Isolation of nanoplastics in fish from the North Aegean SeaPURE AND APPLIED BIOCHEMISTRY | FACULTY OF ENGINEERING | LUND UNIVERSITY ISABELLA GIMSKOG | MASTER THESIS 2019

Isolation of nanoplastics in fish from theNorth Aegean sea

2019

Master thesis by Isabella Gimskog

Division of Pure and Applied Biochemistry, Department of Chemistry, LTH

Supervisor: Tommy Cedervall, Senior lecturer at Biochemistry andStructural Biology

Examiner: Lei Ye, Professor at Pure and Applied Biochemistry

Cover image: Eric Gaba (Wikimedia Commons user: Sting)

Printed by: Media-Tryck KC

2019

Abstract

The breakdown of plastics in our oceans is one of our times biggest challenges. What happens when theseplastics are broken down into pieces too small to see is still a mystery. Nanoplastics is a relatively new area ofresearch and much more studies need to be done before we can get a perspective of how big the problem is.To be able to study these plastics we need a way to attain nanoplastic samples from our oceans. This thesisaims to act as a first step in suggesting a method to isolate eventual nanoplastics accumulated in fish.

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Contents

1 Popular sientific summary 41.1 Is it possible to isolate nanoplastics from the fish we eat? . . . . . . . . . . . . . . . . . . . . . . . . 4

2 Preface 5

3 Introduction 63.1 Plastic in the Mediterranean sea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63.2 Defining micro- and nanoplastics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63.3 Source of micro- and nanoplastics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63.4 Breakdown of plastics in the ocean . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73.5 Interaction with marine life . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

3.5.1 Organisms that ’eat plastic’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83.6 Properties of nanoplastics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93.7 Toxins carried by plastic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

4 Method 104.1 Abbreviations and sample names . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104.2 Analyze of microplastic content in fish in a field lab . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104.3 Water samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

4.3.1 Dynamic Light Scattering and Nano Tracking Analysis . . . . . . . . . . . . . . . . . . . . . . 124.3.2 Centrifugation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124.3.3 Fourier-Transform InfraRed spectroscopy analysis . . . . . . . . . . . . . . . . . . . . . . . . 124.3.4 Size exclusion chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124.3.5 Absorbance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

5 Results and discussion 145.1 Primary microplastic analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145.2 Breakdown of microplastic fibres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155.3 Breakdown of polystyrene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165.4 Absorbance and NTA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175.5 Analysis of the water samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19

5.5.1 Assessment of particle populations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195.5.2 Density assessment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225.5.3 Absorbance analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255.5.4 Desalting the samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265.5.5 Concentrating the particles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

6 Future research 31

7 Conclusions 31

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1 Popular sientific summary

1.1 Is it possible to isolate nanoplastics from the fish we eat?

This thesis investigates the possibility to find nanoplastic particles in the gills and digestive tract of food fishfrom the Mediterranean sea.

During the last few years microplastics has gone from being an almost unknown word to being a hot topic fre-quently discussed in media and possibly one of the worst environmental threats of our time. But what happenswhen the microplastics break down? The answer we all would wish for is that they disappear or become usefulmatter for the ecosystem again. This might unfortunately not be the case as plastics never break down fully butare just fragmented into smaller and smaller pieces and we might stand before an even more incomprehensiblethreat. Nanoplastics.

Nanoplastics are usually defined as plastic particles with at least one dimension under 100nm and they are stillmuch of a mystery in how they are formed and how they affect the ecosystem in which they are present. Itcan be hard to comprehend the difference between micro- and nanoplastics but as microplastics are usuallydefined as plastic pieces smaller than 5mm, the difference in scale is like comparing a Olympic size swimmingpool to a single drop of water. They are so small that they are not restricted to the pathways we are used to andprevious research shows that they can even cross the brain-blood-barrier in fish.

Nanoplastics is a new area of research and it is of great importance that we learn more, fast. This thesis aims tobe a first step in finding a way to detect existence of nanoplastics in the fish we eat and thereby in our oceans.The work analyses the the particles from the digestive tract and gills of fish below the size of 1,2 µm. Thefindings indicate that there are nanoplastics present in the fish and that these plastics are possible to isolate.

The report also includes mechanical breakdown of microplastics found in the same fish and of a polystyrenecoffee cup lid. All experiments show that nanoplastics are released in the break down process.

The hope is that this report can act as a starting point and a guide to anyone who is curious about nanoplasticsand wish to isolate them from complex samples.

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2 Preface

This report is my thesis work which began in February 2019. The first two and a half months were spent onthe island of Samos in Greece working with microplastics for Archipelagos marine conservation institute andcollecting samples from locally sourced fish. Mid April to August was spent at the biochemical department atLund Univeristy analyzing the samples brought back from Samos.

Most of all I would like to thank my supervisor Tommy Cedervall without whom this project would not havebeen possible, for always answering my questions, for his great expertise and for always reminding me to lookon the bright side and finding new ways when my results have not been what I hoped for. I also want to givea big thank you to Martin Lundqvist and Mikael Ekvall who has helped me a lot with input and teaching mehow the machines work. And of course the rest of the department that have supported me and helped me withvaluable input and to find my way when I have gotten lost in the corridors.

I also want to thank Archipelagos Institution of Marine Conservation for letting me do the field work there andhelping me collecting my samples. A special thanks to Dr. Guido Pietroluongo, Belen Quintana and SimoneAntichi who supervised me and helped me make my project possible.

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3 Introduction

This master thesis aims to see if it is possible to isolate nanoplastics from the gills and digestive tract of fishcaptured in an area known to have a high concentration of microplastics.

The work is meant to further the research on nanoplastics and therefore both successful methods and unsuc-cessful methods are included. The report aims to act as a guide to anyone who wants to do similar work andtherefore includes what I believe works and mistakes that I have made that can be avoided.

3.1 Plastic in the Mediterranean sea

The Mediterranean sea is severely polluted due to several factors, some of the most important being its denselypopulated coastlines, large fishing industries, heavy tourism and intense shipping. The very limited outflowof surface water makes the pollution more concentrated than for larger seas. It is suggested that the Mediter-ranean could have among the highest concentration of microplastics in the world [1]. Even tough this is veryalarming it makes it an interesting starting point for research on plastic pollution. It can be seen as an indicatorof how the world’s oceans will react if the plastic pollution continues at our current rate.

The Aegean sea is a part of the Mediterranean located between Greece and Turkey and is in no way an exceptionto the Mediterraneans pollution problem with its many, many islands. In addition the border between Turkeyand Greece as an outer border of the EU makes this area harder to regulate and protect.

The first part of this project is done at Archipelagos Institute of Marine Conservation research base in Samos,Greece. Archipelagos is an non governmental organisation working to protect the wildlife in the Mediterraneanby conducting research and spreading information.

3.2 Defining micro- and nanoplastics

The definition of microplastics is not uniform but commonly defined as plastics smaller than 5 mm. Nanoplas-tics also lack a strict definition, one study on what nanoplastics are suggests the definition as particles unin-tentionally produced (i.e. from the degradation and the manufacturing of the plastic objects) and presenting acolloidal behavior, within the size range from 1 to 1000 nm [2].

Several reports on the subject highlight the importance of establishing unified international standards.

3.3 Source of micro- and nanoplastics

Microplastics enter the oceans either as fabricated microplastics from sources such as cosmetics or toothpaste.They can also enter the oceans as macroplastics and subsequently be broken down into micro- and nanoplas-tics trough UV-degradation, chemical processes and mechanical stress. They can also end up in the sea viawaste water from cleaning plants. Synthetic clothes have been observed to release a lot of microplastic frag-ments. One wash of a single fleece sweater can lose as much as 1900 plastic fibres [3].

Nanoplastics is a much newer research field and their presence have not yet been entirely confirmed in theocean waters but it is likely that they are present to some extent. Some hygiene products containing mi-croplastics have also been confirmed to contain nanoplastics [4]. There has been studies on the breakdown ofpolystyrene that proves the release nanoplastics during its breakdown [5] [6]. The research that has been doneon nanoplastics suggest possibly grim impacts on our oceans that we might grow accustomed to in the future.A possible reason that we have yet not noticed any vast effects of nanoplastics on the marine environment isthat the breakdown of plastics take so long that the most plastics left in the ocean during the last centuriessimply have not gotten to that state yet. According to Koelmans, Besseling and Shim [7] the breakdown of amicroplastic measuring 1 mm to nanoplastic size (100 nm) would take 320 years in the lab and probably evenlonger in the ocean.

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3.4 Breakdown of plastics in the ocean

To learn how long it takes for different types and structures of plastic to break down in the ocean has provendifficult as there are no methods of determining the age of microplastics found in the field [8]. Using the term"break down" can itself be problematic as all types of plastics do not decompose into parts that can be in-cluded into the ecosystem again. It is rather fragmented into smaller and smaller parts. Plastic as a material isunique in many ways and evolution has no possibility to adapt to such a rapid explosion of use and the conse-quences that may come of the careless way we have handled its disposal and recycling. Fig.1 is an illustrationby Maphoto/Riccardo Pravettoni that shows how long a plastic product will stay in, and affect, its marine envi-ronment.

Figure 1: illustration by Maphoto/Riccardo Pravettoni, 2018 from http: // www. grida. no/ resources/ 6914 showinghow long plastic waste stays in the marine environment.

The breakdown of polymers can be divided into six different categories[9]. Of these all except for thermal degra-dation contribute to break down of plastics in the oceans with the most important being photo degradation[8].

1. Thermal degradation:

The breakdown of plastics using heat. This reaction is not naturally occurring in the oceans.

2. Hydrolysis:

Breaking bonds in the polymer by addition of water.

3. Mechanical degradation:

The physical disintegration of plastics, in the oceans caused by the stress from waves and collisions withrocks or other structures, crushing the plastic.

4. Thermo-oxidative degradation: A slow oxidative breakdown that occurs at moderate temperatures, inthe oceans caused by radiation from the sun. This process also works differently on plastics treated withadditives such as UV and heat stabilizers which makes the process slower[8].

5. Photodegradation:

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Caused by light, in the oceans sunlight, that effectively makes the plastic weak and brittle.

6. Biodegradation:

Breakdown caused by microbes, such as bacteria. In the ocean this effect will be more prominent inwaters that are warm and contain a lot of oxygen.

These processes have been extensively studied in the development of plastic products to make the material asdurable as needed but not as much research has gone into investigating the complete breakdown into micro-and nanoplastics in the environment.

That macroplastics break down all the way to nanoplastics in the ocean is still not entirely proven since the sam-pling from the field is such a difficult process for nanoplastics. If most plastics do break down into nanoplasticswe might have a very big problem on our hands that we know very little about. On the other hand, if mostplastic stay in the microplastic scope the problem is still very much acute. It is even accelerating because ofour increasing use of plastics. At least we have some more knowledge about the effects of microplastics thanksto the extensive amount of studies during the last decade. It might be that we have not even seen the tip of theiceberg of problems caused by nanoplastics yet.

3.5 Interaction with marine life

A common sight in the media for the last few years have been evidence of the effects of plastics in our waters.Pictures of dead sea birds with their stomachs full of plastic, seals with fishnets around their throats and turtlesshaped into odd forms from the plastic from a six pack of beer. These photos are very clear evidence of thehorrible impact plastics have on wild life. Even when the plastics appear to have disappeared they are still thereas small particles and is potentially an even bigger threat to the world as we know it as decades of plastic wastecontinue to break down in our oceans. How these small particles affect life is not entirely clear yet, especially inthe case of nanoplastics. A lot of further studies are needed, although many negative effects have already beenreported. Some negative health effects in animals that have been reported from microplastics are increasedimmune response, starvation, energy depletion and even negative impacts on subsequent generations[10].

Studies also suggest that nanoplastics pose a great threat to the aquatic ecosystems and subsequently to all life,as the nanoplastics are able to cross membranes and other biological barriers, permanently damaging themand also providing a vector for the transport of persistent organic pollutants(see section "Toxins carried byplastic"). A study by Mattson et al., 2017 also shows that polystyrene nanoparticles are able to cross the bloodbrain barrier in fish and disturb their behaviour [11].

3.5.1 Organisms that ’eat plastic’

Some attention have been focused on animals that seem to be able to break down plastics in the ocean withoutbeing harmed by them. According to an article in the Guardian[12] one type of Antarctic krill are able to breakdown microplastics. It is suggested that the krill will break the microplastics down into nanoplastics. Thisprocess will however greatly increase the surface area of these plastic particles and might therefore potentiallymake the problem even worse.

There has also been suggested that some types of bacteria has evolved to break down the plastic type PET forexample written about in the Conversation [13], the author warns however if the bacteria can evolve to eatplastic this might become a threat to plastics and we might have to adopt to a plastic free lifestyle not only tosave the planet but also because plastic will not be as durable as it is today.

Trying to find an organism that is able to break down plastics is not an optimal way to attack the problem. Thematerial needs to be properly recycled, not only so it won’t pollute the environment but also so it can be usedto make new things. The demand for plastic is still increasing.

These organisms have not been studied in this work but the section is included to point out that there is noeasy cleanup for our oceans. We can’t expect nature to adapt to the plastic pollution and incorporate plasticwaste as a natural part of the food chain.

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3.6 Properties of nanoplastics

Nanoplastics have different properties than larger structures and even though they are too small to see withthe naked eye they can still cause a lot of problems. The nanoplastics have very large surface area relative totheir volume and when larger plastics are broken into nanoplastics the total surface area becomes extremelylarge. It also seems like these particles accumulate toxins in a marine environment and even though this is alsoobserved for microplastics it seems to be an even bigger problem for nanoplastics[14]. The high curvature ofthe nanoparticles also affects binding to other particles and differentiates nanoplastics from larger structures.

Another concern is that the nanoplastics are so small that they are not restricted by cell membranes and otherbarriers in organisms. It is therefore impossible to predict how the nanoplastics will move within an unstudiedorganism, where the nanoplastics may accumulate and how long they will remain. Long term effects need tobe investigated to be able to give indications and to state conclusions from nanoplastics.

3.7 Toxins carried by plastic

One mayor concern with micro- and nanoplastics is their ability to carry and accumulate toxins and chemicalpollutants, often in literature referred to as persistent organic pollutants (POPs).

POPs are organic compounds often found in for example solvents and pesticides. A POP have characteristicsthat make them an important problem in the modern world, these characteristics can be found in the Stock-holm Convention(2004)[15].

A POP should:

• Remain intact for exceptionally long periods of time (many years).

• Become widely distributed throughout the environment as a result of natural processes involving soil,water and, most notably, air.

• Accumulate in the fatty tissue of living organisms including humans, and are found at higher concentra-tions at higher levels in the food chain.

• Are toxic to both humans and wildlife.

A study on the Monogonont Rotifer Brachionus koreanus showed that the toxicity of POPs can be increased byingestion of nanoplastics[16]. The study also indicates that the nanoplastics not only are able to cross mem-branes but also permanently damage them.

It is very difficult to know how the adsorption of POPs to microplastics and nanoplastics work as the conditionsare hard to mimic in a lab. There are so many dynamic components in the ocean as well as very large timescales. Studies however give us an indication of how certain POPs bind to certain plastic particles.

These small plastic particles bind to hydrophobic toxic pollutants and studies[14] have shown that nanoplas-tics provide an additive effect to bio accumulation to these toxic substances. These studies indicate that thenanoplastics may pose a very large threat to humans as they will act to higher concentrations of toxic sub-stances in the marine food chain and therefore in the food we eat.

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4 Method

4.1 Abbreviations and sample names

A summary of the most common abbreviations used in this report. The samples is referred to by the organ, thenumber of the fish (1-7) and if the water is a surface fraction or a bottom fraction.

• MP= Microplastic

• NP = Nanoplastic

• PS = Polystyrene

• NTA = Nano Tracking Analysis

• DLS = Dynamic Light Scattering

• FTIR = Fourier-Transform InfraRed spectroscopy

• DT = Digestive Tract

• S = Surface

• B = Bottom

4.2 Analyze of microplastic content in fish in a field lab

The sample collection for this project was done in a field lab at Archipelagos Institute of Marine Conservationin Samos, Greece. The method is developed to be simple and cost efficient to promote the research to anyonewho wants to do research without the need of expensive tools and equipment.

All analysis is done on fish bought at the local fish market.

1. Sample collection

To analyze the microplastic content the fish were dissected by extracting the digestive system and thegills.

Seven fish of the species Dentex macrophthalmus were used for sample collection. Before dissection, theoutside of the fish were rinsed with distilled water to remove any plastic contaminants the fish might haveattained in the handling process. All fish were measured and weighed. The measurements are shown intable 1.

Fish total weight (g) total length (cm) gut weight (g) gill weight (g)1 115 19 3,38 52 132 20,5 5,04 33 109 19 8,41 34 110 18,5 3,92 25 91 18,5 1,96 36 111 19 2,05 37 87 18 2,33 4

Table 1: Measurements of fish used in sample collection.

The gills were isolated, weighed and put in a glass container with 100 ml of 20% saline solution.

The digestive tracts were also collected, weighed and put in glass containers. The inside of the fish andthe head were rinsed with distilled water and the water collected in the same container.

2. Pre-filtration

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The digestive systems of the fish were mushed in a stone mortar to make sure the contents of the bowlswere available, everything was then put back in the jar. The samples were then left to sit for 48 hours. Thegill samples were not ground but only shaken to try to free plastics adsorbed to surface of the gills.

All samples were run through a metal sieve with a mesh size of 200 µm using distilled water to rinse.

This pre-filtration step aims to collect only fluid matter, small particles and microplastics, while gettingrid of most organic matter.

3. Filtration preparation

A saline solution (20% NaCl) is used to dilute the DT samples 1:1 and the sample is then left to set forroughly 48 hours. This is a density separation and the microplastics are expected to float to the surface.

The gills were also taken out after 48 hours and rinsed with 20 ml of saline solution using a glass syringe,the saline solution used to rinse was collected with the rest of the sample.

4. Filtration

Before the final filtration a few drops of vinegar was added to all samples to remove mineral sediments.

The samples are then filtered 20 ml at a time using filter paper with a pore size of 1,2 µm, a glass filtrationflask and a vacuum pump. The filter paper is then placed on a glass Petri plate and marked with the nameand series number of the sample.

A control sample, of the saline solution used in diluting the samples, is also collected. This to be able todetect any contributions from the salt, distilled water or the filters.

5. Collection of water samples

Water samples of 10 ml for nanoparticle analysis were collected for each sample jar and the control sam-ple. These were collected after the first three 20 ml fractions were collected and then a second one fromthe last three fractions collected. This was to see if a difference could be seen between the surface andbottom fractions of the samples. The samples were marked with fish number, organ and S or B for surfaceor bottom.

6. Analysis

The filter papers are then analyzed in a microscope with a magnification of x40. Microplastics are recordedand categorized by type, color and size. The different "types" are fibre, film or fragment. To suspect some-thing to be a plastic it should have a homogeneous thickness, color and gloss and not exhibit any cellularor organic structures[10]. The plastics found are tested to confirm that they are plastic by conducting"the hot needle test" where a needle is heated up and brought close to the fragment to see if it curls up,if so it is a plastic, if it does not move it is not recorded as a plastic. All plastics found in this experimentwere fibres.

The samples were then packed and shipped to Lund for more advanced analysis.

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4.3 Water samples

This section lists a short description of the methods used when analysing the water samples.

4.3.1 Dynamic Light Scattering and Nano Tracking Analysis

Dynamic light scattering, more often referred to as DLS is a analyzing tool used to give a size distribution ofparticles in a solution by looking at the scattered light from the particles. This method does not tell the con-centration of particles.

When DLS has been used in this report, several additional methods have been analyzed to investigate the va-lidity of the results.

Nano Tracking Analysis (NTA) was in this report used in combination with the DLS to get an estimate of the sizedistribution of particles in the samples. Both methods are based on observing Brownian motion. The differenceis that the NTA converts this information to a size distribution trough image analysis while the DLS uses timedependent fluctuations in scattering intensity. The NTA is better for individual particles and heterogeneoussolutions but the DLS is faster and forms an average of a greater number of particles [17].

The operator has a big influence of the results in NTA analysis since the measuring parameters can be regulatedfor each sample. This is important to take into consideration when analysing the results. It is also importantto note that the particles found in the NTA are not necessarily the smallest ones but the smallest ones theinstrument is able to detect.

4.3.2 Centrifugation

In this work centrifugation is used to separate particles from each other using differences in density. The in-creased centrifugal force of the rotation will force particles to "sink" to the bottom faster and there be collectedin a pellet. Therefore low density particles will require high centrifugal force to be collected in a pellet.

This method is useful when looking for plastics as they have relatively low density and should require a highcentrifugal force to sink to the bottom.

4.3.3 Fourier-Transform InfraRed spectroscopy analysis

Fourier-Transform InfraRed spectroscopy (FTIR) is a technique, in this work used for material identification,and it works by applying a infrared range of wavelengths to a sample. A Fourier transformation is used to at-tain a spectra. This type of absorbance analysis uses a infrared spectrum of light and the light beam is beingmodified during the measurement to periodically block and transmit different wavelengths. This gives manydifferent data points and by applying the Fourier transformation a spectra with the absorbance for each wave-length is attained. This method of measuring several wavelengths at once gives the method a higher signal tonoise ratio [18].

4.3.4 Size exclusion chromatography

The principle of size exclusion chromatography is simple. A sample is introduced to a column with a porousgel. Larger molecules, with a radius larger than the largest pore radius, will be eluted first as they go trough thecolumn without entering into the gel matrix. The smaller molecules, like salt, will enter into the pores of thematrix, making their way trough the column longer and they will therefore be eluted later.[19] The principle isillustrated in fig.2.

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Figure 2: Basic principle behind size exclusion chromatography with a sample with particles of different sizes being intro-duced in the first step. In the second step the smaller particles enter the pores and the larger particles take the shorter pathbetween the gel particles. Once eluted the large particles will leave the column first and can be collected as separate fractions.

In this thesis PD10 columns, Sephadex G-25 M, from GE healthcare were used for desalting samples. This wasdone for samples of both the digestive tract and the gills as well as for a pure saline solution sample to be ableto compare the results.

4.3.5 Absorbance

Absorbance is measured by looking at the transmittance of a sample in a spectrometer between a range ofwavelengths. For the purpose of finding nanoplastics the peak 230 nm were of certain interest as previous workshowed this peak for pure polystyrene[6].

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5 Results and discussion

5.1 Primary microplastic analysis

The filters from the gills and digestive systems were analysed in a light microscope with a magnification of 40.A summary of how many microplastic fibres found in each fish and organ can be seen in table 2. Some fibreswere hard to see due to organic matter on the filter and transparency or white color of the fibre. It is thereforelikely that there were more fibres present than the ones accounted for.

Fish nr. microplasics in gills microplastics in digestive tract1 33 92 16 223 29 114 14 105 13 46 9 137 20 12

Table 2: Number of microplastic fibres found in each fish during the microscopy.

Several attempts were made to analyse fibres in FTIR but this proved to be a big challenge. Single fibres weretoo thin to see with the naked eye and were therefore not possible to transfer to the FTIR. Further attemptswere made to collect a small "pile" of fibres on the filter and then measure directly on the pile, using a plasticfree region of the filter as background. The results of these measurements were inconclusive. The final plan foranalysis of the fibres was to collect them all in two test tubes, one test tube for the gill and one for the digestivetract and then try to break these down using a blender and comparing them to the water samples.

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a. Single microplastic fibre on filter. b. Tweezers and microplastic fibre

c. Microplastic fibre in 1,5 ml eppendorf tube. d. Collection of five fibres on a filter.

Figure 7: Different steps in the handling process of the microplastic fibres on the filters.

To illustrate the difficulty of handling these fibres fig.7 shows different steps of handling the fibres. They are verysmall and in comparison the tweezers are very big and small irregularities from ware on the tweezers make itvery challenging to get a grip of the fibres. Once collected into a 1,5 ml eppendorf tube a single fibre could notbe found again with the naked eye.

5.2 Breakdown of microplastic fibres

Since the microplastic fibres could not be tested in the FTIR, another approach was taken. All of the fibresfrom the gill samples and all the fibres from the digestive tract samples were collected into one eppendorf tuberespectively, these can be seen in fig.8.

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Figure 8: Images depicting the microfibres once collected in 1,5 ml tubes. The upper images are the fibres from the gills andthe lower are the ones from the digestive tract.

The collected fibres were put in test tubes with 4 ml of milli-Q water each and broken down with mechanicalforce for one plus five minutes using a homogeniser of the model VDI 12 from VWR laboratory equipment.

The homogeniser was first tested by mixing 10 samples of milli-Q water to see if any particles were releasedfrom the mixer or test tubes. This test proved that no particles were released from the homogeniser.

The test tubes used for break down of the microplastic fibres were first rinsed thoroughly to avoid any con-tamination. The results of the break down were analyzed for absorbance and NTA after one and six minutesof blending to see how the breakdown progressed. After the blending, pieces of fibres could still be seen i thewater. During the second blending which lasted for five minutes the sample got very heated and breaks had tobe taken to let it cool off, however the heat might have effected the breakdown.

The purpose of the break down of the microplastics was to be able to compare with the water samples as wellas a sample of raw PS and see if the samples resemble one another. These samples were then tested in NTA andabsorbance.

5.3 Breakdown of polystyrene

To see how pure polystyrene (PS) looked in these analysis, a part of a PS coffee cup lid were also broken downby the same method. The weight of the pice of PS used was 0,3 g.

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Figure 9: Pieces of polystyrene coffee lid. First picture: pieces cut down with scissors, second picture: blended for one minute.Third picture: blended for 6 minutes.

The sample with pure PS had much more material than the other samples and the pieces were larger so they cannot be used as a direct comparison to the fibres. After being blended for six minutes the samples was noticeablybroken down. A range of different sizes could be seen in the water and the water looked more cloudy indicatinga higher concentration of small particles must be present. The breakdown can be seen in fig.9

5.4 Absorbance and NTA

Absorbance and NTA was applied for all samples. The results for the absorbance measurements can be seen infig.10 and fig.11.

Figure 10: Absorbance for water with micro plastics after one and six minutes of blending.

The data shows that the absorbance became higher after longer blending time. This tells us that more fibresbreak up into smaller fragments and are suspended in the water. Note that the absorbance was measured in acuvette and the sample was handled with a 200 µl pipette so the larger pieces, visible to the naked eye, werenot included in the absorbance measurements.

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Figure 11: Absorbance for water with polystyrene after one and six minutes of blending.

For the breakdown of the PS the difference was very easy to see with the naked eye and in the absorbancemeasurements the absorbance became higher for all wavelengths. This is shown in fig.11. The peak in 230 nmis clear in both samples and the absorbance spectra is similar to those of the broken down MPs. As mentionedearlier the absorbance peak in 230 nm is relevant when looking for PS in a sample.

To see the size and concentration of the broken down plastic particles the samples were analysed in the NTA.The results can be seen in figures 12 and 13.

Figure 12: NTA for microplastics broken down for one and six minutes of blending.

The Results for the NTA of the microplastics (fig.12) is not as clear as the absorbance. For the MPs from thedigestive tract the concentration of particles around 120nm seems to be the same but the concentration for theslightly larger ones, around 220 nm, has increased some. For the MPs from the gills the small particles seems tohave increased some and a small population of particles around 400 nm is now present. The results from thisdiagram are not clear enough to draw any conclusions from.

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Figure 13: NTA for PS breakdown after 1 and 6 minutes, the results for the six minute sample is not reliable.

In the case of the PS (fig.13) the results do not seem reliable as the concentration of particles were clearly higherwhen the PS had been broken down for longer. However the results indicate that the concentration of particleswere lower after 6 minutes of breakdown. The reason might be that some larger particles were present after sixminutes and they spread more light than the small particles concealing a large portion of them. These particlesmight have been too big for the NTA to count them, as no population of larger particles could be seen in theresults. It might also be that there are particles smaller than 50 nm present that the NTA is unable to detect.

The microplastic fibres present many sources of potential errors as the handling was very difficult. Some pos-sible errors is that not all of the plastic fibres could be collected. This due to that some were too small or simplycould not be found. Some of the fibres collected might have been misdiagnosed for plastic when is was some-thing else. To limit these errors the filters have been compared to filters used for the control saline solution.

The section handling the microplastic fibres have been included as a reference to the particles suspected to benanoplastics. The section also highlights the problems in this type of analysis and aspects that are importantto think about when analysing microplastics.

5.5 Analysis of the water samples

The water samples collected from the fish contained an unknown range of different particles. To examine ifthere were any indications of the presence of nanoplastics a number of different methods of analysis wereapplied. How they were used and the results of each method are described in this chapter.

5.5.1 Assessment of particle populations

The first analysis were done with NTA and DLS to show an indication of how many particles there were in thesamples and which sizes were dominating.

NTA results show that particles were present in all the samples taken from fish. The concentration in the controlsamples, containing the saline solution, were too low to be able to measure in the NTA.

It was clear, in both DLS and NTA, that even though the filters used had a pore size of 1,2 µm, the particles thatfound were significantly smaller that that. The most commonly observed particles were between 100-250 nm.

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sample Particle size (nm) sample Particle size (nm)) sample Particle size (nm)DT1S 137.3 G1S — control S —DT1B 122.5 G1B — control B 293.9DT2S 183.1 G2S —DT2B 257.5 G2B —DT3S — G3S —DT3B 144.7 G3B —DT4S 259.3 G4S —DT4B 166.3 G4B —DT5S 173.4 G5S 214.3DT5B 189.0 G5B —DT6S — G6S 378.3DT6B 176.8 G6B —DT7S 176.3 G7S 442.0DT7B 193.1 G7B —

Table 3: DLS results from analyse of all untreated samples.

The DLS shows particles present for most of the DT samples and only three of the gill samples. The majority ofthe particles are between 100-300 nm.

Figure 14: NTA results for surface fraction of gill water samples diluted as 1:16.

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Figure 15: NTA results for bottom fraction of gill water samples diluted as 1:8.

Figure 16: NTA results for all digestive tract samples diluted as 1:8.

The results also showed that there were a difference between the water samples from the gills and digestivetract as well as for the surface and bottom fraction for the gill samples. A summary of the results can be seen infig.14, fig.15 and fig.16. This is likely due to the density separation causing the lower density particles to float tothe surface.

Note that the dilution differs between the figures, this is due to the original concentration of particles being toohigh to get good measurements for all samples.

The surface fractions of the gill samples contained the most particles and had to be diluted as 1:16 (fig.14). Forthe samples from the digestive tract a dilution with milliQ water of 1:8 was used(fig.16). In the bottom fractionof the digestive tract samples the concentration of particles were lower but the dilution of 1:8 could still be usedwith good results(fig.15).

The surface and bottom fraction of the gill samples were made into two different diagrams as the concentra-tions differed so greatly.

To attain more information about the material of the particles an attempt was made to use the FTIR on theuntreated water samples. Drops of water sample were deposited on the FTIR and allowed to dry to increasethe concentration of particles near the measuring crystal. This method was not successful in this case, likely

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due to the high concentration of salt in the samples. The drops dried very slowly and left salt crystals on themachine that affected the results. To be able to use this method the salt had to be separated from the particlesand particle concentration had to be increased. These methods will be handled further down in the report, seesection 5.5.4, Desalting the samples.

5.5.2 Density assessment

To get a better understanding of the density of the particles all samples from the digestive tract were centrifugedat different speeds. The rpm(rotations per minute) was set to 2000, 6000, 10000, 14000 and 18000 rpm for 10minutes each with a fraction of 100 µl taken out after each round for DLS analysis. These fractions were thenrun in the DLS plate reader and in NTA. The idea was to see how much gravity was needed to remove particlepopulations from the supernatant into the pellet.

DLS measurements of particle size (nm)SAMPLE/RPM 0 2000 6000 10000 14000 18000

1S 136,2 114,8 164 — 102,8 103,31B 126,6 125,1 119,3 101,2 — —2S 180,2 217,3 166,6 141 145,1 121,22B 199,5 217,9 147 144,5 119,7 111,33S 148,7 143,5 126,9 121,9 117,4 137,63B 162,3 128,5 127,2 133,4 116,7 108,84S — — 405,9 130,3 151 —4B 176,7 163,2 137,1 118,9 108,4 —5S 206,3 172,3 153,3 162,6 106,9 —5B 165,4 186,7 140,9 133,4 110,1 180,56S — 188,5 129,1 116,6 1860,8 —6B 180,2 155,1 133,5 117,9 105,5 —7S 149,3 152,3 177,8 119,2 787,8 3484,97B 219,9 176,5 137,5 167,2 — —

Table 4: DLS table from centrifugations for all speeds and fractions showing the particle size (nm).

The DLS results can be seen in table 4. This table shows that the effect of the centrifugation became clear at14000 and 18000 rpm. As particle populations are lost at high centrifugation speeds. High speed, i.e. gravita-tional force, is needed to force the particles to the bottom of the tube. This indicates low density of the particlesand this would support the hypothesis that they might be plastics.

The test was then repeated, for speeds 10000 rpm and 18000 rpm, for all digestive tract samples. This to confirmthe first results and to attain samples of the pellets and supernatants. For these centrifugations the pellet andsupernatant were tested in DLS and NTA. The pellet was diluted in 1 ml of Milli-Q water and sonicated for oneminute before analysis. The DLS results are shown in table.5.

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DLS measurements of particle size (nm)Rotations per minute(rpm) 10000 18000Sample original supernatant pellet supernatant pellet Average σ

1S 112,4 106,2 244 — 122,1 146 661B 157 118,2 110,7 — 125,9 128 202S 229,6 — 486,5 144,8 420,7 320 1602B — 263,9 413,3 120,5 430,4 307 1453S 270,8 182,7 1040,5 132,8 707,6 467 3933B 250,7 126,7 — 134,7 241,3 188 674S 454,6 159 426,3 991,1 178,6 442 3364B 172,3 135,7 251,8 107,3 161,5 166 545S 182,5 143,8 573 133,9 243,4 255 1835B 157,7 134,9 222,5 — 185 175 386S 631,8 148,6 304,1 — 247,3 333 2096B 204,4 1573,1 458,1 199,9 177,2 523 5987S 202,7 145,5 — — 163,9 171 297B 257 180,6 230,9 — 187,1 214 36

Table 5: Table of DLS results showing the particle size (nm) from the centrifugation runs on the digestive tract system forsupernatant and pellet for 10000 and 18000 rpm respectively with a calculated average and standard deviation(σ).

From table.5 we can see that most of the samples showed a population of larger particles in the pellet than inthe supernatant which is expected. From these results two samples were selected for further analysis usingNTA. To get the most information, one sample with little deviation was selected and one with large deviation.To fit this description and still show results for all wells in the DLS, the two samples chosen were 4B and 6B.

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Figure 17: NTA results from centrifugation runs at 10000 and 18000 rpm for pellet and supernatant for samples DT4B andDT6B. Note that the scale differ slightly between the figures, however the total concentration is not important in this compar-ison.

In fig.17 the results from NTA of samples 4B and 6B are shown. It can be confirmed from this table that thepellet has a larger part of the bigger particles. This was expected since they should be more affected by thecentrifugation than the smaller particles. There is a difference between the two samples, in sample DT4B thereare more particles left in the supernatant for centrifugation at the lower speed and for sample DT6B more areforced into the pellet.

To compare with the results from the DLS in table.5 the results for 4B are very close to the results of the NTA infig.17. One exception was the pellet for the 10000 rpm. Here the largest peak in the NTA is in 180 nm while theDLS shows 252 nm. This can be explained by a closer look at the NTA diagram. Several small peaks can be seenfor the pellet at a centrifugation of 10000 rpm. while they all show up in the NTA the DLS as a bias for largerparticles. The slightly smaller population of larger particles might overshadow the larger population of smallerparticles in the DLS as the larger particles scatter more light. This is a good example of why it is beneficial touse both methods to get a more comprehensive understanding of the samples.

Doing the same comparison for sample 6B, greater deviations can be seen. The peaks consistently show up forsmaller particles in the NTA than in the DLS. One exception is for the 18000 rpm pellet, where they seem to

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match. The DLS does have a bias towards larger particles as explained before but these differences seem quitelarge in this case. This difference could possibly be due to a density difference causing the particles to scattermore light than less dense particles.

5.5.3 Absorbance analysis

As a next step in analysing the samples the absorbance was measured.

Figure 18: Absorbance for a saline solution sample with a clear peak in 230nm.

Figure 19: Absorbance for sample DT1S.

From previous research it is known that polystyrene has a strong absorbance peak in 230 nm [6] [20], thereforesome of the water samples with high particle concentration were tested for absorbance. The peak was presentfor all samples but also other peaks. For a control sample of the saline solution used to filter the fish the peakin 230 nm was very clear and with a narrow distribution, see figures 18 and 19. It was therefore important toseparate the particles in the samples from the salt.

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5.5.4 Desalting the samples

It was important to separate the particles from the salt. This to, in the absorbance measurements, be able toidentify which peaks are from the salt and which ones are from the samples in the absorbance measurements.

To achieve this a PD10 column was used for desalting. As a trial for the desalting one samples with high particleconcentration was used, G1S. In the PD10 column 0,5 ml of sample was added and eluted with 10ml of Milli-Qwater, 10 fractions of 1 ml each were collected and all fractions were tested in DLS, NTA and for absorbance.

The DLS reader showed reliable particle populations for fraction three and four and this was confirmed in theNTA with no particles visible in any other fractions. The absorbance peaked in fractions three and four butalso for seven, eight and nine. This is suspected to be some pollution in the salt causing the peaks in the laterfractions which indicates a successful separation from the salt.

More DT and gill samples, as well as saline solution samples, were desalted in the PD10 column following thesame protocol. Further analysis of these samples showed similar results with no peak visible in fractions threeor four for the saline solution.

Figure 20: A summary for the wavelength 230nm for each fraction in six different samples from digestive tract and gills.

The absorbance for the 230 nm wavelength were summarised for each series of samples to look for a corre-lation, the results are shown in fig.20. This points at a peak in fraction three and one in fraction nine. Theabsorbance is also significant for fractions four and eight. The corresponding figure for the saline solution,fig.21, shows the peak in fraction nine but the peak in fraction three and four is clearly not comparable to thosein sample DT4B or G3S. These two samples are therefore chosen for further analysis by repeating the desaltingand the absorbance analysis. These results are shown in fig.22 and fig.23.

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Figure 21: A summary for the wavelength 230 nm for each fraction of sample of saline solution run trough a PD10 column.

Figure 22: A summary for the wavelength 230 nm for each fraction in six samples of G3S run trough a PD10 column.

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Figure 23: A summary for the wavelength 230 nm for each fraction in six samples of DT4B run trough a PD10 column.

These results are consistent and agrees well with the hypothesis that the first peak is from the particles and thesecond one is from a pollution from the salt, both these peaks could be from PS but is not proof that it must be.

For a simple comparison of the absorbance the results of a fraction four for one of the samples (DT1S) is plottedagainst the absorbance for a water sample only containing pure PS in fig.24.

Figure 24: Absorbance comparison of a digestive tract sample fraction four and pure PS broken down in blender for oneminute. The absorbance have been adjusted by a factor of 2,5 to bring the lines closer to each other for easier comparison.

The sample absorbance have been adjusted by a factor of 2,5 to bring the graphs closer for easier comparison.The graphs are very similar in shape and peaks.

5.5.5 Concentrating the particles

To be able to make a new attempt to identify the material of the particles in FTIR, fractions three and four fromeach repetition were pooled into two test tubes. One of the tubes is for all fraction three and four for DT4B andone for G3S. These are then freeze dried to get rid of the water and thereby concentrating the particles. Thesame was done for saline solution to have a comparison.

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Figure 25: 50 ml tubes containing freeze dried samples from saline solution, gills and digestive tract in order from left to right,the lower pictures 50µ l of water have been added to each sample.

After freeze-drying 50 ml of each sample a dry matter was obtained, this was then re suspended in 50 µl ofMilli-Q water using a pipette. The remains after the removal of water trough freeze-drying can be seen in fig.25,the figure also shows the particles resuspended in 50 µl of water. It is clear that a lot more matter is collectedfrom the fish samples than from the saline solution. The samples were then tested in absorbance, NTA andFTIR. The results of the absorbance analysis can be seen in fig.26, the results of the NTA can be seen in fig.27and the result of the FTIR can bee seen in fig. 28.

Figure 26: Absorbance for concentrated samples of collected fraction 3 and 4 for six different column runs for digestive tract,gills and saline solution respectively.

The absorbance measurements gave a high, clear peak in 230 nm for both digestive tract and gills. The saline

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solution showed very low over all absorbance. These results are shown in fig.26. This proves that the particlesobtained, and the absorbance peak, is not from the salt but from the samples.

Figure 27: NTA for the freeze dried samples of fraction 3 and four pooled together for respective samples.

The NTA shows high concentration of particles around 200 nm for both gill and DT samples. The concentrationof particles in the control saline solution is in comparison insignificant.

From both figures 26 and 27 it is clear that the particles present in fraction three and four for the samples is notpresent in the same fractions for the saline solution. These particles must be from the fish and could potentiallybe plastics.

Figure 28: FTIR graph for concentrated samples of gill, digestive tract(DT) and saline solution. The number after the sampleis the scan number.

The samples were again analysed using FTIR, the results of this is shown in fig.28. By comparing these spectrasto plastic reference spectras[21] it was not possible to see any perfect match. It is therefor not possible toconfirm that they contain PS or any other plastic.

When analysing on the nano scale it is important to remember that everything contains nano particles, theycan even be found in Milli-Q water, therefore precautions have been taken by always having controls to accountfor these contaminations. The measurements can still be a bit off however as the results often differ betweenruns. There is always the possibility to repeat tries and measurements more times to get higher accuracy. Forthis report there was limited time and sample volumes, therefore results of analysis have been used more asindications and no hard conclusions can been drawn.

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6 Future research

More research and experimental work is needed in the field of nanoplastics. Especially on how to isolate themand decide what material they are made from. This work is a significant step on the way to find a good method-ology for these tasks. There is also much work to do in the field of finding the biological implications of NPs aswell as how they break down and act in more complex systems such as the ocean or in soil.

One thing that have been avoided in this thesis is using chemicals to break down the organic matter in thesamples. This in fear that they will affect the nanoplastics in ways we do not know. However this is a possibilitythat would be interesting to study if more in future research.

7 Conclusions

From this work it has been proven that it is possible to isolate plastic-like particles, on the nano scale, from acomplex mixture. The absorbance measurements gave a strong indication that nanoplastics were present inthe fish, both in gills and digestive tract.

From all analysis made on the water samples, no results from the particles isolated, differed from the propertiesof nanoplastics. Therefore it is likely that the protocol of filtration, NTA, DLS, desalting and freeze-drying canbe used to attain nanoplastic samples.

The presence of microplastics was confirmed in both the gills and the digestive tract of all fish analysed. Themechanical breakdown of microplastic fibres and a polystyrene coffee cup lid also released nanoplastics inmechanical breakdown. This suggests that nanoplastics might be very common.

It was not possible, with a 100 percent certainty, to say that nanoplastics were found in the fish, further analysisto confirm the material is needed.

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References

[1] C.G. Mineo A. Lattin G.L. Magaldi M.G. Belmonte G. Moore C.J. Regoli F. Aliani S. Suaria, G Avio. Themediterranean plastic soup: synthetic polymers in mediterranean surface waters. Scientific Reports,6:37551, 2016.

[2] J. Gigault, A. ter Halle, M. Baudrimont, P. Pascal, F. Gauffre, T.-L Phi, H. El Hadri, B. Grassl, and S. Reynaud.Current opinion: What is a nanoplastic? Environmental Pollution, 235:1030 – 1034, 2018.

[3] H. Abrahamson. 1900 små syntetiska partiklar lossnar från en enda fleecetröja vid tvätt och föl-jer med avloppsvattnet ut i havet. Ny Teknik, 2011. https://www.nyteknik.se/innovation/fleecetrojan-fororenar-haven-vid-tvatt-6419543.

[4] L.M. Hernandez, N. Yousefi, and N. Tufenkji. Are there nanoplastics in your personal care products? En-vironmental Science & Technology Letters, 4:280–285, 2017.

[5] S. Lambert and M. Wagner. Characterisation of nanoplastics during the degradation of polystyrene.Chemosphere, 145:265 – 268, 2016.

[6] M.T. Ekvall, M. Lundqvist, E. Kelpsiene, E. Šileikis, S.B. Gunnarsson, and T. Cedervall. Nanoplastics formedduring the mechanical breakdown of daily-use polystyrene products. Nanoscale Adv., 1:1055–1061, 2019.

[7] Koelmans A.A., Besseling E., Shim W.J. Nanoplastics in the Aquatic Environment. Critical Review. In:Bergmann M., Gutow L., Klages M. (eds) Marine Anthropogenic Litter. Springer, Cham, 2015.

[8] GESAMP . “Sources, fate and effects of microplastics in the marine environment: a global assessment”(Kershaw, P. J., ed.). (IMO/FAO/UNESCO-IOC/UNIDO/WMO/IAEA/UN/UNEP/UNDP Joint Group of Ex-perts on the Scientific Aspects of Marine Environmental Protection). Rep. Stud. GESAMP No. 90, 96 p.,2015.

[9] K. Mattsson, L.-A. Hansson, and T. Cedervall. Nano-plastics in the aquatic environment. Environ. Sci.:Processes Impacts, 17:1712–1721, 2015.

[10] A. L. Lusher, N. A. Welden, P. Sobral, and M. Cole. Sampling, isolating and identifying microplastics in-gested by fish and invertebrates. Anal. Methods, 9:1346–1360, 2017.

[11] E.V. Malmendal A. Linse S. Hansson L. Cedervall T. Mattsson, K. Johnson. Brain damage and behaviouraldisorders in fish induced by plastic nanoparticles delivered through the food chain. Scientific Reports,7(1).

[12] C. Whalquist. Krill found to break down microplastics – but it won’t save the oceans.The Guardian, 2018. https://www.theguardian.com/environment/2018/mar/12/krill-can-turn-microplastics-into-nanoplastics-study.

[13] E. Flashman. How plastic-eating bacteria actually work – a chemist explain. The Guardian, 2018. http://theconversation.com/how-plastic-eating-bacteria-actually-work-a-chemist-explains-95233.

[14] L.M. Rios Mendoza, H. Karapanagioti, and N. Ramírez Álvarez. Micro(nanoplastics) in the marine envi-ronment: Current knowledge and gaps. Current Opinion in Environmental Science & Health, 1:47 – 51,2018.

[15] United Nations Industrial Development Organization (UNIDO). The pop priority: Unido and the stock-holm convention. Vienna International Centre · P.O. Box 300 · 1400 Vienna · Austria.

[16] C.-B. Jeong, H.-M. Kang, Y.H. Lee, M.-S. Kim, J.-S. Lee, J.S. Seo, M. Wang, and J.-Se. Lee. Nanoplastic inges-tion enhances toxicity of persistent organic pollutants (pops) in the monogonont rotifer brachionus kore-anus via multixenobiotic resistance (mxr) disruption. Environmental Science & Technology, 52(19):11411–11418, 2018. PMID: 30192528.

[17] Malvern Panalytical. Nanoparticle tracking analysis (nta) and dynamic light scattering (dls) - comparisonbetween nta and dls, 2009.

[18] R. Dris, H.K. Imhof, M.G.J. Löder, J. Gasperi, C. Laforsch, and B. Tassin. Chapter 3 - microplastic contami-nation in freshwater systems: Methodological challenges, occurrence and sources. In Eddy Y. Zeng, editor,Microplastic Contamination in Aquatic Environments, pages 51 – 93. Elsevier, 2018.

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[19] GE Healthcare. PD-10 Desalting Columns-Instructions 52-1308-00 BB, September 2007.

[20] S. M. Mintenig, P. S. Bäuerlein, A. A. Koelmans, S. C. Dekker, and A. P. van Wezel. Closing the gap betweensmall and smaller: towards a framework to analyse nano- and microplastics in aqueous environmentalsamples. Environ. Sci.: Nano, 5:1640–1649, 2018.

[21] M.R. Jung, F. D. Horgen, S.V. Orski, V. Rodriguez C., K. L. Beers, G. H. Balazs, T. T. Jones, T.M. Work, K.C.Brignac, S. Royer, K. D. Hyrenbach, B.A. Jensen, and J.M. Lynch. Validation of atr ft-ir to identify polymersof plastic marine debris, including those ingested by marine organisms. Marine Pollution Bulletin, 127:704– 716, 2018.

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