chemistry, 34(5), 945-953.
  73. Talvitie J., Heinonen M. HELCOM. (2014). BASE project 2012-2014: Preliminary study on synthetic microfibers and particles at a municipal waste water treatment plant. Retrieved from http://www.helcom.fi/mwg-internal/de5fs23hu73ds/progress?id=eiKN8huaqatMPUlZ2_PuNH-hQ_iUzC03ZnsE9o_IWw0,
  74. Teuten, E. L., Rowland, S. J., Galloway, T. S., Thompson, R. C. (2007). Potential for plastics to transport hydrophobic contaminants. Environmental science & technology, 41(22), 7759-7764.
  75. Teuten, E. L., Saquing, J. M., Knappe, D. R., Barlaz, M. A., Jonsson, S., Björn, A., Rowland, S.J., Thompson, R.C., Galloway, T.S., Yamashita, R., Ochi, D., Watanuki, Y., Moore, C., Viet, P.H., Tana, T.S., Prudente, M., Boonyatumanond, R., Zakaria, M.P., Akkhavong, K., Ogata, Y., Hirai, H., Iwasa, S., Mizukawa, K., Hagino, Y., Imamura, A., Saha, M. Takada, H. (2009). Transport and release of chemicals from plastics to the environment and to wildlife. Philosophical Transactions of the Royal Society B: Biological Sciences, 364(1526), 2027-2045.
  76. Thompson, R. C., Olsen, Y., Mitchell, R. P., Davis, A., Rowland, S. J., John, A. W., McGonigle, D. Russell, A. E. (2004). Lost at sea: where is all the plastic?. Science, 304(5672), 838-838.
  77. United Nations Environment Programme [UNEP] (2015).  Plastic in Cosmetics. Retrieved from:  http://www.unep.org/newscentre/Default.aspx?DocumentID=26827&ArticleID=35180
  78. United Nations Environment Programme, International Environmental Technology Centre [UNEP-IETC] (2012). Project Converting Waste Plastic into Fuel. Retrieved May 21, 2015, from http://www.unep.org/ietc/OurWork/WasteManagement/Projects/wastePlasticsProject/tabid/79203/Default.aspx
  79. Van Cauwenberghe, L., Janssen, C. R. (2014). Microplastics in bivalves cultured for human consumption. Environmental Pollution, 193, 65-70.
  80. van Franeker, J. A., Bell, P. J. (1988). Plastic ingestion by petrels breeding in Antarctica. Marine Pollution Bulletin, 19(12), 672-674.
  81. von Moos, N., Burkhardt-Holm, P., Koihler, A. (2012). Uptake and effects of microplastics on cells and tissue of the blue mussel Mytilus edulis L. after an experimental exposure. Environmental science & technology, 46(20), 11327-11335.
  82. Woodall, L. C., Sanchez-Vidal, A., Canals, M., Paterson, G. L., Coppock, R., Sleight, V., Calafat, A., Rogers, A.D., Narayanaswamy, B.E. Thompson, R. C. (2014). The deep sea is a major sink for microplastic debris. Royal Society Open Science, 1(4), 140317.
  83. Zbyszewski, M., Corcoran, P. L. (2011). Distribution and degradation of fresh water plastic particles along the beaches of Lake Huron, Canada. Water, Air, & Soil Pollution, 220(1-4), 365-372.
  84. Zbyszewski, M., Corcoran, P. L., Hockin, A. (2014). Comparison of the distribution and degradation of plastic debris along shorelines of the Great Lakes, North America. Journal of Great Lakes Research, 40(2), 288-299.
  85. Zubris, K. A. V., Richards, B. K. (2005). Synthetic fibers as an indicator of land application of sludge. Environmental pollution, 138(2), 201-211.

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