Toxicity of Microplastics to Aquatic Organisms

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Toxicity of Microplastics to Aquatic OrganismsSarah AuClemson University, [email protected]

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TOXICITY OF MICROPLASTICS TO AQUATIC ORGANISMS

A Dissertation Presented to

the Graduate School of Clemson University

In Partial Fulfillment of the Requirements for the Degree

Doctor of Philosophy Environmental Toxicology

by Sarah Au May 2017

Accepted by: Dr. Cindy M. Lee, Committee Chair

The late Dr. Stephen J. Klaine Dr. Peter H. Adler

Dr. William S. Baldwin Dr. Peter van den Hurk Dr. John E. Weinstein

ii

ABSTRACT

Plastic is used in products that span from cutlery to the main components in

automobiles, and airplanes. Both municipal solid waste streams and marine debris are

comprised of largely plastic products, and as annual global plastic production and use is

unlikely to decrease, the presence of plastic in the environment will likely increase.

Microplastics (MPs) are plastic fragments that have at least one dimension that is less

than 5 mm, and have been found throughout water bodies around the world. MPs are in

the same size fraction as most sediment particles and microorganisms, allowing them to

be easily ingested by a variety of organisms. The potential for MPs to alter the

bioavailability of contaminants, and whether MPs and adsorbed contaminants can

undergo trophic transfer, have been investigated in very few studies. Fluoranthene (FLU)

is one of the US Environmental Protection Agency’s 16 priority polycyclic aromatic

hydrocarbons; and its hydrophobic nature allows it to sorb easily to sediment and organic

matter, which suggests FLU will sorb to the hydrophobic surfaces of most MPs. While

most studies conducted have focused on acute MP exposures to marine organisms, little

is known regarding the toxicological chronic effects of MP exposures. Recent studies

have demonstrated that MP concentrations in freshwater water bodies are comparable to

those found in the ocean; even less information is available regarding MP toxicity to

freshwater organisms.

The first objective of my research was to characterize the physical toxicity of MPs

to aquatic invertebrates and fish. After establishing a baseline for acute MP toxicity, the

second objective focused on the determination of whether fluoranthene that is adsorbed to

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MP surfaces is bioavailable to aquatic invertebrates and fish. Finally, the third objective

evaluated whether MPs and the adsorbed fluoranthene can undergo trophic transfer

between prey and predatory organisms in both freshwater and marine ecosystems.

My results demonstrated that acute exposures to polyethylene spherical MP do

result in mortality at environmentally relevant concentrations. Specifically, 10- and 7-

day LC50s of approximately 4.6 x 104 and 7.9 x 104 MPs/mL for spherical polyethylene

MPs were quantified for amphipods and copepods, respectively. No mortality was

observed for either fathead minnows or mummichogs at MP exposures used during these

bioassays. Interestingly, polypropylene MP fibers were more toxic to exposed

amphipods than the polyethylene MP spheres, where mortality was observed at

concentrations as low as > 22.5 MPs/mL. When polyethylene spherical MPs are

contaminated with FLU, acute exposures can result in an increase in FLU bioavailability

to both fish and invertebrates. Chronic polyethylene spherical MP exposures resulted in

significant growth reductions in both Hyalellla azteca and Amphiascus tenuiremis, and

decreased reproduction in exposed H. azteca. In comparison to polyethylene spherical

MPs, fibrous polypropylene MP exposures to H. azteca resulted in significantly greater

mortality and reductions in both growth and reproduction. Exposure to FLU-

contaminated MPs resulted in an increase in FLU bioavailability to invertebrates and fish,

where more than 90% of the invertebrate FLU body burdens were due to MP exposure.

Finally, my results demonstrated that FLU that was adsorbed to MP surfaces can undergo

trophic transfer using both freshwater and marine invertebrates and fish. Approximately

1.5 mg fluoranthene was bioavailable to fathead minnows and mummichogs that had

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ingested amphipods, and copepods that had been pre-exposed to fluoranthene-

contaminated MPs, respectively. Hybrid striped bass that has consumed fathead

minnows that were pre-exposed to fluoranthene-contaminated MPs resulted in an average

bile concentration of approximately 1.3 mg fluoranthene. MPs represent another route of

exposure for ubiquitous persistent organic pollutants in the environment for lower and

higher trophic level organisms.

My research provided a platform on which the acute and chronic toxicological

effects due to MP exposures of freshwater and marine invertebrates and fish can be

compared because similar methods were used in all the bioassays. Generating data that

resulted from the use of similar exposure methods allows the scientific community to

make cross-species comparisons that are difficult to determine with the currently

available microplastic toxicity data. My results demonstrate that microplastics can have

both acute and chronic toxicological effects on aquatic organisms, and that the presence

of microplastics in the environment needs to be taken into consideration when evaluating

the exposure to environmental contaminants.

v

DEDICATION

I dedicate my dissertation to my family and friends.

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ACKNOWLEDGMENTS

I cannot express how thankful I am to have had the opportunity to study under my

PhD adviser, Dr. Stephen J. Klaine. It is easy to say that he was a great teacher and

mentor, but it is a rare gift to be able to say that one’s adviser was also a friend, father-

figure, and colleague. The science that I learned from him will carry me through my

future career decisions, and the people I have met through him are not just professional

acquaintances, but are some of the most amazing friends one could ever have.

Professionally and personally, I am a better scientist, and person because of Steve Klaine.

Hopefully I am a little better at seeing one tree at a time, and not just the whole forest.

My committee members also deserve a round of applause. Everyone has

contributed to being a solid support system that I hesitate to think of what the last year

would have been like without them. I wholeheartedly thank Dr. Cindy M. Lee for being

an amazing mentor and friend. You have been a steadying presence in my life for the

past year, and I don’t think I could have finished my dissertation without your help.

Thank you for taking the time to correct all my writing errors. This may seem trivial, but

I “believe” I am a better writer now. Any future collaborator or correspondent will have

you to thank as well. I am grateful that through Sea Grant, and my dissertation work, I

was given the opportunity to collect microplastics in Charleston and Georgetown, SC,

with Dr. John Weinstein and his students. Getting a different perspective on microplastic

sampling techniques made me more aware of the various abiotic and biotic processes that

influence contaminant transport and fate in the environment. Thank you for your

guidance at my platform presentations. It is always good to have a friendly face around.

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A special thanks to Dr. Peter Adler for opening his lab to me, so that I could develop

better methods to image my invertebrates. I believe that our discussions on science as a

whole and how my work is influenced by other worldly events has given me a better view

on how to connect with individuals that do not have the same research background.

Thank you for taking the blinders off occasionally, and giving me a chance to truly reflect

on different aspects of my research. I am grateful that Dr. William Baldwin has also

opened his lab for me, so that I could explore different methods to better extract

fluoranthene from my tissue samples. Thank you for allowing me to use some of the

instruments in your lab. Finally, a special thank you to Dr. Peter van den Hurk for being

a mentor throughout my five years as a graduate student, and becoming a committee

member for me in my last year. Thank you for also opening your lab to me, and helping

me with method development.

I would like to acknowledge and thank Ron Gossett, Norm Ellis, and John Smink

for their assistance with animal culture and maintaining facilities at both the Institute of

Environmental Toxicology and Cherry Farm. None of my work could have been

accomplished without you because you were always helping us whenever an extra hand

was needed. ‘Thank you’ is a phrase that does not quite amount to the gratitude I feel.

I would also like to thank all past and present Klaine lab members. The research

projects that everyone had are so different, and I believe that I benefitted from having so

many types of methods and perspectives on toxicology surrounding me every day. Thank

you to Kim Newton, Austin Wray, Lauren Sweet, Anna Lee McLeod, Katherine Johnson-

Couch, Erica Linard, Lauren Stoczynski, Jason Coral, and Chad Mansfield for helping

viii

with fish culture, lab inspection cleanings, and giving me a support system throughout the

last five years. A special thanks to Lauren Sweet and Chad Mansfield for helping me

count copepods during a few bioassays.

I would also like to thank Dr. Tom Chandler and Emily Stewart at the University

of South Carolina for providing me with copepods to use in my toxicity bioassays.

Thank you, Paul Kenny, for assisting us in the collection of pluff mud and mummichogs

at the Belle W. Baruch Institute of Coastal Ecology and Forest Science. There are many

experiments that would not have been conducted if you had not helped me on so many

occasions. Thank you, Austin Gray, for helping me on countless field collection trips in

Charleston and Georgetown, SC. I am so glad that we were able to become both

colleagues and friends through these trips and various conference meetings. I would also

like to thank Dana Szymkowicz and Kaleigh Sims from Dr. Lisa Bain’s lab for helping

me with mummichog collection and embryo fertilization. Thank you, Drs. Lisa Bain and

Charlie Rice, for helping me develop methods for various experiments in the last few

years, and giving me advice regarding mummichog collection and culture.

Establishing the What’s in Our Water Jr. Program would not have been possible

without the help of Janine Sutter, Cathleen Reas, Laura Nash, and the numerous mentors

from Clemson University that helped make science fun for elementary students.

Participating in this program has given me a refreshing outlook on my research and a

different platform on which to relate my research to the interests of people in different

age groups and personal interests.

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Finally, a huge thank you to my mother, father, brother, and sister. Your patience

with my schedule has helped me get through the last few years and I am so glad to finally

be back in the Maryland-DC area so that the time between visits are not as long. Thank

you to all my friends that are scattered between the two coasts for taking the time to

maintain our friendships despite everyone having such crazy schedules. I would also like

to thank Jacob for making the multiple trips to South Carolina every year so that we can

explore various cities in the south. Thank you for listening to all my problems, and

helping me cope with all the stress. A general thank you to everyone I have met during

my five years at Clemson University. Your patience, guidance, and friendship though out

this whole experience has made this dissertation possible.

x

TABLE OF CONTENTS

Page

TITLE PAGE .................................................................................................................... i ABSTRACT ..................................................................................................................... ii DEDICATION ................................................................................................................. v ACKNOWLEDGMENTS .............................................................................................. vi LIST OF TABLES ......................................................................................................... xv LIST OF FIGURES ...................................................................................................... xvi CHAPTER I. LITERATURE REVIEW .............................................................................. 1 1.1 Introduction ........................................................................................ 1 1.2 Plastic disposal and entry into the environment ................................ 5 1.3 Plastic degradation and biofouling ..................................................... 6 1.3.1 Polymer degradation ....................................................................... 6 1.3.2 Biofouling of plastics ...................................................................... 9 1.4 Factors that influence microplastic exposure, ingestion, and trophic transfer .......................................................................... 13 1.4.1 Microplastic size, shape, and concentrations ................................ 13 1.4.2 Microplastic polymer composition ............................................... 19 1.4.3 Regional climate patterns .............................................................. 23 1.4.4 Uptake and depuration .................................................................. 24 1.5 Microplastic toxicity to aquatic organisms ...................................... 26 1.5.1 Energy reserves and growth .......................................................... 31 1.5.2 Reproduction ................................................................................. 30 1.5.3 Bioavailability of additives, adsorbed environmental contaminants, and viruses ..................................... 33 1.5.4 Trophic transfer ............................................................................. 36 1.5.5 Population-level effects ................................................................ 40 1.6 Dissertation goals and objectives ..................................................... 43 1.7 References ........................................................................................ 45 II. MICROPLASTIC PHYSICAL TOXICITY TO FRESHWATER AND MARINE INVERTEBRATES AND FISH ............ 61

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Table of Contents (Continued) 2.1 Abstract ............................................................................................ 61 2.2 Introduction ...................................................................................... 62 2.3 Methods............................................................................................ 68 2.3.1 Organism culture ........................................................................... 68 Freshwater amphipod Hyalella azteca ................................................... 68 Marine copepod Amphiascus tenuiremis ............................................... 69 Freshwater fish Pimephales promelas ................................................... 69 Marine fish Fundulus heteroclitus ......................................................... 70 2.3.2 Microplastic exposure water preparation ...................................... 72 Polyethylene microplastic spheres ......................................................... 72 Polypropylene microplastic fibers (Hyalella azteca only)..................... 73 2.3.3 Toxicity Bioassays ........................................................................ 74 Freshwater amphipod Hyalella azteca ................................................... 74 Marine copepod Amphiascus tenuiremis ............................................... 77 Freshwater fish Pimephales promelas ................................................... 79 Marine fish Fundulus heteroclitus ......................................................... 79 2.3.4 Microplastic ingestion analysis ..................................................... 80 Freshwater amphipod Hyalella azteca & marine copepod Amphiascus tenuiremis ....................................... 80 Freshwater fish Pimephales promelas & marine fish Fundulus heteroclitus ................................................ 81 2.3.5 Microplastic egestion analysis ...................................................... 82 2.3.6 Growth and reproduction analysis ................................................ 83 Freshwater amphipod Hyalella azteca ................................................... 83 Marine copepod Amphiascus tenuiremis ............................................... 83 Freshwater fish Pimephales promelas & marine fish Fundulus heteroclitus ................................................ 84 2.3.7 Data analysis ................................................................................. 84 2.4 Results .............................................................................................. 85 2.4.1 Freshwater amphipod Hyalella azteca .......................................... 85 Acute toxicity of microplastics: polyethylene spherical MPs and polypropylene fibrous MPs ............................ 85 Chronic toxicity of microplastics: polyethylene spherical MPs and polypropylene fibrous MPs ............................ 86 2.4.2 Marine copepod Amphiascus tenuiremis ...................................... 89 Acute toxicity of polyethylene spherical MPs ....................................... 89 Chronic toxicity of polyethylene spherical MPs .................................... 90 2.4.3 Freshwater fish Pimephales promelas .......................................... 91 2.4.4 Marine fish Fundulus heteroclitus ................................................ 91 2.5 Discussion ........................................................................................ 92 2.6 Limitations ..................................................................................... 102 2.7 Conclusions .................................................................................... 103

xii

Table of Contents (Continued) 2.8 Figures............................................................................................ 104 2.9 References ...................................................................................... 120 III. TOXICITY OF FLUORANTHENE-CONTAMINATED MICROPLASTICS TO FRESHWATER AND MARINE INVERTEBRATES AND FISH ............................................... 126 3.1 Abstract .......................................................................................... 126 3.2 Introduction .................................................................................... 127 3.3 Methods.......................................................................................... 132 3.3.1 Characterization of fluoranthene adsorption to microplastics ......................................................... 132 Adsorption isotherm.................................................................... 132 Isotherm modeling ...................................................................... 133 3.3.2 Fluoranthene-contaminated microplastics bioassays .................. 135 3.3.2.1 Freshwater amphipod Hyalella azteca ..................................... 135 Fluoranthene-contaminated polyethylene microplastic exposure .............................................................. 135 Water-borne fluoranthene exposure (No microplastics) .................................................................... 136 3.3.2.2 Marine copepod Amphiascus tenuiremis ................................. 138 Fluoranthene-contaminated polyethylene microplastic exposure .............................................................. 138 Water-borne fluoranthene exposure (No microplastics) .................................................................... 139 3.3.2.3 Freshwater fish Pimephales promelas ..................................... 140 Fluoranthene-contaminated polyethylene microplastic exposure .............................................................. 140 Water-borne fluoranthene exposure (No microplastics) .................................................................... 141 3.3.2.4 Marine fish Fundulus heteroclitus ........................................... 142 Fluoranthene-contaminated polyethylene microplastic exposure .............................................................. 142 Water-borne fluoranthene exposure (No microplastics) .................................................................... 143 3.3.3 Fluoranthene bioavailability quantification ................................ 144 3.3.4 Data analysis ............................................................................... 145 3.4 Results ............................................................................................ 146 3.4.1 Fluoranthene adsorption to polyethylene microplastics ............. 146 3.4.2 Freshwater amphipod Hyalella azteca ........................................ 147 3.4.3 Marine copepod Amphiascus tenuiremis .................................... 150 3.4.4 Freshwater fish Pimephales promelas ........................................ 152

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Table of Contents (Continued) 3.4.5 Marine fish Fundulus heteroclitus .............................................. 153 3.5 Discussion ...................................................................................... 153 3.6 Limitations ..................................................................................... 159 3.7 Conclusions .................................................................................... 160 3.8 Figures............................................................................................ 162 3.9 References ...................................................................................... 175 IV. MICROPLASTIC TROPHIC TRANSFER ............................................... 180 4.1 Abstract .......................................................................................... 180 4.2 Introduction .................................................................................... 181 4.3 Methods.......................................................................................... 188 4.3.1 Microplastic exposure preparations ............................................ 188 4.3.2 Microplastic trophic transfer from Hyalella azteca to Pimephales promelas .................................... 189 Group and individual training sessions ....................................... 189 Hyalella azteca exposure to polyethylene microplastics with and without fluoranthene ........................ 191 3-d Trophic transfer experiment ................................................. 192 Bile analysis ................................................................................ 192 4.3.3 Microplastic trophic transfer from Pimephales promelas to Morone chrysops x Morone saxatilis .................................... 193 Morone chrysops x Morone saxatilis culture ............................... 193 Pimephales promelas preparation ................................................ 194 Group and individual training sessions ....................................... 194 Pimephales promelas exposure to polyethylene microplastics with and without fluoranthene ........................ 195 1-d Trophic transfer experiment .................................................. 196 Bile analysis ................................................................................. 197 4.3.4 Microplastic trophic transfer from Amphiascus tenuiremis to Fundulus heteroclitus............................................................ 197 Group and individual training sessions ....................................... 197 Amphiascus tenuiremis exposure to polyethylene microplastics with and without fluoranthene ........................ 199 3-d Trophic transfer experiment .................................................. 200 Fluoranthene body burden analysis ............................................. 200 4.3.5 Data analysis ............................................................................... 201 4.4 Results ............................................................................................ 202 4.4.1 Microplastic trophic transfer from Hyalella azteca to Pimephales promelas .................................... 202

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Table of Contents (Continued) 4.4.2 Microplastic trophic transfer from Pimephales promelas to Morone chrysops x Morone saxatilis .......................................... 202 4.4.3 Microplastic trophic transfer from Amphiascus tenuiremis to Fundulus heteroclitus ....................... 203 4.5 Discussion ...................................................................................... 203 4.6 Limitations ..................................................................................... 208 4.7 Conclusions .................................................................................... 209 4.8 Figures............................................................................................ 211 4.9 References ...................................................................................... 214 V. CONCLUSIONS........................................................................................ 219 5.1 Future research needs ..................................................................... 222 5.2 References ...................................................................................... 226

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LIST OF TABLES

Table Page 1.1 Environmentally relevant concentrations of microplastics in various field studies ....................................................... 18 2.1 Summary of microplastic acute and chronic exposure concentrations for both freshwater and marine invertebrates and fish ................................................................ 77 2.2 Ramped Diet Used in 10-day and 42-day Hyalella azteca exposures ............................................................... 75 3.1 Hyalella azteca acute exposure to fluoranthene- contaminated microplastics (FLU-MPs) and chronic exposure to only water-borne fluoranthene (FLU) .................................... 137 3.2 Amphiascus tenuiremis acute exposure to fluoranthene- contaminated microplastics (FLU-MPs) and chronic exposure to only water-borne fluoranthene (FLU) .................................... 140 4.1 Microplastic exposure concentrations (#MPs/mL) and number of prey organisms fed to prey and predatory organisms, respectively ............................................... 189

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LIST OF FIGURES

Figure Page 2.1 Mortality and polyethylene microplastic (MP) ingestion following 10-d acute exposure ............................................................. 104 2.2 Polypropylene MP fiber ingestion, mortality, growth, and egestion following 10-d acute exposure ........................................ 105 2.3 Polyethylene MP ingestion, growth, and reproduction Following 42-d chronic exposure ........................................................ 106 2.4 Images of fluorescent polyethylene MP spherical particles (10-27 µm in diameter) ingested by H. azteca ...................... 108 2.5 Mortality following 42-d chronic exposure to polypropylene fibrous MPs .................................................................. 109 2.6 Growth following 42-d chronic exposure to polypropylene fibrous MPs ........................................................................ 110 2.7 Reproduction following 42-d chronic exposure to polypropylene fibrous MPs ........................................................................ 111 2.8 Gut residence time of polypropylene MP fibers during 42-d chronic exposure .................................................................... 112 2.9 Mortality following 7-d acute exposure to polyethylene MPs....................................................................................... 113 2.10 Polyethylene MPs ingested throughout 32-d chronic exposure ............................................................................................... 114 2.11 Images and growth resulting from the ingestion of fluorescent polyethylene MPs by nauplii and adult copepods ..................................................................................... 115 2.12 Reproduction following 32-d exposure to polyethylene MPs....................................................................................... 117 2.13 Ingestion of polyethylene MPs following 4-d acute exposure to fathead minnows ..................................................... 118

xvii

List of Figures (Continued) 2.14 Ingestion of polyethylene MPs following 4-d acute exposure to mummichogs .......................................................... 119 3.1 Adsorption isotherm of fluoranthene adsorbed onto polyethylene microplastics ................................................................ 162 3.2 Images taken of polyethylene microplastics with and without fluoranthene ................................................................... 163 3.3 Mortality following 42-d chronic exposure to Fluoranthene and 14-d depuration period .................................................. 164 3.4 Fluoranthene body burden following 10-d acute exposure to fluoranthene-contaminated microplastics and water-borne fluoranthene .............................................................. 165 3.5 Fluoranthene body burden following 10-d acute exposure to water-borne fluoranthene ....................................................... 167 3.6 Fluoranthene body burden following 42-d chronic exposure to fluoranthene and 14-d depuration period ............................... 168 3.7 Reproduction following 42-d chronic exposure to Fluoranthene and 14-d depuration period .................................................. 169 3.8 Fluoranthene body burden following 7-d acute exposure to fluoranthene-contaminated microplastics and water-borne fluoranthene .............................................. 170 3.9 Fluoranthene body burden following 7-d acute exposure to water-borne fluoranthene ....................................................... 171 3.10 Comparison of fluoranthene body burden between copepods and amphipods ............................................................. 172 3.11 Fluoranthene body burden following 4-d acute exposure to fluoranthene-contaminated microplastics and water-borne fluoranthene .............................................. 173

xviii

List of Figures (Continued) 3.12 Fluoranthene body burden following 4-d acute exposure to fluoranthene-contaminated microplastics and water-borne fluoranthene .............................................. 174 4.1 Fluoranthene in the bile of fathead minnows due to trophic transfer ................................................................................ 211 4.2 Fluoranthene in the bile of hybrid striped bass due to trophic transfer ........................................................................ 212 4.3 Fluoranthene whole body burden of mummichogs due to trophic transfer ......................................................... 213

1

CHAPTER ONE

LITERATURE REVIEW

1.1 Introduction

As an innovative product of the 20th century, worldwide plastic production

reached 322 million metric tons in 2015 (Statista 2015). Plastic is a synthetic organic

polymer that can have highly variable chemical and physical properties that influence its

use, durability, and eventual presence in the environment (Hidalgo-Ruz et al. 2012). The

plastics industry is involved with products used in many sectors of society including

medicine, energy generation, aerospace, automotive, construction, electronics, packaging,

and textiles (PlasticsEurope 2015). Polyolefins (i.e. polyethylene and polypropylene),

polyvinyl chloride (PVC), polystyrene (PS) solid, expandable polystyrene, and

polyethylene terephthalate (PET) account for about 80% of the total global demand

(Plastics Europe 2105). Furthermore, packaging (39.5%) and construction (20.1%) are

the main plastic applications in which plastics are produced worldwide (Plastics Europe

2015). Nearly a quarter (22.7%) of plastic production is used in sectors involved with the

production of consumer and household appliances, furniture, sport, and health and safety

equipment (Plastics Europe 2015). Polyethylene and polypropylene, followed by PET,

polymethyl methacrylate (PMMA), and nylon are used most prominently in health

products (Leslie 2012). As additional uses for plastic continue to be discovered, plastic

production is predicted to steadily increase (PlasticsEurope 2015), which may account for

increasing amounts of microplastics found in marine sediment (Claessens et al. 2011).

2

While the presence and toxicity of larger fragments of plastic debris have been

well documented in marine ecosystems, and higher trophic level organisms, respectively,

there is little information readily available on whether smaller fragments of plastic debris

pose an ecological and human health threat (Gregory and Ryan 1997; Derraik 2002; Ryan

1988; Laist 1987; Fry et al. 1987; Bugoni et al. 2001; Baird and Hooker, 2000).

Microplastics (MPs) have been generally defined as plastic particles that have at least one

dimension less than 5 mm (Andrady 2011; Barnes et al. 2009), and can be further

classified by the intent in which the microplastics were formed. Nanoplastics (if defined

as nanoparticles are) have also been characterized as plastic particles that have at least

one dimension that is between one and one hundred nanometers in length (Besseling et al

2017). Primary microplastics are intentionally manufactured to have micron-sized

dimensions (i.e. industrial pellets, microbeads in pharmaceutical and health care

products). Microplastics or “microbeads” used in health and beauty products are

typically composed of polyethylene and can comprise up to 4.2 and 4% of the total

weight of facial scrubs and toothpaste, respectively (5 Gyres Institute, 2013). Secondary

microplastics have fragmented from a larger plastic product, eventually reaching micron-

sized dimensions. Most microplastics found in the environment are secondary

microplastics, and may pose a greater environmental threat than primary microplastics, as

discussed below.

Approximately 80% of plastic debris is attributed to land-based sources (i.e.

industrial discharges, garbage management, litter), thus it is likely that terrestrial

organisms will also be exposed to microplastics due to the application of microplastic-

3

containing sludge, and the potential release of microplastic fibers from concrete aging

(Andrady 2011; Rillig 2012; Zubris and Richards 2005; Pelisser et al. 2010). Only a few

studies have focused on terrestrial organisms. Sandhoppers (Talitrus saltatory) exposed

to polyethylene spherical microplastics (10-45 µm) for 24 hours were able to excrete all

microplastics within 24 hours after the initial exposure (Ugolini et al. 2013).

Categorizing sandhoppers as strictly a terrestrial organism would be inaccurate as they

also dwell below the wet sand on beach coastlines, making them susceptible to plastic

debris of marine as well as terrestrial origin. The incorporation of surface microplastics

into the burrows of the common earthworm (Lumbricus terrestris) demonstrated that

there may be indirect toxicological effects (i.e. altering microplastic bioavailability to

sediment organisms) associated with microplastic exposures (Lwanga et al. 2017).

Organisms that inhabit such transient and dynamic ecosystems throughout their lifetime

or at different developmental stages will be exposed to microplastics that are specific to

certain environmental conditions, thus making it imperative that additional research be

conducted to best determine how terrestrial and aquatic exposures to microplastics should

be done.

Microplastics have been found in all aquatic ecosystems sampled, but it is

premature to say whether the sediment or surface water will be the larger sink of

microplastics due to the different methods in which microplastics are collected and

quantified (Claessens et al. 2011; Law et al. 2010; Moore et al. 2001; Thompson et al.

2004). Upwards of 300,000 microplastics/m3 have been found in urbanized rivers; it is,

therefore, likely that organisms that either live in, or frequent urbanized coastal areas are

4

at great risk for microplastic exposures (McCormick et al. 2016; Dris et al. 2015; Sanchez

et al. 2014). Coastal food webs may be more susceptible to the trophic transfer than

those in other habitats. This is not to say that remote regions will not also have

microplastic trophic transfer as microplastics and plastic debris in general have been

found in underdeveloped areas. Free et al. (2014) suggest that the general durability and

buoyant nature of plastics contribute to the fast transportation of plastic to areas where

plastic use is low.

The presence of microplastics in the environment has been confirmed in both

aquatic and terrestrial ecosystems, and in even some remote areas as well (Browne et al.

2010; Van Cauwenberghe et al. 2013; Eerkes-Medrano et al. 2015; Free et al. 2014). It is

hypothesized that the ubiquitous presence of microplastics globally reflects not only an

increase in plastic production and coastal human populations, but also the inability to

properly manage the waste stream (Claessens et al. 2011; Claessens et al. 2013; Browne

et al. 2011; Carr et al. 2016; Sutton et al. 2016; Mason et al. 2016; Paul-Pont et al. 2016).

Thus far, most studies have investigated how macro-sized plastic debris impact higher

trophic level marine organisms (i.e. suffocation, blockage of digestive tract, reduction in

food intake, and growth) (Fry et al.1987; Eriksson and Burton 2010; Goldsworthy et al.

1997; Lusher et al. 2017). Conversely, research on the effects of MPs on both aquatic

and terrestrial organisms is in its infancy. The purpose of this review is to critically

examine existing research on the effects of MPs on aquatic organisms.

5

1.2 Plastic disposal and entry into the environment

Plastic disposal methods include burial in landfills, incineration and recycling

(Hopewell et al. 2009); however, each method has different consequences (i.e. residence

times, release of chemicals) that may further alter plastic surface properties and their fate

and transport in the environment (Webb et al. 2013). Floating, non-biodegradable

particulate matter less than 0.5 mm in size will evade waste water treatment capture and

as smaller microplastics are being produced in greater quantities, it is likely that

microplastic presence in the environment will continue despite manufacturer efforts to

decrease microplastic incorporation in various health care products (Fendall and Sewell

2009; Eriksen et al. 2013). Additional research on the removal efficacy of microplastics

as a function of different treatment processes is needed because it has been observed that

over 99% of microdebris can be removed after secondary treatment; however, only 80%

of the removed microdebris is contained in the sludge (Talvitie et al. 2017). The

percentage of the microdebris that is comprised of plastic was not quantified in this study

and for microplastic regulatory purposes, it will be useful to know which wastewater

treatment processes are more effective for microplastic removal. Given the different

wastewater treatment processes available and their use in different facilities, the amount

of microplastics that are captured in the sludge and re-released through their application

in agricultural processes in different regions and countries will vary.

Waste does not always go through waste water treatment facilities, thus MPs may

be directly released into bodies of water (Browne et al. 2011; Leslie et al. 2012).

Approximately 80% of plastic debris is attributed to land-based sources (Andrady 2011),

6

and although the concentration of MPs and plastic debris varies greatly, the fact that

plastics comprise up to 85% of seabed debris in Tokyo Bay (Kanehiro et al. 1995), and

70% of trawled seabed debris in the Mediterranean Sea (Galil et al. 1995) demonstrates

the need to further understand how high releases of synthetic artificial substrates will

affect different organisms. MPs specifically comprise more than 50% (number of

particles) of the field-retrieved plastic debris, and have been found in field-collected fish

and invertebrates from all over the world, thereby demonstrating the environmental

relevance of investigating microplastic interactions and their acute and chronic

toxicological effects (Rummel et al. 2016; Neves et al. 2015; Lusher et al. 2017;

Desforges et al. 2015; Martins and Sobral 2011; Browne et al. 2010).

1.3 Plastic degradation and biofouling

1.3.1 Polymer degradation

The rate at which polymers degrade in the environment has not been extensively

studied because it is difficult to monitor and attribute degrees of degradation to different

environmental factors and processes (e.g. photooxidation, biodegradation, mechanical

degradation). Plastics that are derived from natural products such as starch, chitin, and

cellulose degrade more readily than petroleum-based polymers because they contain

oxygen and nitrogen functional groups; however, less than 1% of all plastics in 2002

were made from natural materials (Scott 2002). Unfortunately, due to the

biodegradability of natural polymers, products normally incorporate petroleum-based

plastics to increase their strength and durability, although the natural polymers may

7

degrade (i.e. CO2, nitrogen, H2O), the petroleum-based polymer residues may not

(Tokiwa et al. 2009). The production of natural polymers such as poly (lactic acid),

which has properties similar to polyethylene, has been incorporated into many petroleum-

based polymer products, yet despite being able to biodegrade, the conditions at waste

management facilities may not facilitate such processes (Tokiwa et al. 2009). Petroleum-

based polymer degradation needs to be further researched because recent data have

shown that bacteria found in the gut of waxworms and mealworms are capable of

degrading ingested polyethylene films, and polystyrene blocks, respectively (Yang et al.

2104; Yang et al. 2015). Due to the incorporation of various stabilizing plasticizers and

the durable nature of plastic in general, some plastics may be unable to undergo complete

mineralization (conversion of organic compounds to carbon dioxide, water, and other

inorganic chemicals) (Andrady 1994). Degradation rates are further influenced by

temperature, oxygen availability, and the presence of microorganisms (Tokiwa et al.

2009).

Degradation rates of plastic in aquatic systems are slower than that of plastic

debris on land due to lower temperatures and less available oxygen to induce oxidative

stress (Rios et al., 2007; Andrady 2011; Webb et al., 2013). Frequent occurrences of

particle-particle collisions as microplastics approach land (saltation and traction) also

contribute to greater physical degradation occurring in coastal ecosystems (Cooper and

Corcoran 2010). Natural degradation often begins with photodegradation and the

incorporation of oxygen atoms into the polymer surface, which can then lead to other

degradation pathways (Webb et al. 2013; Singh and Sharma 2008). Most plastics are

8

consistently exposed to ultraviolet and visible light and degradation will vary based on

the wavelength experienced and the polymer in question; UV light specifically causes

many bonds within plastics to break due to the energy in the light being similar to those

contained in bonds between most of the atoms (Singh and Sharma 2008). Compared to

photooxidation, other processes such as hydrolysis, thermal degradation and

thermooxidative degradation also occur in nature but at a much slower rate (Andrady

2011). The combination of various weathering processes results in continuous changes in

polymer properties, due to chemical, physical and biological reactions; the formation of

new functional groups (hydroxyl, carbonyl groups, and carbon-oxygen), bond scission,

and other alterations can also be visually identified through the deterioration of the plastic

and confirmed using Fourier Transform Infrared (FTIR) spectroscopy (Pospisil and

Nespurek 1997; Shah et al. 2008).

As previously mentioned, the incorporation of additives to prevent product

degradation is common. A common material added to improve the resistance of the

polymer to UV light degradation is carbon black. Carbon black is cheap, has high UV

absorption efficiency, and can also be used to improve mechanical strength, and as a

colorant (Yousif and Haddad 2013). However, the use of carbon black is often paired

with non-carbon-black additives: ultraviolet light absorbers (UVA), and hindered amine

light stabilizers (HALS). Mixtures of these additives are effective in preventing damage

from UV light, and the degree of degradation due to UV light will depend on the

concentrations of the mixtures of additives. Antioxidants can also be added to increase

thermal stability within polymers by removing the propagated free radicals formed by

9

oxidation products, and inhibiting the formation of free radicals (Pornsunthorntawee et al.

2014; Raheem 2012). Another additive that can be used to prevent polymer degradation

are antimicrobials, which are used in products that are primarily involved with food and

medical applications. Silver is known to have antibacterial properties and has been seen

to decrease bacterial growth on various polymers (Marchetti et al., 2015). Additional

additives such as pigments, and stabilizers (i.e. flame retardants, clarifiers,

compatibilizers) also need to be taken into consideration when assessing the formation of

microplastics and toxicological effects of both microplastics and associated contaminants.

The physical weathering of plastics is one of the more obvious ways by which

macroplastics fragment into smaller micron- and nano-sized plastics. However, it is

difficult to identify the type of product a plastic may have been part of originally due to

some weathering processes changing the color, shape and surface of the plastic, thereby

making it difficult to determine whether certain types of products produce more

microplastics in the environment.

1.3.2 Biofouling of plastics

Once released into the environment, plastic debris is all likely to accrue various

microorganisms colonizing the surface, creating biofilms on which algae and invertebrate

populations can be sustained (Andrady 2011; Artham et al. 2009). The formation of

marine biofilms follows four phases: 1. adsorption of dissolved organic molecules, 2.

attachment of bacterial cells, 3. attachment of unicellular eukaryotes, and 4. attachment of

larvae and spores (Dobretsov 2010). The matrixes that are produced by bacterial cells

10

and other attached organisms allow for the attachment of other invertebrates like

barnacles (Fazey and Ryan 2016). The composition of the biofilm and other biofouling

organisms will depend on seasonal changes (i.e. nutrient availability, light intensity and

temperature), thereby also influencing the degradation pathways of plastic debris (Eich et

al. 2015). Despite the development of biofouling organisms on the surfaces of plastic

debris, biodegradation may not necessarily occur as certain organisms that have shown

the capacity for breaking down certain polymers may not be present (Lobelle and Cunliff

2011).

The intensity of UV light that plastic debris is exposed to, which can then in turn

initiate the oxidative process by producing hydroperoxides, is a function of biofilm

growth (Billingham et al. 2000; O’Brine and Thompson 2010). The development of

biofilm on plastic debris surfaces results in a 90-99% decrease in UV transmittance

within 4-40 weeks in the marine environment (O’Brine and Thompson 2010; Weinstein

et al. 2016), potentially inhibiting degradation pathways that involve exposure to heat and

light, by shielding the plastic surface, and causing the plastic debris to sink below the

water surface, making the physical (i.e. mechanical action by waves at the surface and in

the littoral zone) and biodegradation of plastic debris the dominant plastic degradation

pathways in aquatic ecosystems where biofouling occurs quickly (Corcoran et al. 2009;

Copper and Corcoran 2010).

Plastic debris sampled from the western North Atlantic showed that biomass

attached to the plastic surfaces resulted in the sinking of the debris due to an increased

density (Moret-Ferguson et al. 2010). The growth of bacteria on polyethylene shopping

11

bags resulted in their sinking below the water surface, and remaining buoyant within the

water column after three weeks of being released into the marine environment (Lobelle

and Cunliffe 2011). However, as the overall density of plastic fragments increases to the

point of being greater than that of the surrounding body of water (1.0-1.03g/cm3), they

can sink below the water surface (Railkin 2004; Ye and Andrady 1991).

The rate at which microplastics undergo sedimentation from the water column is

species- and polymer-specific in terms of the organisms that colonize the MPs.

Polystyrene MPs colonized by the marine phytoplankton Chaetoceros neogracile

produced larger and sturdier MP aggregates with attached phytoplankton than those

colonized by Rhodomonas salina (Long et al. 2016). Consequentially, the larger MP

aggregates also sunk faster through the water column than smaller aggregates containing

R. salina (Long et al. 2016). As discussed below, particle surface charge can also

influence the binding affinity of nanoplastics to algal cell walls (Nolte et al. 2017;

Bhattacharya et al. 2010), thereby affecting plastic particle aggregate formation,

microplastic biofouling, and sedimentation into the pelagic and benthic zones. Low-

density polyethylene (LDPE) microplastics will also become negatively buoyant (within

12 weeks of being exposed to sea water) before high-density polyethylene (HDPE)

microplastics due to biofouling, despite both polymers originally being less dense than

sea water (Fazey and Ryan 2016). Thus it is likely that the presence and toxicity of MPs

will continuously change as the plastic itself is exposed to different abiotic and biotic

processes (Ye and Andrady, 1991; Barnes et al. 2009).

12

MP properties such as size, shape, and density will influence the sedimentation of

MPs in aquatic ecosystems (Fazey and Ryan 2016; Kowalski et al. 2016), thus also

influencing the type of MPs to which pelagic and benthic organisms will most likely be

exposed. Biofouling was observed to occur faster on polyethylene and polypropylene

plastic debris in the pelagic zone, as compared to those in the benthic zone (Zich et al.

2015). In regards to microplastic shape, those with more angular shapes have greater

drag, thus reducing the initial settling velocity (Khatmullina and Isachenko 2016).

However, microplastics that are more rectangular and have more ridges (i.e. fragments,

fibers) will have greater biofouling potential than spheres (Fazey and Ryan 2016),

potentially increasing particle density and sedimentation. Microplastics that have been

exposed to degradation pathways that lead to greater amounts of cracking, and alterations

of surface structure may provide both additional surface area and the presence of oxygen-

containing functional groups that favor the biofouling process and sorption of certain

contaminants (Schorer and Eisele 1997).

The rate of biofouling within the water column is a function of light and algal

growth; the colonization of plastic with macroinvertebrates may further depend on the

amount and presence of plastic-inhabiting organisms (Ye and Andrady, 1991).

Furthermore, water density and temperature is a function of depth, thus the sedimentation

of MPs and biofouling will likely decrease as water depth increases (Wang et al. 2016).

Yet, the greater pressure, and mechanical shearing experienced at greater depths may also

result in the formation of microplastics in the benthic zone (Murata et al. 2004; Eich et al.

2015). It has yet to be determined what degradation pathways will be dominant in

13

different ecosystems, and further work is necessary to understand location-specific

microplastic formation.

Although de-fouling may occur once submerged below the surface (i.e.

consumption of fouling agents on the surface or the whole plastic) (Andrady, 2011), once

the plastic reaches the benthos, it is possible that biofouling will continue resulting in a

different biofilm composition, and the exposure of microplastics to a different variety of

organisms (Eich et al. 2015).

1.4 Factors that influence microplastic exposure, ingestion, and trophic transfer

1.4.1 Microplastic size, shape, and concentrations

The physical weathering of plastics is one of the more obvious ways in which

plastic debris will break down into smaller micron- and nano-sized plastics. The

exposure to multiple types of degradation pathways makes it difficult to identify from

what type of product a microplastic particle may have originated. When exposed to 30°C

in situ (with the presence of visible and ultraviolet light) for 14 and 56 days, a 1 cm

square of polystyrene foam produced concentrations of 0.32 x 108 and 1.26 x 108

particles/mL, respectively (Lambert and Wagner 2016). Recently it has also been shown

that microplastic fragments and fibers can be produced from high density polyethylene,

polypropylene, and polystyrene strips after only 8 weeks of being exposed in an intertidal

salt marsh (Weinstein et al. 2016). Evaluations of sediment samples have shown similar

concentrations of polyester and acrylic fibers as those found in sewage effluent, most

likely because more than 1,900 fibers are potentially produced by a single garment during

14

one wash (Browne et al. 2011). There may be a way to limit microplastic fibers from

entering the wastewater stream if certain filtration methods are implemented at the

household level.

As plastic products degrade, the number of micro- and nanoplastics produced will

continue to increase, resulting in an increasing presence of plastic with heterogeneous

forms and sizes in the environment. Although the mineralization of plastic products may

take upwards of 50 years, microplastics have been shown to not only form within a few

weeks of being released into an aquatic system, but that the size and shape of particles

formed can also differ based on polymer type (Lambert and Wagner 2016; Weinstein et

al. 2016; Müller et al. 2001). Fish taken from the wild had a greater amount of

microplastics in examined gastrointestinal tissue than macroplastics (> 0.5mm) (Rummel

et al. 2016). Such findings suggest the increasing presence of smaller microplastics may

pose a greater threat than previously thought, especially when considering the higher

surface area to volume ratio that smaller particles have, and their contribution of a higher

amount of adsorbed contaminants to exposed organisms (Norén 2016; Ogonowski et al.

2016).

Size and shape are the most investigated microplastic physical characteristics

amongst laboratory toxicity studies, and are thought to be influential factors when

investigating microplastic ingestion and egestion rates for exposed organisms (Watts et

al. 2015). The shape- and size-specific ingestion of microplastics makes it likely that

degradation rates of specific polymers will be influential in determining the types of

microplastics organisms will more likely encounter and mistake for food (Cole et al.

15

2013; Moore 2008; Hall et al. 2015). Many invertebrates can select ingested particles

based on size, shape and nutritional value. Mussels and lugworms collected from the

field had microplastics between 10-30 µm, and 30-90 µm, respectively, suggesting the

organisms have a size preference (although the exact cause had not been determined)

(Van Cauwenberghe et al. 2015). Polychaetes preferentially consume smaller particles

that stick to their mucus-lined proboscis papillae (Zebe and Schiedek, 1996), and bivalves

can inhibit particles that hold little nutritional value from entering the digestive tract by

forming pseudofeces (Xu et al. 2016).

The greater presence of 3.0 µm microplastics in comparison to 9.6 µm

microplastics in the hemolymph of exposed mussels demonstrates how size may

influence body residence time of microplastic, in addition to ingestion capacity (Browne

et al. 2008). Furthermore, when Daphnia magna were exposed to either 1.0 µm or 20 nm

carboxylated polystyrene particles, it took exposed organisms longer to egest 20 nm

beads, despite preferentially consuming the larger microplastics (Rosenkranz et al. 2009).

Microplastic size can also determine the extent to which microplastics can interact with

algae, which is consumed by almost all aquatic organisms either accidentally or

intentionally; 1.0 µm polyvinyl chloride (PVC) and 0.05 µm polystyrene (PS)

microplastics decreased algal growth, whereas 1 mm PVC and 6 µm PS microplastics did

not (Zhang et al. 2017; Sjollema et al. 2016). By increasing the microplastic to algae

ratio in aquatic environments, organism exposure to microplastics will increase, thereby

not only increasing ingestion rates, but also their susceptibility to suffering from chronic

sublethal effects (i.e. reduction in growth, reproduction, immune function). Yet, there

16

may be a threshold at which exposure to microplastics is less toxic to certain organisms

when food is readily available in high concentrations; microplastic toxicity observed in

exposed Daphnia magna was lessened when provided with high concentrations of algae

(Ogonowski et al. 2016). Despite limited in situ exposures to microplastics of different

sizes, shapes and other physical characteristics, most of the studies have demonstrated

that there is a size-dependent uptake of microplastics. Copepods (Tigriopus japonicas)

easily consumed PS microplastics between 0.5 and 6.0 µm, yet only the smaller PS

microplastics resulted in significant mortality; the exact mechanism for the greater

mortality was not determined, yet it is possible that copepods were better equipped to

egest the larger microplastics since the copepods normally ingest Tetraselmis suecica (~

7 µm) (Lee et al. 2013). This study demonstrated that nauplii (first larval stage of many

species of crustaceans) were able to ingest a few 6 µm microplastics, but it is likely that if

mouthpart and filtration size are limiting factors in determining microplastic uptake, then

the developmental state will influence what age groups of organisms will be most

susceptible to certain types of microplastics.

In the field, the most common reported shapes of microplastics are fragments,

fibers, and other non-spherical shapes (Sutton et al. 2016; Rummel et al. 2016). The

ingestion of fibers results in gut blockages and increased residence times in exposed

invertebrates, whereas smaller microplastic spheres have also been shown to have

increased body residence times at high exposure concentrations of 1000 particles/L

(Murray and Cowie 2011; Au et al. 2015; Xu et al. 2016; Rosenkranz et al. 2009). Yet

the body retention time of either microplastic beads or fibers obtained from face wash or

17

washed clothing, respectively, in exposed goldfish were not significantly different

(Grigorakis et al. 2017). Characterizing the physical microplastic properties that increase

the likelihood of both ingestion rate and residence times will increase our understanding

of microplastic trophic transfer. Since most of the laboratory studies have investigated

homogeneously shaped and sized particles that are <100 um, there is a need for research

investigating the ingestion of smaller particles with a variety of environmentally-relevant

shapes and sizes.

Table 1.1 depicts the varying MP concentration ranges found in different field

studies. Recent studies have demonstrated that freshwater concentrations of MPs are

similar to those found in marine ecosystems; when average MP concentrations found in

freshwater ecosystems are normalized to the average surface area (MPs/km2), total MP

particles range between 1.1 million and 2.7 billion MP particles (Faure et al. 2012;

Eriksen et al. 2013). However, note that in some studies, MP concentrations as low as 1-

6 MPs/m2 have been quantified (Collignon et al. 2012; Martins and Sobral 2011). MP

collection and preservations method differences (i.e. size of meshes, tow speeds) may

potentially influence the MP concentrations found in the environment, making it difficult

to compare MP concentrations across different ecosystems (Collignon et al. 2012;

Martins and Sobral 2011).

18

Table 1.1: Environmentally relevant concentrations of microplastics in various field studies

Source Type of aquatic ecosystem

Type of sampling method

Microplastic shapes collected

Average microplastic (MP) concentrations (MPs/km2)

Reference

Open Ocean, Northwestern Atlantic

Marine 0.947mm mesh net; pelagic

Spherical, films, fragments

67,000 Colton et al. 1974

Lake Hovsgol, Mongolia

Freshwater 333µm mesh net; pelagic

Fragments, films, fibers

20,264 Free et al. 2014

Lake Huron Freshwater 333µm mesh net; pelagic

Pellets, fragments

2,779 Eriksen et al. 2013

Lake Superior


Pellets, fragments


Lake Erie Freshwater 333µm mesh net; pelagic

Pellets, fragments


Lake Geneva, Switzerland


Fragments, films

51,556 Faure et al. 2012

The color of microplastics may also determine which plastics organisms will

preferentially ingest. Sea birds have been shown to selectively ingest plastics that are a

shade of brown and other colors as opposed to those that are clear or dark (Robards et al.

1995; Ryan 1988 Day et al. 1985; Vlietstra and Parga 2002). It is believed that particles

that are colors resembling naturally found items are more likely to be eaten.

Invertebrates have been observed to distinguish food items by color, making it likely that

certain microplastics will be preferentially ingested (mistaking them for food), and

undergo trophic transfer (Brembs 2014). Microplastics found in the Goiana Estuary

19

(Brazil) have the same density as that of fish eggs, suggesting that predators may

accidentally ingest microplastics (Lima et al. 2014); microplastic density, which is be

affected by various degradation pathways and biofouling, will influence what organisms

will be exposed to microplastics.

1.4.2 Microplastic polymer composition

Polymer crystallinity is an indication of how monomers are arranged, and

influences the hardness, rigidity, tensile strength, and opacity of a product. Polyethylene,

polypropylene, polystyrene, polyamide, polyvinyl alcohol, and polyvinyl chloride are

common microplastic polymers found in aquatic ecosystems, yet specific

characterizations other than the general type of polymer are rarely made (i.e. surface area,

surface functional groups, density) (Thompson et al. 2004). Despite both being

polyethylene, high-density polyethylene (HDPE) is more crystalline than low-density

polyethylene (LDPE) because HDPE has longer unbranched hydrocarbon chains, and is

thus stronger and degrades at a different rate than LDPE (Ojeda et al. 2011). Different

polymers are likely to degrade and produce different amounts of microplastics in the

environment, thus polymer crystallinity will ultimately influence the profile and

distribution of microplastics in the environment (Weinstein et al. 2016).

Further, the microstructure of a plastic product will also influence the sorption of

other environmental contaminants to microplastics. The degree of crystallinity will

dictate the capacity for a polymer to store or leach contaminants, as amorphous regions

have more free volume and are thus more permeable than crystalline regions (Gowariker

20

et al. 1986). The size of the penetrant (i.e. contaminant, additive) will also influence its

ability to permeate into or out of the polymer (Sperling 2006). Plasticizers are often

added to alter the flexibility and stability of polymers to design a product with a specific

use. The migration of plasticizers to the surface of a product results in a loss of initial

properties (Vieira et al. 2011). Fries et al. (2013) reported that microplastics found in

sediment samples (polyethylene, polypropylene, polystyrene, polyamide, etc.) contained

an assortment of phthalates as well as inorganic additives such as titanium dioxide

nanoparticles. The incorporation of materials such as carbon black (to improve resistance

of polymers to UV light degradation), antioxidants (to increase thermal stability within

polymers by removing free radicals that might form), and antimicrobials (to reduce

bacterial growth) generally increases the integrity of the plastic by slowing chemical and

biological degradation processes (Pornsunthorntawee et al. 2014; Raheem 2012). As

additives leach out or degrade, microplastic formation is likely to occur at a faster rate.

Such additives and how they are incorporated into or onto a polymer must be taken into

consideration when investigating microplastic formation, fate and transport, organism

susceptibility, and microplastic trophic transfer.

Surface characteristics of plastic particles will also influence their uptake.

Nanoplastics that were neutrally or positively charged had a higher binding affinity to

algal cell walls than negatively charged plastic particles (Nolte et al. 2017). Functional

groups either added to plastic particles during or after being manufactured will not only

influence interactions with commonly ingested algae for filter feeders such as Daphnia,

but may also interact and form aggregates with other particles in the environment such as

21

clay, which are similar to algal cell walls in being negatively charged. There has been no

investigation of what type of microplastic aggregates will form in the environment,

despite this phenomenon occurring in laboratory studies. Artificially produced secondary

polyethylene microplastics aggregated to a greater extent than primary microplastics,

resulting in a significant decrease in the amount of secondary polyethylene microplastics

ingested by Daphnia magna (Cole and Galloway 2015; Ogonowski et al. 2016). This

may be due to the inability of Daphnia to ingest larger aggregates. The aggregation of

microplastic with other particles may open up new pathways in which organisms that

prefer to consume larger objects will ingest significant quantities of smaller

microplastics. Microplastics can easily adhere to the surface of suspended seaweed,

Fucus vesiculosus, resulting in the consumption of microplastics by gastropods,

suggesting a potential inability of organisms to distinguish between food and

microplastics (Gutow et al. 2015). Further, microplastics have been shown to easily

adhere onto the external surfaces of exposed organisms, potentially restricting their

movement and providing another route of indirect ingestion of microplastics (i.e. trophic

transfer) (Rehse et al. 2016; Watts et al. 2014). Continuous changes of microplastic

characteristics (i.e. surface functional groups, cracking, surface area changes) make it

difficult to study microplastic presence in the environment, and the exposure and

consequential toxicity that may incur. Depending on where plastic debris or

microplastics are released, and where they are transported, the resulting alterations made

in surface characteristics will likely be influenced by the regional climate patterns; thus

22

microplastic environmental concentrations, and exposures will be unique to different

areas.

Field studies have shown that secondary microplastics are predominantly found

in the environment, whereas laboratory studies have focused on characterizing the

toxicity of primary microplastics (Au et al. 2015; Cole et al. 2015; Cole et al. 2013, Lee

et al. 2013; Kaposi et al. 2013; Setälä et al. 2014). Studies in which secondary

microplastics are made in the laboratory are also likely to have different physical

(cracking, fragmentation) and chemical (surface functional groups, additives)

compositions than those that occur in the environment (Ogonowski et al. 2016; Au et al.

2015). Although it is reasonable for researchers to use easily characterized and

homogenously shaped and sized (usually fluorescent) primary microplastics to establish a

threshold of effect of microplastic exposures in aquatic organisms, it is difficult to gauge

which particle characteristics actually contributed to the different toxicological effects

observed. Unfortunately, the analysis of the surface properties of microplastics found in

the field is hindered by variations in collection methods, equipment used to analyze the

plastics, and cleaning and storage methods, all of which potentially alter the surface

characteristics (i.e. functional groups or biofilms present), or damage the particles

(making it difficult to simulate environmentally-relevant microplastic exposures in situ).

Researchers must also be cautious in the interpretation of their results as it has been

recently pointed out that the misidentification of natural particles or polymers as synthetic

polymers using visual identification will result in the overestimation of microplastic

ingestion in the wild (Wesch et al. 2015).

23

1.4.3 Regional climate patterns

Regional weather patterns greatly influence the presence and transport of

microplastics in aquatic ecosystems. Large precipitation events, and fluxes within the

water column can result in extreme changes in microplastic concentrations in both the

water column and the benthos. Microplastics found in the sediments of coastal areas

were approximately 3x greater after a flooding event in India; however, it is unknown if

most of the microplastics washed onto the beach from the freshwater tributaries or the sea

via wind and surface currants (Veerasingam et al. 2016). Increased river flow, and

turbulence during storm events result in greater microplastic transport from freshwater

sources, and re-suspension of sediment that may contain high concentrations of

microplastics (Lima et al. 2014; Lima et al. 2015; Cole et al. 2011). The 2011 tsunami in

Japan generated an estimated 1.5 million tons of floating marine debris, which will likely

reach North American shorelines for many years to come (Maximenko et al. 2015).

Yet, in the Southern Sea of Korea, rainy seasons resulted in a decreased

microplastic to zooplankton ratio (0.086 to 0.022 in 2012, and 0.016 to 0.004 in 2013)

(Kang et al. 2015). An approximate 5% decline in microplastic concentrations in water

samples following a large precipitation event suggests that such events may reduce

organism encounters with microplastics (Kang et al. 2015). Higher ratios of microplastics

to zooplankton are also found in sampling sites closer to municipal wastewater treatment

plants, and urbanized areas (Kang et al. 2015), demonstrating the significant role that

terrestrial sources may play in microplastic exposures in aquatic ecosystems. The

24

bioavailability of microplastics before and after large precipitation events may cause

large microplastic fluxes between the pelagic and benthic zone. Wave action, tides, and

other temporal and spatial processes will also influence microplastic transport and fate, as

well as microplastic formation from plastic debris (Moreira et al. 2015). In addition,

diurnal changes in the tide may also influence the bioavailability of both microplastics

and adsorbed contaminants (Boucher et al. 2015).

1.4.4 Uptake and depuration

The uptake and depuration of microplastics from organisms are a function of the

physical and chemical properties of the microplastics, anatomy and physiology of the

exposed organism, and a wide variety of environmental variables. Microplastic uptake

will likely be higher in areas with greater abundance of microplastics or plastic debris,

and the rate at which microplastics are ingested will in turn depend on organism

preferences and ingestion capacities for certain particle sizes, shapes, and surface

characteristics. Exposure to microplastics has resulted in either increasing or decreasing

gastrointestinal depuration rates in some studies (Sussarellu et al. 2016; Xu et al. 2016;

Wegner et al. 2012; Ogonowski et al. 2016), whereas in other exposures, there was no

difference observed (Browne et al. 2008). Polymer type may account for some of the

different observations in regards to microplastic depuration rates; Browne et al. (2008)

exposed organisms to polyvinyl chloride microplastics, whereas significant effects on

depuration rates occurred when organisms were exposed to polystyrene or polyethylene

microplastics. Unfortunately, it is difficult to concretely state whether specific polymer

will primarily influence microplastic egestion because the three polymers used most

25

likely had different types and amounts of additives and different organisms were exposed

to microplastics for different amounts of time.

Microplastic clearance may be size-dependent; as mentioned above, Daphnia

excreted 90 and 40% of the ingested 1.0 µm and 20 nm polystyrene microplastics,

respectively, within four hours (Rosenkranz et al. 2009). Microplastic clearance may

also be a dose-dependent, where clams excreted 85 and 50% of the ingested microplastics

when exposed to 1000 and 10 microplastics/L, respectively, within the same depuration

period (Xu et al. 2016). Interestingly, when polyethylene microplastic concentrations

were four times greater than that of algae, Daphnia magna took twice as long to egest

microplastics (Ogonowski et al. 2016). Although not a dose-dependent observation, these

results demonstrate that the presence of food will also influence microplastic depuration

rates. Microplastic shape will also determine their residence time within the

gastrointestinal tract, where secondary fragmented and fibrous microplastics took longer

to be egested than primary microplastics (Ogonowski et al. 2016; Au et al. 2015).

Microplastic spheres are found in smaller quantities in the environment compared to

fibers and fragments. In general, increased exposure to and reduced elimination of

ingested microplastics will result in a longer body residence time of microplastics.

Organism anatomy and physiology will influence microplastic retention times as

younger and smaller organisms will be limited by gape (size of mouth parts). As smaller

microplastics are found in higher concentrations than larger microplastics, it is likely that

smaller microplastics will not only be ingested at higher concentrations and by more

organisms, but will also have a longer residence time if able to translocate from the

26

gastrointestinal tract, and into other organs (Browne et al. 2008; Brennecke et al. 2015).

Gut morphology can play a large role in microplastic retention, as microplastic fibers

have been shown to form ball-like aggregates and block the gut, preventing microplastic

egestion (Murray and Cowie 2011; Watts et al. 2015). Lobsters (Nephrops norvegicus)

sampled from the wild showed that larger individuals, males, and those that had recently

molted contained less microplastics in the gut, indicating that ecdysis (i.e. cuticle removal

process) is one of the main excretion routes for N. norvegicus. Younger N. norvegicus

are more susceptible to gastrointestinal blockage by microplastic aggregates, which may

limit their ingestion of necessary nutrients (Welden and Cowie 2016). After being

exposed to 15 µm latex microplastics, estuarine copepods (Eurytemora affinis) were able

to regurgitate ingested microplastics an hour later, despite ingesting up to 59,000 particles

per individual per hour (Powell and Berry 1990). Many organisms do not have the

capacity to undergo ecdysis or regurgitation where toxic or purely non-nutritional

gastrointestinal-containing materials can be directly eliminated. In addition to an increase

in microplastic uptake, the inability or delayed ability of exposed organisms to excrete

ingested microplastics, will result in an overall increased microplastic body residence

time, leading to an increase in the bioaccumulation and biomagnification of microplastics

and associated contaminants.

1.5 Microplastic toxicity to aquatic organisms

Microplastic toxicity and environmental presence have been studied to a greater

extent in aquatic ecosystems, in comparison to terrestrial ecosystems. Although research

27

regarding plastic pollution has focused primarily on marine ecosystems, recent studies

have suggested that MP distribution and toxicity may be just as prevalent in freshwater

systems (Eerkes-Medrano et al. 2015). Plastic pollution is a result of both terrestrial and

aquatic sources, with a majority of plastic debris eventually migrating to waterways,

influencing various biological functions and processes (Webb et al. 2013). The

entanglement and consumption of plastics have been extensively monitored and

documented in higher trophic level organisms such as marine mammals (Laist 1997) and

birds (Rios et al. 2007), respectively. In regards to invertebrates, microplastic ingestion

has been evaluated in various species, and negatively affects cellular to population level

biological processes and functions.

Few uptake routes of microplastics other than their direct consumption have been

investigated in situ, thus more research is needed to concretely determine whether the

consumption of microplastics is the most toxic exposure route for aquatic organisms.

The ingestion of microplastics has been observed to have polar effects on organism

uptake of nutrients and food materials. Many organisms serendipitously ingest non-food

with food materials, consequentially ingesting small organisms and other nutrients that

adhere to the particles (Phillips et al. 1980). Specifically, aquatic invertebrates often

intentionally consume non-food or nutrient-poor materials to feed upon the microbial

biofilm present on the surface and excrete the non-food or nutrient-poor materials fairly

easily (Cummins and Klug 1979). Plastic consumption has been seen in many

invertebrates including amphipods, lugworms, barnacles (Thompson et al. 2004; Au et al.

2015), mussels (Browne et al. 2008), crustaceans (Murray and Cowie, 2011), and sea

28

cucumbers (Graham and Thompson 2009), representing organisms that live throughout

the pelagic and benthic zones, with multiple feeding mechanisms.

1.5.1 Energy reserves and growth

In the presence of food, polystyrene micro- and nanoplastic exposures negatively

affected algae uptake by exposed clams (Atactodea striata) and blue mussels (Mytilus

edulis) respectively, by reducing filtering activity, and increasing the production of

pseudofeces (Xu et al. 2016; Wegner et al. 2012). The exposures to polystyrene micro-

and nanoplastic particles were between 8 hours and 10 days, thus it is not known whether

chronic exposures to such particles will result in starvation should organisms experience

both reduced food uptake and depleted energy from increased microplastic excretion (Xu

et al. 2016; Wegner et al. 2012). Similarly, copepods (Calanus helgolandicus) exposed

to polystyrene microplastics had reduced algae consumption, as well as a depletion in

energy over time (Cole et al. 2015).

In the presence of food (Artemia spp.), polyethylene microplastic exposures

resulted in a reduction in predatory performance and efficiency in juvenile common goby

(Pomatoschistus microps) collected from Lima River (Iberian coast), thereby reducing

food intake (de Sá et al. 2015). Interestingly, the consumption efficiency of Artemia was

not significantly affected in juvenile gobys collected from Minho River, a nearby estuary,

suggesting that there may also be other factors influencing organism susceptibility that

are water body-specific (de Sá et al. 2015). Exposure to polystyrene and polyethylene

microplastics resulted in a reduction in body size in exposed D. magna, and Hyalella

29

azteca, respectively (Besseling et al. 2014; Au et al. 2015). It has yet to be determined

whether there is a threshold at which given the amount of food and nutrients available,

microplastic ingestion will not influence nutrient uptake.

Exposures with higher concentrations of algae (9 µg C/mL vs. 0.4 µg C/mL)

resulted in growth regardless of the exposure concentrations of polyethylene

microplastics (1 x 102 – 105 microplastics/mL) (Ogonowski et al. 2016). On the other

hand, when exposed to microplastics concurrently with a lower concentration of algae,

growth was lower in all treatments of microplastics and kaolin, suggesting that the uptake

of any non-nutrient particle can reduce growth if the diet is not supplemented with

additional food (Ogonowski et al. 2016). The adsorption of microplastics to algae

surfaces, and the formation of aggregates result in a decrease in photosynthesis

(chlorophyll content and photosynthetic efficiency) and overall growth in treatments with

exposed marine microalgae (Skeletonema costatum) (Zhang et al. 2017). Polystyrene

microplastics can also decrease algal growth and photosynthesis (Sjollema et al. 2016;

Besseling et al. 2014). The presence of microplastics on the surface of algal cells may

block light, CO2, O2, and light transfer (Zhang et al. 2017).

A two-month exposure to 2 and 6 µm polystyrene microplastics resulted in an

increase in microalgae consumption in exposed Pacific oysters (Crassostrea gigas),

suggesting that oysters may be compensating for the ingestion of non-nutritional items

such as microplastics by increasing overall ingestion rates to attain enough microalgae

(Sussarellu et al. 2016). These data suggest that for some organisms, if there is enough

uptake of nutrients ingested through increased food availability or mechanical ingestion

30

capabilities, then microplastic ingestion may not inhibit growth. Exposure to polystyrene

microplastics did not cause an overall change in the calculated cellular energy allocation

of exposed M. edulis and Arenicola marina, despite increasing the energy consumption of

exposed M. edulis by 25%, and protein content in A. marina by 18% (Van Cauwenberghe

et al. 2015). Increasing energy consumption (respiration) in response to microplastic

ingestion has been linked to stress and may lead to the reallocation of energy from

reproduction to survival and growth as discussed below (Sussarellu et al. 2016;

Ogonowski et al. 2016; Smolders et al. 2002).

Microplastic shape may influence microplastic retention once ingested, and

organism development. Freshwater amphipods exposed to polyamide microplastic fibers

(500 x 20 µm) for 14 days showed a decreased assimilation efficiency for leaf materials,

despite the fibers being egested within 16 hours of exposure (Blarer and Burkhardt-Holm

2016). Juvenile H. azteca exposed to polypropylene microplastic fibers for 10 days also

showed reduced growth (Au et al. 2015). Yet, ingestion of both primary and secondary

polyethylene microplastics resulted in a decrease in algae uptake by D. magna

(Ogonowski et al. 2016). It is unclear as to whether microplastic ingestion can reduce

food intake or interfere with food processing; however, ingested microplastics may

physically damage structures within the gastrointestinal tissue that results in the

impairment of assimilation efficiency (Blarer and Burkhardt-Holm 2016).

Inflammatory responses due to microplastic particles were detected in A. marina

(Wright et al. 2013) and in the mussel, M. edulis (Von Moos et al. 2012). Wright et al.

(2013) suggested inflammation combined with reduced feeding activity and longer gut

31

residence times was responsible for depleted energy reserves (i.e. reduced lipid reserves)

of the lugworm.

1.5.2 Reproduction

Thus far research has demonstrated that microplastic exposures can have

multigenerational effects. Exposure to polyethylene and polystyrene microplastics

significantly reduced the number of D. magna, H. azteca , and C. helgolandicus neonates

produced (Ogonowski et al. 2016; Au et al. 2016; Cole et al. 2015). D. magna exposed to

polystyrene nanoplastic particles produced fewer neonates with smaller body sizes and

more malformations (Besseling et al. 2014). Oysters exposed to polystyrene microplastics

for two months resulted in a decrease in fecundity and offspring development (Sussarellu

et al. 2016). Specifically, oocyte number, size, and sperm velocity decreased by 38, 5,

and 23%, respectively, resulting in decreases in larval oyster development by 18%

(Sussarellu et al. 2016). Exposure to polystyrene microplastics (0.005-0.5µm) increased

the amount of time it took for nauplii to develop into copepodites and adults, and

decreased the number of nauplii produced in two generations of Tigriopus japonicus (Lee

et al. 2013). The ingestion of non-nutritional items most likely results in the reallocation

of energy from reproduction to consumption capacity and survival (Sussarellu et al. 2016;

Au et al. 2015). Among potential beneficial uses, microplastics and plastic debris in

general can serve as a new durable surface on which organisms can use for egg

deposition, potentially increasing the number of offspring produced and the distance at

32

which offspring will travel because plastic will not degrade as quickly as surfaces such as

wood or kelp, and they may travel to areas with less natural predators.

Paint chips contain high concentrations of polymeric (polyethylene, polyethylene

terephthalate, expanded polystyrol, and polyurethane) compounds and additives, and

exposure to snails, Assiminea grayana, (particles were not characterized by size) resulted

in changes to snail reproductive structure development (i.e. necrotic oocytes, atypical

spermatozoa) (Watermann et al. 2017). However, the changes in reproductive organ

growth were not necessarily attributed to microplastic toxicity; the presence of other

contaminants on the microplastics (Diuron, tributyltin, and various metals) that are

endocrine disruptors may have contributed to the changes observed (Watermann et al.

2017). Similarly, aged polyethylene microplastic exposures also resulted in greater

developmental toxicity in exposed Japanese medaka due to the leaching of endocrine

disrupting contaminants (Rochman et al. 2014a). Specifically, exposure to aged

polyethylene microplastics for two months decreased the expression of genes involved in

the production of the estrogen receptor (ERα), vitellogenin (Vtg 1) and choriogenin H

(Chg H), which are transformed into egg yolk and egg envelope proteins, respectively, in

exposed Japanese medaka (O. latipes) (Rochman et al. 2014a). Changes in fertility may

cause life history shifts on various populations and affect entire food webs where certain

organisms depend on the availability of certain prey at specific development stages.

Exposure to aged polyethylene microplastics resulted in significantly higher development

abnormalities and mortality in exposed juvenile brown mussels (Perna perna) compared

to virgin polyethylene microplastics (Gandara e Silva et al. 2016)

33

1.5.3 Bioavailability of additives, adsorbed environmental contaminants, and viruses

Many priority pollutants have been associated with microplastics that can be

introduced during the manufacturing process and sorb from surrounding environmental

media (Rochman et al. 2013. Microplastics are associated with significant amounts of

persistent organic pollutants such as polycyclic aromatic hydrocarbons, polychlorinated

biphenyls, dichlorodiphenyltrichloroethanes, perfluorooctanesulfonate,

perfluorooctanesulfonamide (Fries et al. 2013; Frias et al. 2010; Rios et al. 2007;

Rochman et al. 2013b; Wang et al. 2016), as well as inorganic contaminants such as

silver, titanium dioxide nanoparticles, barium, sulfur, and zinc (Khan et al. 2015; Fries et

al. 2013). The presence of microbeads (taken from cosmetic products) resulted in the

sorption of cadmium from contaminated intertidal sediment, and decreased the amount of

cadmium in pore and surface water (Boucher et al. 2016). In addition to the above

chemicals, styrene (a monomer in many plastic types), bisphenol-A, pthlatates and

nonylphenol (additives in plastic) have also been shown to disrupt endocrine system

function (Iguchi et al. 2006; vom Saal and Hughes 2005; Harris et al. 1997; Fent et al.

2014). It is not yet clear that microplastics always increase the uptake of the associated

contaminants because the concentrations of chemicals found in field organisms do not

always correspond with the amount of microplastics collected (Paul-Pont et al. 2016;

Rochman et al. 2014b). The differences in contaminant bioavailability due to

microplastic exposure may be associated with the biofilm growth on microplastic

surfaces and seasonal changes; higher biofilm growth is associated with greater persistent

organic pollutant concentrations (Schorer and Eisele 1997).

34

Some studies have shown that contaminants adsorbed to microplastics are

bioavailable to exposed organisms, and affect molecular and cellular pathways (Avio et

al. 2015; Khan et al. 2015; Rochman et al. 2014a; Rochman et al. 2014b; Oliveira et al.

2013). Exposure routes other than digestion (i.e. dermal, ventilation) may also contribute

to the bioavailability of microplastics and associated contaminants. Yet, according to

other studies, the high concentrations of contaminants adsorbed to microplastics in situ

kinetically favor the transfer of contaminants from the microplastics to the exposed

organism, suggesting that at ecologically relevant concentrations, microplastics are not a

significant vector of contaminants, at least when using thermodynamic models

(Koelmans et al. 2016; Ziccardi et al. 2016). Exposures of polyvinyl chloride

microplastics in the presence of phenanthrene or 17α- ethinylestradiol (EE2) to larval

zebrafish resulted in greater sorption of EE2 on microplastics despite phenanthrene

having a higher log KOW, suggesting that contaminant bioavailability in the presence of

microplastics cannot be predicted using physiochemical properties alone (Sleight et al.

2017). More research is needed to establish the best laboratory and field practices that

are appropriate for determining whether the presence of microplastics will influence the

bioavailability of other environmental contaminants.

Overall, it is likely that smaller plastic debris may be a greater concern in regards

to contaminant bioavailability because the general buoyant nature of most plastic debris,

and small size (greater surface area) enables microplastics to be a vector for high

concentrations of pollutants that can travel great distances. Smaller microplastics take

longer to reach equilibrium with surrounding water due to higher surface areas. Further,

35

as most plastic debris will be released from urban coastal areas, it is likely that these

microplastics will accrue large concentrations of contaminants quickly (due to a greater

input of contaminants in urban coastal areas), prolonging the amount of time it takes to

reach equilibrium (Karapanagioti et al. 2010). By traveling great distances quickly, many

organisms and food webs may be exposed to high concentrations of microplastics with

adsorbed contaminants prior to the contaminants adsorbed to the microplastics reaching

equilibrium with the external media (i.e. sediment, water).

Pomatoschistus microps (common goby) exposed to pyrene-contaminated

polyethylene microplastics (1-5 µm) for 96 hours resulted in lethargic swimming

behavior, reduced acetylcholinesterase (AChE) a and isocitrate dehydrogenase (IDH)

activity, and increased amounts of pyrene metabolites in the bile (Oliveira et al. 2013).

AChE is involved with neuro transmission in fish; IDH is involved with the aerobic

pathway of cellular production, and cellular antioxidant defense (Oliveira et al. 2013).

Changes in swimming behavior, possibly due to the decrease in AChE activity, combined

with a decrease in antioxidant defense may reduce organism fitness in the environment.

Greater concentrations of polybrominated diphenyl ethers (PBDEs), the polychlorinated

biphenyl 28 (PCB28), and the PAH chrysene were found in Japanese medaka (Oryzias

latipes) exposed to field-aged polyethylene microplastics in comparison to virgin

polyethylene microplastics (Rochman et al. 2014b). Depending on how degradation

pathways alter microplastic physical (i.e. shape, size) and chemical (i.e. surface

functional groups, presence of biofouling organisms) properties, aged microplastics may

also act as a source for environmental contaminants.

36

1.5.4 Trophic transfer

Limited research is available regarding the trophic transfer of microplastics;

however, the discovery of microplastics in the scat and tissues of various coastal higher

trophic level organisms that represent a diverse array of habitats (i.e. mammals, birds,

and fish) suggest the strong potential for trophic transfer occurring concurrently with the

direct ingestion of microplastics (Fry et al.1987; Eriksson and Burton 2003; Goldsworthy

et al. 1997; Lusher et al. 2012). Microplastics are in the size range of many species of

plankton and other types of particles that are normally ingested by aquatic invertebrates,

thus posing the potential for microplastics and associated contaminants to undergo

trophic transfer to higher biologically classified organisms (Desforges et al. 2015;

Browne et al. 2008). Invertebrates and fish with different feeding mechanisms and

habitats have also been observed to easily consume microplastics in vivo and in vitro

(Browne et al. 2008; Thompson et al. 2004; Graham and Thompson 2009; Setälä et al.

2014; Jabeen et al. 2017).

The presence of microplastics and zooplankton in both the sediment and water

column makes it inevitable that microplastics will be consumed by both zooplankton and

other potential predators. The trophic transfer of polystyrene microplastics to

macrozooplankton occurred after only three hours of exposure to mesozooplankton that

had previously ingested polystyrene microplastics (Setälä et al. 2014). Although the

amount of microplastics ingested per zooplankton individual may not be high in the

environment (less than one particle per organism examined), the continuous consumption

of high numbers of zooplankton by predators such as humpback whales and juvenile fish

37

make it likely for microplastics to bioaccumulate in larger fish and mammals (Desforges

et al. 2015). The presence of microplastics and plastic additives in whale feeding grounds

also makes it likely that whales will ingest high concentrations of microplastics,

persistent organic pollutants, and contaminated prey (Fossi et al. 2016). The ingestion of

microplastics by organisms throughout the water column in addition to the benthos

indicates the ease at which microplastics may be transferred throughout the food web and

water column (Setälä et al. 2014).

The residence time of microplastics in the tissues and gut of biota is an important

factor in trophic transfer and a function of several factors. Microplastics were found in

the hemolymph of crabs (Carcinus maenas) 21 days after being provided with mussels

(Mytilus edulis) that had consumed polystyrene microplastics (Farrell and Nelson 2013).

The residence time of the microplastics found within the crabs varied among different

organs (i.e. digestive tract, hepatopancreas, ovary, gills), but were less than that of the

hemolyph (Farrell and Nelson 2013). The translocation of polystyrene spherical

microplastics, <10 and 0.5µm, into the hemolymph of invertebrates for up to 48 and 21

days, respectively, suggest that perhaps the longer residence time of microplastics due to

translocation may depend on polymer composition or size (Farrell and Nelson 2013;

Browne et al. 2008). Body residence time may also be influenced by uptake routes,

because it was observed that the ventilation of microplastics through gills resulted in a

longer microplastic residence time, than that from microplastic ingestion, when crabs

were exposed to 8-10 µm polystyrene microplastics (Watts et al. 2014). Microplastic

shape is another characteristic that may influence body residence time, as the ingestion of

38

fiber-shaped microplastics by the crab (C. maenas) and lobster (Nephrops) resulted in the

formation of an microplastic aggregate within the gut and a reduction in microplastic

egestion times (Murray and Cowie 2011; Watts et al. 2015). Longer body residence

times of microplastics will increase the chance that microplastics within a prey can be

consumed by a higher level organism, thus providing another route of exposure to

microplastics in the environment. Whether this contribution is sufficient to be toxic at

environmentally relevant concentrations is uncertain as there is a high variability in

microplastic consumption by exposed organisms (Farrell and Nelson 2013; Au et al.

2015).

Numerous chemicals such as plasticizers and the antibacterial chemical, triclosan,

are often incorporated into plastic products during their production (i.e. additives) and

can also adsorb onto plastics once released into the environment (i.e. persistent organic

pollutants). Concerns have been voiced about the trophic transfer of chemicals

associated with microplastics, and the toxicological ramifications of microplastics acting

as a source and vector for a mixture of contaminants. It is likely that the uptake of

microplastic-associated contaminants will be greater for higher trophic organisms as

desorption rates of persistent organic contaminants from microplastics were faster in

treatments that simulated gut conditions of warm blooded organisms (i.e. higher body

temperature, lower pH, and the presence of gut surfactants) (Bakir et al. 2014). Only one

published study thus far has demonstrated the trophic transfer of polyethylene

microplastics and adsorbed benzo[a]pyrene from Artemia sp. to zebrafish, where the

desorption of benzo[a]pyrene was observed in the intestinal epithelium and resulted in an

39

induction of cytochrome P450 1A (CYP1A) in the liver of the exposed zebrafish (Batel et

al. 2016). Plastic ingestion has also been related to trace metals and PBDEs present in

sea bird feathers and abdominal adipose tissue, respectively (Lavers and Bond 2016;

Tanaka et al. 2013), demonstrating how plastic ingestion can contribute to toxicant

uptake in larger marine organisms. However, when persistent organic pollutant (POP)

uptake from plastic ingestion by fulmars was compared to that from the ingestion of prey,

it was found that the bioaccumulation of POPs due to plastic ingestion is negligible in

comparison to other POP uptake routes (Herzke et al. 2016). The limited evidence to date

indicates that it is likely that microplastics may act as a source of contaminants to

exposed organisms through their direct and indirect (trophic transfer) consumption;

however, the reviewed studies have exposed contaminated microplastics to clean test

organisms (where the transfer of contaminants to the organism is favored) (Koelmans

2015). In environments where contaminants are already present, microplastic ingestion

may act as a sorbent for contaminants already present within the exposed organism,

resulting in an overall depuration effect (Koelmans 2015). Rainbow trout first exposed to

polychlorinated biphenyls (PCB) and then polyethylene microplastics through their feed

for up to nine weeks after the initial PCB exposure demonstrated that there was no

significant effect on PCB elimination rates, indicating that in this experiment, the

ingestion of uncontaminated MPs did not result in an expedited depuration of PCBs from

exposed fish (Rummel et al. 2016). Further investigations on whether microplastics will

act as a source of or depuration method for environmental contaminants are needed.

40

The presence of microplastics in the wild as well as areas designed for

aquaculture indicates the potential for microplastics to be present in organisms commonly

ingested by humans (Davidson and Dudas 2016; De Witte et al. 2014). Microplastics

found in both wild and cultured Manila clams (Venerupis philippinarum) (0.07-5.47

particles/g), in British Columbia indicate that microplastics are in seafood consumed by

humans (Davidson and Dudas 2016). Microplastic fibers between 200 and 1500 µm were

detected in blue mussels (Mytilus edulis) purchased from a store (De Witte et al. 2014).

Microplastics were found in higher concentrations in supermarket-purchased clams

(Crassostrea gigas) in comparison to M. edulis from a mussel farm, suggesting that

microplastics present in human consumption-ready shellfish may be species-specific

(Van Cauwenberghe and Janssen 2014). Although fish are normally gutted, thereby

removing most of the microplastics ingested, prior to human consumption, this is not the

case with shellfish, where the whole organism is generally consumed intact. It is possible

that the trophic transfer of microplastics from the environment to humans may occur

through the ingestion of whole organisms, as well as through ventilation (fibers from

clothing, and other products).

1.5.5 Population-level effects

The presence of microplastics in sediment results in their incorporation into

burrows built by both terrestrial and aquatic invertebrates, thereby altering sediment

habitat structure. Approximately 73% of the LDPE microplastics present on the sediment

surface was introduced into the burrow walls by exposed common earthworms

41

(Lumbricus terrestris) with over 50% of the microplastics having a diameter ≤ 50 µm

(Lwanga et al. 2017). Size selectivity observed by L. terrestris, a terrestrial invertebrate,

demonstrates that organisms deemed as ecosystem engineers may further change their

environment for themselves and other organisms when exposed to microplastics.

Depending on the microplastics present, the permeability and heat transfer within soils

may change (Carson et al. 2011). The construction of heavier burrows with the

incorporation of LDPE as compared to treatments with only sediment and organic matter

suggest that the pore spaces within sediments that include microplastics will be smaller,

potentially affecting the bioavailability of contaminants and future burrowing capabilities

by other organisms (Lwanga et al. 2017). By impacting water movement and soil

temperature within sediment environments, there may be many inadvertent detrimental

effects on other organisms that may not even ingest microplastics.

The presence of plastic in the environment also makes it possible for invasive

fauna to migrate to new environments (Masó et al. 2003; Barnes and Milner 2005).

Historically, organisms have been geographically limited, yet with increasing human

travel, the introduction of invasive species has become more frequent and destructive

(Barnes and Milner, 2005). Plastic debris found in both isolated islands and heavily

populated areas have been inhabited by various organisms (Ryan 1987; Benton 1991).

Organisms such as dinoflagellate algae (Masó et al. 2003), barnacles, polychaetes,

bryozoans, hydroids (Barnes and Milner, 2005), and iguanas (Censky et al. 1998) were

observed to move on rafts of plastic debris in the Atlantic Ocean. The plastic type also

impacts the species abundance of colonizing microorganisms; plastics sampled from the

42

North Atlantic showed a dominance of the genus Vibrio (bacteria associated with various

pathogens) on polypropylene samples (Zettler et al. 2013), despite constituting a small

fraction of worldwide marine bacterial populations (Thompson and Polz 2006),

suggesting that plastics may provide a preferred surface for certain organisms.

Microplastics may also be an oviposition resource for insects, where Halobates sericeus

eggs deposition was a function of microplastic presence in the North Pacific Subtrophical

Gyre and on beaches (Goldstein et al. 2012; Majer et al. 2012).

The introduction of invasive species poses many problems to native wildlife, and

although it is not possible to eliminate all sources of species introduction to exotic places,

the addition of plastic products has increased the ways in which organisms can reach new

locations. The addition of invasive species often results in the loss of native species,

specifically in regions like the Southern Ocean that has high levels of endemism. Like

wood, plastic can host a variety of organisms and in high abundances, whereas other

floating entities like kelp, carry significantly less encrusting colonists (Barnes and Milner

2005). Higher colonization of debris correlates with higher abundances of plastic debris,

with no differences found between the North and South Hemispheres (Barnes and Milner

2005). Plastic debris in marine systems were also seen to recruit bacteria colonies that

are normally on benthic substrates (Zettler et al. 2013); the presence of microorganisms

that are typically absent from the water column although increasing microorganism

diversity, may have other unforeseen ecological effects (i.e. spreading pathogens to other

regions and higher trophic organisms).

43

The durability of plastic debris provides a stable and longer lasting habitat for

organisms than wood or aquatic vegetation that degrade faster. However, there may also

be limitations for the colonization of plastics by certain organisms that cannot withstand

specific temperatures, or varying salinity levels as the debris travels through different

regions. Organisms with faster growth rates and the capacity to develop specific

attributes (formation of holdfast filaments, or vegetative cysts) may be able to disperse

into areas and water depths that they are normally unable to colonize. Polymer

characteristics may also play a role in microorganism colonization, as polypropylene has

been found to have greater species richness than polyethylene debris found in the North

Atlantic Ocean (Zettler et al. 2013).

1.6 Dissertation goal and objectives

Clearly from the preceding literature review, there is much yet to learn about the

role of microplastics in both marine and freshwater ecosystems. The overall goal of this

dissertation was to gain a better understanding of how exposures to microplastics affect

freshwater and marine invertebrates and fish. To achieve the goal, the first objective was

to further understanding of how microplastic ingestion and egestion will toxicologically

affect invertebrates and fish by establishing a threshold of effect for mortality, growth,

and reproduction. The second objective was to investigate whether microplastics could

serve as a vector for the polycyclic aromatic hydrocarbon, fluoranthene, to exposed

invertebrates and fish. I sought to gain a better understanding of how the presence of

microplastics would influence the bioavailability of persistent organic pollutants in

44

aquatic ecosystems. The third objective was to evaluate whether microplastics and the

adsorbed fluoranthene could undergo trophic transfer.

The following chapters explain each objective, and the tasks that were necessary

to accomplish each one. Detailed methodology regarding the bioassays conducted for

each organism and the results from each experiment are also described in each chapter.

A discussion that relates my results to those that are available in published literatures as

well as limitations of each experiment, and general conclusions wrap up each chapter.

Two publications to date resulted from the compilation of the literature review (Au et al.

in press), and results from Chapter Two, regarding microplastic toxicity observed with

the freshwater amphipod, Hyalella azteca (Au et al. 2015). The last chapter provides an

overall conclusion aligned with the objectives and recommendations for future research.

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1.7 References

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CHAPTER TWO

MICROPLASTIC PHYSICAL TOXICITY TO

FRESHWATER AND MARINE INVERTEBRATES AND FISH

2.1 Abstract

Microplastic ingestion has been well documented in aquatic ecosystems in both in

vivo and in situ in a variety of pelagic and benthic invertebrates and fish. However, due

to the many limitations involved in field sample collection and characterization methods,

and those associated with the exposure to uniform and homogeneous microplastic

exposures in situ, it is difficult to truly evaluate the toxicological effects of microplastics

in the environment. Acute polyethylene microplastic (MP) sphere exposures (0-100,000

MPs/mL) to the freshwater amphipod, Hyalella azteca, and fathead minnow, Pimephales

promelas, and marine copepod, Amphiascus tenuiremis, and mummichog, Fundulus

heteroclitus, were conducted in order to quantify MP ingestion, egestion, and mortality.

Within the range of polyethylene MP exposure concentrations used, MP ingestion was

dose-dependent and gut residence time did not significantly differ from that of naturally

ingested food materials (i.e. Tetramin®, algae) in all four organisms. Mortality was not

observed during the 4-d acute exposures to P. promelas, and F. heteroclitus. The 10-d

and 7-d lethal concentration 50% (LC50) values were 4.64 x 104 and 3.98 x 104 MPs/ml

for H. azteca and A. tenuiremis, respectively. Chronic polyethylene MP sphere exposures

(0-20,000 MPs/mL) to H. azteca and A. tenuiremis were conducted to evaluate

toxicological effects on growth and reproduction. A 42-d chronic exposure to

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polyethylene MPs significantly decreased H. azteca growth and reproduction at the low

(5,000 MPs/mL) and intermediate (10,000 MPs/mL) exposure concentrations. Exposure

of A. tenuiremis to polyethylene MPs for 32 days resulted in significant reductions in

growth (represented by significant increases in time spent in each developmental stage) at

intermediate (5,000 MPs/mL) and high (10,000 MPs/mL) concentrations, whereas

reproduction was not affected at any MP exposure concentrations. Ten-d acute (1-90

MPs/mL), and 42-d chronic (0-22.5 MPs/mL) polypropylene fiber MP exposures to H.

azteca were also conducted in order to evaluate whether MP characteristics (i.e. shape

and polymer) may influence organism survival, MP egestion, as well as growth and

reproduction. Polypropylene MP acute exposures resulted in an LC50 of 71.43 MPs/mL,

and the data suggest that the increased gut residence time of MP fibers resulted in

significant reductions in growth and reproduction. These data demonstrate MP toxicity is

specie-specific. Although MP ingestion may have prevented exposed organisms from

ingesting sufficient nutrients, resulting in reductions in survival, growth and

reproduction, it is difficult to attribute these effects without evaluating additional

endpoints (i.e. energy reserves, food uptake).

2.2 Introduction

Quantifying and characterizing microplastic (MP) presence in the environment

and the resulting toxicity from their exposure has been a topic of much discussion. Due

to the long residence time of plastic debris in the environment and exposure to various

types of physical, chemical and biological degradation pathways, most of the plastic that

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is present in aquatic ecosystems is small. Approximately 65 and 72% of the plastic

debris recovered from beaches is less than 5 and 1 mm, respectively (Martins and Sobral

2011; Browne et al. 2010). The upper limit of 5 mm has been widely used to define a

“microplastic”, although it is more intuitive to use 1 mm as the upper size limit, thus

making it difficult to compare field data regarding environmental MP concentrations and

determining what MP exposure concentrations to use in situ (Arthur et al. 2009; Costa et

al. 2010).

It has been estimated that over five trillion pieces of plastic debris currently

pollute oceans worldwide, with more than one billion of MPs per day discharged from

the Los Angeles River into the Pacific Ocean (Eriksen et al. 2014; Moore et al. 2005).

Approximately 8 and 10% of freshwater and marine fish, respectively, sampled from the

Gulf of Mexico had MPs in their gut tract, and fish sampled from urbanized streams

ingested more MPs than those from non-urbanized streams (Phillips and Bonner 2015).

Wild marine invertebrates (Mytilus edulis and Arenicola marina) contained an average of

0.2 and 1.2 MPs/g tissue, respectively, where the greater ingestion of MPs observed in A.

marina was attributed to their being non-selective feeders (Van Cauwenberghe et al.

2015). Up to 27.6% of the detritivorous invertebrates sampled in the Calvi Bay, France,

had ingested MP fibers (Remy et al. 2015). Although the number of organisms with

ingested MPs, and number of MPs ingested per organism may seem insignificant, these

data demonstrate the need to understand the varying susceptibility of organisms

inhabiting areas close to urbanized areas that will generate high amounts of plastic waste

and potential sinks for different types of MP.

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One of the reasons why studies available so far have demonstrated such a

spectrum of toxicological effects is because it is difficult to examine every type of MP

with different amounts and types of additives (that are also incorporated into a plastic

product in different ways), shapes, and sizes in situ. Few studies have investigated the

effects of multiple types of MPs on different organisms, thus making it difficult to

compare thresholds of effect.

Few published studies thus far have investigated the effect of MPs on freshwater

or marine fish in situ, and although many studies have examined MP exposures on

primarily marine invertebrates, it is challenging to make comparisons across studies due

to differences in MPs examined, exposure reporting methods (i.e. concentrations in terms

of mass, total MP counts), and exposure conditions and periods. Objective one for my

study focused on establishing a threshold of acute effects for freshwater and marine

invertebrates and fish to provide a platform on which the effects can be compared by

using similar bioassay methods for four organisms (Hyalella azteca, Amphiascus

tenuiremis, Pimephales promelas, and Fundulus heteroclitus). Chronic exposures to H.

azteca and A. tenuiremis were conducted to examine sublethal effects (i.e. development,

reproduction) in order to investigate the potential for multigenerational effects. There are

no published data available regarding microplastic toxicity to any of the invertebrates and

fish used in this study.

H. azteca is ubiquitously found in freshwater habitats and has a broad diet that

includes particulates in the size range of MPs. Their sensitivity to different types of

environmental pollutants and relative simplicity in culture maintenance have resulted in

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the common use of H. azteca in both freshwater sediment and water toxicity bioassays

(USEPA 2000). Amphipods are representative of large portions of biomass present in

freshwater ecosystems, and are a key food source for fish and birds (MacNeil et al. 1997;

Chapman 2007). Furthermore, amphipods are primarily benthic organisms that are

responsible for a large amount of leaf litter breakdown (Blarer and Burkhardt-Holm

2016). Amphipods contribute many ecological services, and the consumption of MP

fibers in vivo and in situ marine and freshwater amphipods, respectively, demonstrates

the need to further explore how these data compared to the toxicity of spherical MPs

(Blarer and Burkhardt-Holm 2016; Remy et al. 2015).

Zooplankton provide a necessary and important service to aquatic ecosystems,

and are the base of many aquatic food webs. As free-floating heterotrophic aquatic

organisms, zooplankton health is often monitored as a biomarker for pelagic

contaminants. A. tenuiremis is one of the marine zooplankton used for water toxicity

tests for similar reasons to those listed above for H. azteca (Chandler 1986). A few

studies have demonstrated that the exposure of MPs to marine zooplankton has resulted

in the blockage of the digestive tract, and mechanically affected feeding and digestion

processes (Setälä et al. 2014). Zooplankton exposed to 0.05µm polystyrene MPs resulted

in a dose-dependent developmental delay and reduction in fecundity, whereas growth and

reproduction effects were not seen with the larger MPs (0.5 and 6µm) (Jeong et al. 2017).

Further, the gut retention of polystyrene MPs (7.3-30.6 µm) of up to seven days by

zooplankton implies that longer exposures could result in the constant presence of MPs

within the digestive tracts of various important primary consumers in aquatic ecosystems

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(Cole et al. 2013). Available data suggest MPs can be readily consumed by both benthic

and pelagic invertebrates, suggesting that larger organisms may be susceptible to the

consumption of MPs through their diet.

Fathead minnows (Pimephales promelas) are ubiquitously found in North

America and are often used in freshwater toxicity bioassays (USEPA 1989). However,

no studies thus far have investigated the toxicological effects of microplastics on fathead

minnows. The common freshwater fish, Hoplosternum littorale, collected from South

America had ingested more MPs less than 5mm (86% of the total plastic debris ingested),

than MPs in larger class sizes (Silva-Cavalcanti et al. 2017). Furthermore, more H.

littorale ingested MPs in urbanized sections of the river sampled (Silva-Cavalcanti et al.

2017). Again, organisms inhabiting urbanized areas will likely have higher and more

consistent exposures to MPs. However, there is very little information known about the

toxicity of microplastics to freshwater organisms. Thus far, the only freshwater fish that

has been exposed to MPs in situ is zebrafish, Danio rerio (Lu et al. 2016; Karami et al.

2017). Polystyrene MPs 7-d acute exposures resulted in the accumulation of MPs in the

gills, gut and liver, as well as inflammation and lipid accumulation in the liver (Lu et al.

2016). Exposure to polystyrene and low-density polyethylene MPs also resulted in

oxidative stress (Lu et al. 2016; Karami et al. 2017). Further, metabolomics analysis

revealed that lipid and energy metabolism may be negatively affected by MP exposure in

zebrafish (Lu et al. 2016). Although more information is available on marine fish and

invertebrates in regards to MP toxicity, current data have shown that MPs are also

prevalent in freshwater ecosystems. Thus, given that freshwater organisms are just as

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likely (if not more) to be exposed to MPs, the following data regarding MP toxicity to

fathead minnows will help fill that research gap.

Mummichogs (Fundulus heteroclitus) are found in many coast tidal creeks, and

have been used in estuarine and marine toxicity tests. A commonly used bait fish,

mummichogs are exposed to environmental pollutants that come from both freshwater

and marine sources as the tide moves in and out. MPs have been found in marine fish

collected from many geographical locations, and habitats (i.e. pelagic, demersal, benthic),

resulting in MP ingestion ranging between 1 and 7 MPs ingested/organism (Jabeen et al.

2017; Nadal et al. 2016). Further, the number of MPs ingested by marine fish may be

determined by feeding preferences (i.e. omnivorous, herbivorous, carnivorous), with

greater numbers of ingested MPs in the digestive tracts of omnivorous fish (Mizraji et al.

2017). Feeding mechanism as well as habitat may also influence fish susceptibility and

exposure to MPs; however, this may only occur in specific areas, as feeding preference

did not influence MP ingestion in fish found in the Gulf of Mexico (Phillips and Bonner

2015). The only study thus far that has investigated MP toxicity in situ with marine fish

species demonstrated that polystyrene MP exposures can result in decreased growth, and

alter feeding preferences of larval European perch, Perca fluviatilis (Lönnstedt and Eklöv

2016). Although information is available on MP ingestion by marine organisms, there is

little data on the toxicological ramifications of the prevalent and ubiquitous MP

exposures in marine ecosystems worldwide. Mummichogs and fathead minnows are

surface and benthic omnivorous feeders, respectively, however both have been

accustomed to come to the surface in situ; thus their feeding strategies had to be adjusted

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in the following bioassays to evaluate microplastic toxicity (USFWS 1985; Hanson et al.

2005).

2.3 METHODS

2.3.1. Organism Culture

Freshwater amphipod Hyalella azteca

H. azteca was obtained from the U.S. Geological Survey Columbia

Environmental Research Center, Columbia, Missouri (USGS- Columbia) and cultured

following protocols from USEPA standard methods (USEPA 2000). Stock cultures were

maintained in 10-L aquaria containing hard reconstituted water (HRW) at 25± 1°C under

a 16:8-h light:dark photoperiod. HRW was prepared using a revised version of the

Borgmann (1996) recipe, where the bromide concentration was decreased from 800 to 40

µg Br/L. The recipe for HRW consisted of 55 L MILLIPORE MILLI-Q® water (18

mega-ohm), 8.8 g CaCl, 1.64 g MgSO4, 0.21 g KCl, 4.62 g NaHCO3, and 3.3 mg NaBr

(hardness (mg/L): 125; alkalinity (mg/L): 48). Ten aged maple leaves were added to the

culture aquaria weekly. The aging process for maple leaves followed methods outlined in

USEPA standard methods to reduce the occurrence of organisms in the substrate, and to

remove the residuals of naturally occurring tannic acid prior to placement in the culture

(USEPA 2000). To obtain known-age juvenile amphipods for the 10- and 42-day

exposures, mature amphipods were placed in 1.0-L glass mason jars for one week. After

reproduction, the adults were removed, and the less than 7-day old neonates were used in

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the bioassays. The neonates were counted and placed in temporary holding chambers for

24 hours prior to starting the exposure to account for possible mortality due to handling.

Marine copepod Amphiascus tenuiremis

A. tenuiremis used in this study was obtained from the University of South

Carolina (Colombia, SC) and cultured from original field stocks maintained in sediments

and collected from a pristine site in the North Inlet estuary, South Carolina, USA

(Chandler 1986). A tenuiremis undergo three life stages (naupliar, copepodite, and adult)

and can reproduce after 15 days, when raised in 30 ppt seawater at 25°C (Chandler et al.

2012). Stock cultures were maintained in 5-L aquaria containing 30 ppt seawater at 25±

1°C under a 16:8-h light:dark photoperiod. Instant Ocean Aquarium Salt (Instant Ocean,

Aquarium Systems) was used to prepare 30 ppt artificial seawater. Prior to being used,

the 30 ppt artificial seawater was filtered (0.22 µm) and aerated to >95% O2 saturation

for 24 hours. Stock cultures were fed twice a week with the marine microalgae

Dunaliella tertiolecta. D. tertiolecta was cultured in continuously aerated 30 ppt

artificial seawater (same as that used for the copepod culture) under fluorescent light and

fed to copepods when cultures reached a cell density of approximately 7 x 106 cells/mL.

Freshwater fish Pimephales promelas

P. promelas were cultured at the Clemson University Institute of Environmental

Toxicology (CU-ENTOX) in a static renewal system, containing moderately hard water

(MHW) with a water turnover rate of 5 times a day (hardness = 80-100 mg/L as CaCO3;

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alkalinity =57-64 mg/L as CaCO3; pH = 8) using USEPA standard methods (USEPA

1989). Temperature was maintained at 25°C ± 1°C with a 16/8 h light/dark photoperiod.

Tanks with actively breeding fish contained 15-20 males, 60-80 females (3:8, male:

female), and substrate (polyvinyl chloride pipe tiles) on which eggs were laid and

fertilized. Polyvinyl chloride pipe (PVC) tiles containing fertilized fathead minnow

embryos were removed from the troughs daily, and placed in 1.5 L tanks containing

moderately hard water (artificially prepared using the USEPA recipe) that was statically

renewed on a daily basis. Fertilized embryos took approximately 3-5 days to hatch. The

PVC tiles were examined daily, and the resulting fry removed from the tanks containing

unhatched fertilized embryos, and placed in 4 L tanks with aerated moderately hard water

and fed less than 48 hour old Artemia spp. for 30 days. Artemia spp. cysts (i.e. dormant

eggs) were purchased and kept refrigerated until needed for juvenile fish feedings. When

needed, Artemia spp. cysts were placed in a conical flask containing 1 L of 30 ppt salt

water. The flasks were well aerated and were maintained in the same temperature and

photoperiod as P. promelas. Juvenile fish between 30 and 90 days old were fed crushed

Tetramin®, and fish older than 90 days old were fed Tetramin® and frozen adult Artemia

spp.

Marine fish Fundulus heteroclitus

Fundulus heteroclitus were collected using baited minnow traps in tidal creeks

and pools of the North Inlet estuary located near Georgetown, South Carolina.

Mummichogs are lunar spawners, and reproduce primarily during the spring and summer.

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Fertilization occurs externally between mating pairs in salt marshes when high tides are

characteristically higher (in association with full and new moons). Bait traps were laid at

low tide during the new and full moon between May and August 2016 for a period of 3-6

hours per day. Adult mummichogs captured were separated by sex and kept in buckets

with aerated salt water while unfertilized embryos were stripped from female fish greater

than 3 inches in length (smaller fish were released back into the wild upon capture).

Female fish were removed briefly from the water, wrapped in a soft cloth, and their

abdomens were stroked to extrude unfertilized embryos. Specifically, the area between

the pectoral fins and the anus was stroked gently. Approximately 200 embryos were

collected per glass petri dish, and 10 mL of artificially prepared 18 ppt salt water using

Instant Ocean Sea Salt were added to each petri dish until fertilization occurred. Female

fish were kept out of the water for less than 30 seconds in order to reduce distress to the

animals. When unfertilized embryos were successfully collected, the female fish were

placed in a separate bucket containing aerated salt water; female fish were released back

into the wild after all the fish had been examined. Typically, one male provides enough

sperm to fertilize 250-300 embryos; however, two adult male mummichogs were

euthanized with an overdose of buffered MS-222 (1 g/L). The testes were removed,

placed in a separate glass petri dish containing 1-2 mL of 18 ppt salt water and mashed

into pieces using a razor blade to release the milt (i.e. semen of a male fish). The milt

was then poured over the unfertilized embryos and gently moved in a singular circular

motion to make sure all unfertilized embryos made contact with sperm. Fertilization took

approximately 10-15 minutes, and the milt was then removed from the fertilized embryos

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by gently pipetting all the milt out of the petri dishes. The fertilized embryos were then

gently rinsed with 18 ppt salt water. Temperature was maintained at 25°C ± 1°C with a

16/8 h light/dark photoperiod. The embryos were monitored daily; unfertilized and

unhealthy embryos (those with a dark coloring) were removed. During this time, the

water was also statically renewed every 24 hours. Fertilized embryos required

approximately 2-3 weeks to hatch. The petri dishes were placed on a shaker table

(slowest setting) every day after the second week in order to stimulate hatching. Hatched

fry were removed from the petri dishes, and placed in a 1.5 L tank with aerated 18 ppt

salt water. Less than 48 hour old Artemia spp. were also fed to newly hatched

mummichogs before and during MP bioassays.

2.3.2 Microplastic Exposure Water Preparation

Polyethylene microplastic spheres

The fluorescent blue polyethylene MP particles used were 10-27 µm in diameter,

and had a density of 1.13 g/cc (blue fluorescence: 435 nm peak emission; Cospheric

LLC). The polyethylene MP particles were provided as a powder. The size of fluorescent

blue polyethylene MPs was confirmed using an Olympus LEXT OOLS4000 3D Laser

Measuring Microscope (Clemson Light Imaging Facility). MP suspensions were

prepared as stock suspensions in ethanol; the ethanol was allowed to evaporate prior to

pouring fresh or salt water in the exposure beakers. The low density of ethanol allowed a

greater amount of MPs to remain in suspension as compared to MP solutions made in

either fresh or salt water (18 and 30 ppt), resulting in consistent measurements of MP

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concentrations. As described in Table 2.1, MP concentrations for all bioassays were

prepared and measured twice using a hemocytometer.

Table 2.1: Summary of microplastic acute and chronic exposure concentrations for both freshwater and marine invertebrates and fish. Organism Exposure

Water Acute Exposure Concentration (MPs/mL)

Acute Exposure Period (days)

Chronic Exposure Concentration (MPs/mL)

Chronic Exposure Period (days)

Hyalella azteca (Freshwater)

Hard Reconstituted Water (HRW)

0 - 100,000 10 0 - 20,000 42

Amphiascus tenuiremis (Marine)

30 ppt artificially prepared salt water

0 – 80,000 7 0 – 10,000 32

Pimephales promelas (Freshwater)

Moderately Hard water (MHW)

0 – 100,000 4

Fundulus heteroclitus (Marine)

18 ppt artificially prepared salt water

0 – 100,000 4

Polypropylene microplastic fibers (Hyalella azteca only)

Marine rope that had been sitting on an area of unshaded concrete for three years was

used to create the black polypropylene MP fibers that H. azteca were exposed to during

the acute bioassay. The rope was thoroughly cleansed with MILLIPORE MILLI-Q®

water for 24 hours, and then manually cut to the smallest possible lengths with sharp

scissors. The polymer composition of the rope was characterized by using attenuated

total reflection (ATR) infrared spectroscopy (Agilent Technologies Cary 680 FTIR;

74

Clemson University’s Advanced Materials Research Laboratory), equipped with a

diamond crystal. Thirty-two scans were done with a resolution of 4 cm-1, and no ATR

correction. It was thus determined that the marine rope was composed of polypropylene.

The polypropylene MP fiber size was characterized as 20 - 75 µm in length, with a

diameter of 20 µm, using an Olympus LEXT OOLS4000 3D Laser Measuring

Microscope. Individual polypropylene MP fibers were individually counted to prepare

each exposure concentration in triplicate and weighed using a Mettler Toledo AT201

Analytical Balance; the average mass of polypropylene MP fibers was then used to

prepare the exposure concentrations for the acute and chronic exposures to polypropylene

MP fibers 0 – 90, and 0- 22.5 MPs/mL, respectively.

2.3.3 Toxicity Bioassays


Toxicity of MPs to H. azteca was determined using revised USEPA methods for

conducting 10- to 42-day water-only toxicity exposures (USEPA 2000). Acute and

chronic bioassays were conducted in a static-renewal mode, where HRW renewals

occurred three days each week and the organisms were fed daily with both Tetramin®

and diatoms (Ivey et al. 2016). Revisions to the standard feeding methods (USEPA

2000) included altered feeding regimes with incremental increases in Tetramin® Tropical

Fish Food Flakes and diatoms concentrations were increased as exposed organisms

increased in age (Table 2) (Ivey et al. 2016). Preparations for diatom and Tetramin®

feeding regiments are described further below due to these methods being different than

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those used in standard USEPA methods (USEPA 2000). All organisms were exposed in

250 mL beakers at 25°C and a photoperiod of 16L/8D in the same HRW in which

organisms were cultured.

To initiate the 10-d acute and 42-d chronic exposures, ten juvenile amphipods

(less than 7-d old) were randomly chosen and placed in 250 mL beakers. The 10-d acute

exposures involved three replicate beakers per treatment to examine mortality, growth,

and MP ingestion. The 42-d chronic exposures involved 12 replicate beakers per

treatment, at which three replicates were destructively sampled after 10 days, three

replicates were offered after 28 days, and six replicates were offered after 42 days. After

10, 28, and 42 days, survival, growth, and MP ingestion and egestion were examined.

Additionally, reproduction was examined between 28 and 42 days of MP exposure.

Acute 10-d, and chronic 42-d exposures to polyethylene MP particles and polypropylene

MP fibers were conducted.

Diatom culture

Diatoms (Thalassiosira weissflogii 1200TM) were purchased from

ReedMariculture, Inc (Campbell, CA). This diatom liquid stock had an original

concentration of 320 million cells/mL and a shelf life of four months when stored at 4°C.

Forty mL of the diatom stock were diluted with 160 mL of HRW and centrifuged for 15

minutes at 2800 revolutions per minute (rpm). The supernatant was discarded and the

diatoms diluted again with 60 mL of HRW. After being centrifuged and decanted, 210

mL of control water were added to the concentrated diatom pellet. The final suspension

had a conductivity of less than 700 µS/cm. The dry weight (DW) concentration of

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diatoms in the final slurry was determined by drying 10 mL of the final diatom slurry in

an oven for 24 hours at 70°C. The final concentration of the diatom slurry used during

the bioassays was adjusted to 6.0 mg DW/mL. Diatom feeding concentrations were

increased over time during the bioassays to accommodate increased appetites of larger

organisms (Table 2.2). A renewed diatom slurry was prepared every two weeks and

stored at 4°C during all bioassays.

Tetramin® Tropical Fish Food Flake preparation

Tetramin® Tropical Fish Food Flakes were purchased and crushed until all flakes

were less than 355 µm in diameter. Suspensions of Tetramin® with HRW were added to

exposure beakers, and were stored at 25°C for three days. These suspensions were made

as needed throughout the 10- and 42-day bioassays, in addition to being used during the

egestion assay described below. Tetramin® feeding concentrations were increased over

time during the bioassays to accommodate increased appetites of larger organisms (Table

2.2).

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Table 2.2: Ramped Diet Used in 10-day and 42-day Hyalella azteca exposures

Week of Exposure (Days) Ramped Diatom Diet Ramped Tetramin® Diet

1 (0-6) 0.5 mg/beaker-day 0.25 mg/beaker-day






Adapted from revised USEPA and ASTM Hyalella azteca toxicity test methods, by

Ivey et al. 2016.


While there is a standard chronic toxicity bioassay for using juvenile A.

tenuiremis, there are no standard toxicity bioassays for using adult A. tenuiremis. It was

believed that nauplii do not consume very much in the first few days prior to developing

into a copepodite. A 7-d exposure was conducted to quantify the toxicity of polyethylene

microplastics. One hundred adult copepods were exposed in petri dishes with 20 mL of

30 ppt artificial seawater (same water used for copepod culture) and a 16:8-h light:dark

photoperiod. Exposure concentrations (0 – 80,000 microplastics/mL) to polyethylene

microplastic particles were measured twice using a hemocytometer. Each copepod was

observed every 24h at 40x total magnification with a Meiji EMT Stereo Microscope to

monitor microplastic ingestion and egestion, and survival. During daily water renewals,

glass Pasteur pipettes were used to slowly remove live copepods from the exposure petri

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dishes while minimizing the amount of exposure water taken up. The copepods were

placed in clean petri dishes, and petri dishes were covered with glass petri dish covers in

order to minimize water loss. Copepods were fed approximately 20,000 live algal cells

(D. tertiolecta) per copepod every day following daily water renewals.

Protocols from the Standard Guidelines for Conducting Renewal Microplate-

Based Life-Cycle Toxicity Tests with a Marine Meiobenthic Copepod were used to

conduct full 35-d life-cycle bioassays (Chandler 2004). Amphiascus tenuiremis nauplii

less than 24 hours old were individually placed into 30 microwells of triplicated 96-well

hydrogel-coated microplates (Corning). Three hours prior to the addition of nauplii and

MP exposure solutions, hydrogel coatings were hydrated with water to make the gel

hydrophilic. All organisms were individually exposed in microwells containing 250 µL

of artificial seawater (same water used for copepod culture) and a 16:8-h light:dark

photoperiod. Microplastic exposure concentrations were prepared in fresh renewal

seawater every 3d over the 35d bioassay. Exposure concentrations (0 – 10,000

microplastics/mL) to polyethylene microplastic particles were measured twice using a

hemocytometer. Copepods were fed approximately 20,000 live algal cells (D. tertiolecta)

per copepod every 6d. Each copepod was observed every 24h at 40x total magnification

with a Meiji EMT Stereo Microscope to monitor stage-specific mortality, time to juvenile

copepodite stage, time to adult, sex upon adulthood, and microplastic ingestion and

egestion. At adulthood, individuals were mated pairwise within each microplate

treatment, and placed in fresh microwells. Each pair was observed every 24h for clutch

size, and number of viable hatched offspring (nauplii) was quantified for two clutches.

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Freshwater fish Pimephales promelas

Toxicity of MPs to P. promelas was determined using revised USEPA methods

for conducting 96-hr toxicity exposures (USEPA 2002). Using static renewal mode, 96-

hr acute bioassays were conducted, where MHW renewals occurred every day, and the

organisms were fed daily with less than 48-hr old Artemia spp. Revisions to the standard

feeding methods included the use of 96-hr old instead of less than 24-hr old fathead

minnows. Fathead minnows when cultured at 25°C still attain nutrients from their yolk

sac up until three days after hatching (USEPA 1996); the exposure started on day four

after hatching in order to ensure MPs would be ingested by exposed juvenile fathead

minnows. All organisms were exposed in 250 mL beakers at 25°C and a photoperiod of

16L/8D in the same MHW in which organisms were cultured. Ten randomly chosen

fathead minnows were used per replicate, with five replicates per treatment. Exposure

concentrations (0 – 100,000 microplastics/mL) of polyethylene microplastic particles

were measured twice using a hemocytometer.

Marine fish Fundulus heteroclitus

There is no standardized acute or chronic bioassay for F. heteroclitus, despite this

organism being used in countless aquatic toxicological studies (e.g., Klaunig et al. 1975).

Acute toxicity of MPs to F. heteroclitus was determined by adapting USEPA methods for

conducting 96-hr toxicity exposures with fathead minnows (USEPA 2002). Similar to the

fathead minnow 96-hr acute bioassays described above, 96-hr bioassays were conducted

where 18 ppt salt water (same as that used for mummichog embryo development) was

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statically renewed and mummichogs were fed daily with less than 48-hr old Artemia spp.

Revisions to the standard feeding methods included the use of 96-hr old instead of less

than 24-hr old mummichogs. When cultured at 25°C, mummichogs still attain nutrients

from their yolk sac up until five days after hatching (US Fish and Wildlife 1985); the

juvenile mummichogs that hatched were observed to lose the yolk sac three days after

hatching. The exposure started on day four after hatching in order to ensure MPs would

be ingested by exposed juvenile mummichogs. All organisms were exposed in 250 mL

beakers at 25°C and a photoperiod of 16L/8D in the same 18 ppt salt water in which

organisms were cultured. Five randomly chosen mummichogs were used per replicate,

with three replicates per treatment. Exposure concentrations (0 – 100,000

microplastics/mL) of polyethylene microplastic particles were measured twice using a

hemocytometer.

2.3.4 Microplastic ingestion analysis

Freshwater amphipod Hyalella azteca & marine copepod Amphiascus tenuiremis

Tricaine-S (MS 222) was used to temporarily anesthetize and immobilize the

invertebrates to facilitate ingestion analysis for amphipods exposed to polyethylene MP

particles. Briefly, a stock of Tricaine-S (MS 22) was prepared by dissolving 400 mg

Tricaine-S (MS 222) in 97.9 mL distilled water, and adjusting to a pH 7.0 with

approximately 2.6 mL of 1 M Tris. Invertebrates were washed with clean water at each

sampling point and placed into Lab-Tek® 4 Chambered #1 Borosiliate Coverglass

System with a prepared solution of 15% Tricaine for 30 minutes. Preliminary 30 minute

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exposures to 15% Tricaine demonstrated 100% survival for organisms. Uptake of

fluorescent polyethylene MPs was quantified using a Leica SP8X multiphoton spectral

confocal microscope (Leica Microsystems, Buffalo Grove, IL; Clemson Light Imaging

Facility) equipped with a 10X (HC PL APO; NA=0.40) objective lens. Digital zoom

from 1.2 to 1.7 was used, depending on the size of the specimen. The fluorescent

particles were excited with a 405 nm laser and emittance from 430 to 475 nm was

detected using a photomultiplier tube (PMT). The images were further analyzed using

Leica LAS AF software (Leica Microsystems; Clemson Light Imaging Facility). This

process took approximately 2 min per replicate. After quantifying polyethylene MP

particle ingestion, the immobilized invertebrates were immediately washed with clean

water three times prior to being placed in a 12-well polystyrene flat bottom plate with

water.

Polypropylene MP fiber ingestion by amphipods was evaluated following the

egestion assay, described below. Due to the non-fluorescent nature of the polypropylene

MP fibers, the ingestion of polypropylene MP fibers could not be done using confocal

imaging. The fecal pellets egested following the egestion assay were dissected and the

number of ingested polypropylene MP fibers was quantified by manually counting

individual fibers.

Freshwater fish Pimephales promelas & marine fish Fundulus heteroclitus

Ingestion of high density polyethylene microplastics by fathead minnows and

mummichogs was quantified after euthanizing the fish in buffered MS-222. The fish

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were rinsed three times using MILII-Q-PORE water, and then individually homogenized

in 8 mL of MILLI-Q PORE water. Four mL of 70% HNO3 were then added (final

concentration: 17.5% HNO3), and vortexed for approximately one minute in order to

dissolve the tissue. The number of microplastics present in the digestive tract was

counted using a hemocytometer.

2.3.5 Microplastic egestion analysis

Following microplastic exposures, the same egestion analysis was used to analyze

microplastic egestion by all four organisms. Surviving organisms in the replicates were

exposed to 1.0 g/L of activated carbon (Norit™, Decolorizing carbon neutral) for 20

minutes, and washed with clean water three times. Exposure to the activated carbon

produced a dark section in the digestive tract. Organisms were then placed in clean water

(the water used for each respective organism as described in the culture section above)

with 1.0 mg of Tetramin® (amphipods, fathead minnows, mummichogs) that was less

than 355 µm in diameter, or 20,000 live algal cells (D. tertiolecta) per organism

(copepods). Egestion time was measured by examining each organism under a Meiji

EMT Stereo Microscope at 40X total magnification until the dark plug was completely

egested. Egestion rates were quantified post-exposure to Tricaine in preliminary studies;

there was no significant difference in the egestion rates between the organisms exposed

and not exposed to Tricaine.

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2.3.6 Growth and reproduction analysis


During chronic exposures, neonates in each exposure beaker were counted and

removed at every water change so as not to confuse them with adults later in the

bioassay. The total number of neonates per replicate was then normalized to the number

of females (determined at between days 28 and 42) per replicate. The sex of the adult

organisms was determined by examining each organism at 40X total magnification with a

Meiji EMT Stereo Microscope. The number of adult females was determined by

counting the adult males; adult male amphipods have enlarged second gnathopods, and it

was assumed that the other adults were females (USEPA 2000). Organisms were then

transferred into microcentrifuge tubes containing 50% phosphate buffer solution (PBS)

and 50% glycerol to preserve the integrity of the tissue samples until DW was ready to be

quantified. These organisms were then washed with HRW three times and placed in an

oven at 65°C for 48 hours. The DW per surviving organism in each replicate was

measured using a Mettler Toledo AT201 Analytical Balance.


During chronic exposures, growth was quantified by counting the number of days

it took for each amphipod to reach different developmental stages; the amount of time it

took for nauplii and copepodites to develop into copepodites and adult copepods,

respectively, were evaluated. After all copepods within one 96-well plate developed into

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adults, copepods were randomly paired, and the number of nauplii produced per brood

for two broods was quantified.

Freshwater fish Pimephales promelas & marine fish Fundulus heteroclitus

Reproductive effects were not quantified for fathead minnows and mummichogs;

however, effects on growth were evaluated by comparing dry weights of fish between

treatments, prior to and after microplastic exposures.

2.3.7 Data analysis

The following endpoints: 1) mortality, 2) reproduction, 3) growth, 4) MP

ingestion, and 5) MP egestion were compared with an one-way analysis of variance

(ANOVA) to determine if there were significant effects of MP type and concentration.

This was followed by a series of contrasts evaluated with a least-squared means

difference Student’s t-tests to determine mean differences between treatments exposed to

polyethylene MP particles or polypropylene MP fibers and those that were not exposed to

either MP. These calculations were performed using JMP Pro 11 Statistical Discovery

software (SAS, Cary, NC, USA).

Trimmed Spearman-Karber method was used to estimate LC50 and obtain the 10-

d median lethal concentration with 95% confidence intervals (CI) for both polyethylene

spherical and polypropylene fiber MP particles. These calculations were performed using

ESTSK Version 3.10 (Erich D. Strozier 1992-1997).

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2.4 Results

2.4.1 Freshwater amphipod: Hyalella azteca

Acute toxicity of microplastics: polyethylene spherical MPs and polypropylene

fibrous MPs

Acute exposure to fluorescent blue polyethylene MP particles resulted in a dose-

dependent increase in both number of MP particles consumed by H. azteca and mortality

with a 10-d LC50 of 4.6 x 104 MPs/mL (Figure 2.1). Significantly higher mortality

occurred when the amphipods were exposed to polyethylene MP particle concentrations

greater than 1,000 MPs/mL suggesting this is the threshold of effect. Despite large

variation in the number of polyethylene MPs consumed per surviving amphipod, in

general, the number of polyethylene MPs ingested per organism was significantly higher

at concentrations greater than 10 MPs/mL. While the number of ingested polyethylene

MP particles was significantly greater at concentrations ≥100 MPs/mL, mortality was

only significantly greater at concentrations ≥ 1,000 MPs/mL. Growth (measured in dry

weight (mg)/organism) was not significantly different between organisms exposed and

not exposed to polyethylene MP particles (data not shown). Furthermore, the amount of

time for organisms to egest polyethylene MP particles (approximately 2 hours) was not

significantly different from the amount of time it takes for organisms to egest natural

food items, nor was it significantly different across exposures to different concentrations

of polyethylene MP particles (data not shown).

Acute exposure to polypropylene MP fibers resulted in a 10-d LC50 of 71

MPs/mL (Figure 2.2). Furthermore as a result of the acute polypropylene MP fiber

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exposure, mortality (Figure 2.2B), and gut clearance time (Figure 2.2D) increased in a

dose dependent manner, whereas growth (Figure 2.2C) decreased in a dose dependent

manner. Organisms exposed to 45 and 90 polypropylene MPs/mL, experienced

significantly higher mortality (p<0.001) as compared to organisms that were not exposed

to polypropylene MP fibers. These same exposures to polypropylene MP fibers also

resulted in significant reductions in growth (p<0.05) at the end of the 10-d acute

bioassay; organisms exposed to 45 and 90 MPs/mL weighed approximately 50 and 65%

less than the control, respectively. The amount of time that it took organisms to excrete

the polypropylene MP fibers was also significantly greater (p<0.001) than the amount of

time it took for control organisms to excrete natural food items. When exposed to 45 and

90 MPs/mL, amphipods took two and four times longer, respectively, to egest gut content

containing polypropylene MP fibers. These data suggest that the shape, and possibly

polymer type, have a large influence on MP residence time within the gut.

Consequentially, a longer residence time may suggest a longer time in which the

organism is not taking in food items and proper nutrients necessary for growth, and

reproduction.

Chronic toxicity of microplastics: polyethylene spherical MPs and polypropylene

fibrous MPs

Chronic 42-d exposures to polyethylene MP particles by H. azteca resulted in a

non-dose-dependent effect on reproduction. Specifically, polyethylene MP particle

exposures of 5,000 and 10,000 MPs/mL resulted in a significant decrease in reproduction

87

(p<0.05) occuring until day 28 (Figure 2.3C); no neonates were produced in either of

these exposures, despite the presence of mature female and male amphipods.

Reproduction was also significantly reduced (p<0.05) at polyethylene MP particle

exposures of 10,000 MPs/mL at day 42, where only one out of the six replicates had any

neonate production. The effect on reproduction at day 42 was not dose-dependent. The

chronic bioassay was repeated and the second bioassay yielded results that were not

statistically different (p>0.05) from the first bioassay.

Similarly, chronic 42-d exposures to polyethylene MP particles resulted in a non-

dose-dependent effect on organism growth at all exposure concentrations (Figure 2.3B).

Organism growth was significantly less than the control in the lowest concentration

(5,000 MPs/mL) at days 10 and 28 (p<0.05), and in the intermediate concentration

(10,000 MPs/mL) at days 28 and 42 (p<0.05). Interestingly, growth was not significantly

different than that of the control at any of the sampling timepoints at the highest

concentration of polyethylene MP particles (20,000 MPs/mL). Although the effect of

polyethylene MP particles on growth was not dose-dependent, and followed the same

trend as reproduction in the first chronic bioassay, it should be noted that there was no

significant difference in the second chronic bioassay with regards to organism growth.

During the chronic exposures to polyethylene MP particles, the ingestion of MPs

(measured as number of MPs per organism) was only significantly different (p<0.05)

from the controls for day 28 organisms exposed to 10,000 MPs/mL (Figure 2.3A). The

number of MPs ingested per organism tended to decrease over the course of the 42-d

exposure in all polyethylene MP exposures (Figure 2.4A-C).

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Gut clearing times did not differ significantly between any of the exposure

concentrations to polyethylene MP particles and controls. There was also no significant

difference in mortality between any of the treatments in the chronic bioassay (data not

shown). The presence of MPs in the hind gut and in the egested fecal pellets (Figure

2.4D-E) suggests that the amphipods were able to egest polyethylene MP particles

measuring 10-27 um in diameter.

Chronic 42-d exposures to polypropylene MP fibers resulted in significantly

higher mortality (p<0.05) at the highest exposure concentration (22.5 MPs/mL) on day 10

and 28, and at the intermediate exposure concentration (11.25 MPs/mL) on day 10

(Figure 2.5). Interestingly, no significant mortality was observed among the treatments

on day 42.

Further, chronic 42-d exposures to polypropylene MP fibers resulted in a

significant decrease (p<0.05) in growth (dry weight/organism), where growth was

reduced by approximately 40- 60% on day 10, 23-40% on day 28, and 8-10% on day 42

(Figure 2.6).

Chronic 42-d exposures to polypropylene MP fibers resulted in a dose-dependent

effect on reproduction, where exposures greater than 5.6 polypropylene MP fibers/mL

resulted in significant decreases (p<0.05) in neonate production (Figure 2.7).

Specifically, by day 28 in the 42-d exposure, there was no neonate production in both the

intermediate and highest exposures to polypropylene MP fibers (11.25 and 22.5 MPs/mL,

respectively). At the end of the 42-d exposure to polypropylene MP fibers, neonate

production was 70% less than that observed in the control (0 MPs/mL) in both the

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intermediate and highest exposures. Similar to the chronic exposure to polyethylene

microplastic particles, treatments in which growth was negatively affected, also

experienced negative effects for reproduction when organisms were exposed to

polypropylene microplastic fibers. Interestingly, despite significant reductions in growth

observed in amphipods exposed to the lowest concentration of polypropylene MP fibers

(5.6 MPs/mL), there was no reduction seen in neonate production on days 28 and 42.

Where chronic exposures to polyethylene MP particles did not result in a

significant difference in gut clearing times (when compared to natural feeding materials),

exposures to polypropylene MP fibers concentrations greater than 5.6 MPs/mL resulted

in significantly higher egestion times (p<0.05) (Figure 2.8). There was a dose-dependent

effect on polypropylene MP fiber egestion. The presence of MPs in the egested fecal

pellets suggest that the amphipods could egest polypropylene MP fibers, although at a

slower pace than that of polyethylene MP particles or natural food materials.

2.4.2 Marine copepod Amphiascus tenuiremis

Acute toxicity of polyethylene spherical MPs

Acute exposure to fluorescent blue polyethylene MP particles by A. tenuiremis

resulted in a dose-dependent increase in both number of MP particles consumed and

mortality (7-d LC50 of 7.9 x 104 MPs/mL) (Figure 2.9). Although not statistically

significant, MP ingestion by exposed copepods was dose-dependent, suggesting the dose-

dependent mortality observed may be attributed to MP ingestion; MP ingestion may have

satiated exposed copepods, thus potentially resulting in decreased intake of sufficient

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nutrients. MP egestion was quantified, and adult copepods could egest MPs in the same

amount of time it took to egest algae (data not shown). Data regarding fluoranthene

bioavailability in Figure 2.9 will be further discussed in Chapter Three.

Chronic toxicity of polyethylene spherical MPs

The number of polyethylene MPs ingested per organism depended on both MP

exposure concentrations and developmental stage for A. tenuiremis. Nauplii observed at

day 8 did not ingest a significant amount of MPs across any of the MP exposure

treatments (Figure 2.10); however, Figure 2.11A shows that some nauplii were able to

ingest one MP at day 8 of the exposure. Despite the nauplii showing little MP ingestion,

there was a dose-dependent effect on growth; nauplii exposed to MP concentrations

greater than 2,500 MPs/mL resulted in significantly longer developmental times (Figure

2.11B). Exposure to 10,000 MPs/mL resulted in significantly greater quantities of MPs

(p<0.05) being ingested by both copepodites and adult copepods (Figure 2.10). Figure

2.11C depicts the ingestion of MPs by adult copepods. Adults also ingested significantly

higher quantities of MPs at both the lowest and intermediate MP exposure concentrations

(2,500 and 5,000 MPs/mL, respectively). Copepodites that were exposed to 5,000 and

10,000 MPs/mL resulted in significantly longer developmental times (p<0.05) to become

adults (Figure 2.11D). Interestingly, exposure to MPs did not result in any reproductive

effects; organisms exposed to MPs had brood sizes that were statistically the same

(Figure 2.12).

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Gut clearing times did not differ significantly between any of the exposure

concentrations to polyethylene MP particles and controls (data not shown). There was

also no significant difference in mortality between any of the treatments in the chronic

bioassay.

2.4.3 Freshwater fish Pimephales promelas

MP ingestion was dose-dependent, with significantly higher MP ingestion

(p<0.001) observed in MP exposures greater than 100 MPs/mL (Figure 2.13). Juvenile

fathead minnows acutely exposed to polyethylene MPs did not result in significant

mortality at any of exposure concentration (0-100,000 MPs/mL) (data not shown). There

was also no significant difference in MP gut residence time as compared to that of brine

shrimp (data not shown). No chronic bioassays were conducted for fathead minnows,

thus no effects on growth or reproductions were evaluated.

2.4.4 Marine fish Fundulus heteroclitus

MP ingestion was dose-dependent, with significantly higher MP ingestion

(p<0.001) observed in MP exposures greater than 1000 MPs/mL (Figure 2.14). Juvenile

mummichogs acutely exposed to polyethylene MPs did not result in significant mortality

at any of exposure concentration (0-100,000 MPs/mL) (data not shown). There was also

no significant difference in MP gut residence time as compared to that of brine shrimp

(data not shown). No chronic bioassays were conducted for mummichogs, thus no effects

on growth or reproductions were evaluated.

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2.5 Discussion

MP ingestion was dose-dependent in acute exposures for both invertebrates and

fish; however, mortality was only observed when the invertebrates, H. azteca and A.

tenuiremis, were exposed to concentrations of polyethylene MPs greater than 10,000

MPs/mL. Interestingly, during chronic exposure to polyethylene MP particles, there was

a significant increase in MP ingestion only when H. azteca were exposed to 10,000

MPs/mL at day 28. With regards to polyethylene MP particle ingestion, higher

variability was observed during the H. azteca chronic exposure, and a distinct trend was

observed where fewer polyethylene MP particles were ingested as the duration of the

chronic exposure continued. Although the opposite trend was seen during the chronic

exposure with A. tenuiremis, the dose-dependent ingestion of MPs may be due to the MPs

having a similar size (10-27 µm) as those of the algal cells (D. tertiolecta) normally

consumed by A. tenuiremis. As amphipods grew to larger sizes throughout the chronic

exposure to polyethylene MP particles, it is possible that upon reaching a larger size,

organisms were capable of and preferred to ingest larger food materials (i.e. Tetramin®

and diatoms), resulting in fewer polyethylene MP particles being ingested at later

developmental stages. Amphipods exposed to polypropylene MP fibers (approximately

20-75µm in length, 20µm in diameter) experienced greater mortality at lower

concentrations (45 MPs/mL) than the exposure to polyethylene spherical MPs. But

similar to the polyethylene MPs, by day 42, no significant mortality was observed for

either MP exposure.

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Juvenile fathead minnows and mummichogs have mouths that are magnitudes

larger than those of copepods and amphipods, suggesting that the MPs may have been

passively ingested with other food materials and water. Although size selection was not

specifically evaluated in the present study, field collected organisms have been found to

consume plastic particles with specific diameters (Bråte et al. 2016; Cole et al. 2013).

Atlantic cod (Gadus morhua) sampled from the Norwegian coast preferentially consumed

mesoplastics (5-25 mm), despite having available plastic particles ranging from 3.2 to

41.7 mm in size (Bråte et al. 2016). In regards to invertebrates studies, zooplankton

exposed to polystyrene MPs ranging in size from 7.3 to 30.6 µm overall preferably

ingested 7.3 µm polystyrene MPs, with different species displaying a varying preference

for 20.6 and 30.6 µm polystyrene MPs (Cole et al. 2013). Thus, size selection may

dictate which species or at which developmental stages organisms are susceptible to a

particular MP size.

With regards to MP egestion, the egestion times for polyethylene MP particles

were not significantly different between any of the exposure concentrations and the

control for any of the four organisms evaluated. This trend was also observed in

zooplankton exposed to polystyrene MP spherical particles (Cole et al. 2013), where MP

egestion rates were similar to those of natural food items. Organisms exposed to

microplastic particles that are rounded also seem to be able to egest microplastics easily.

Spherical microplastics were present in the fecal matter and egestion times were not

significantly longer for microplastics as compared to normal food materials (Au et al.

2015; Hamer et al., 2014). Following the egestion assays, no polypropylene MP fibers

94

remained in the guts of exposed amphipods in either acute or chronic exposures. Despite

significantly longer gut clearance times for ingested polypropylene MP fibers as opposed

to polyethylene MP particles during the acute exposure, my study demonstrated that

polypropylene MP fibers can be completely egested by amphipods when given enough

time to depurate the particles. Copepods exposed to MPs egested them in densely packed

fecal pellets, demonstrating that MPs were pushed to certain areas within the gut prior to

being egested (Cole et al. 2013). Although the fecal pellets egested by the amphipods

and copepods both had MP aggregates present, images of the gut tract did not suggest the

MPs were packaged in any specific manner prior to being egested. In examination of the

amphipod feces, both types of MPs were evident and fully incorporated into the pellet.

How MPs are arranged in the gut after being ingested may influence an organism’s

ability to egest them, thus also affecting their retention time within the gut. The same

observation was the case for both fathead minnows and mummichogs, where

polyethylene MPs were quickly egested and randomly distributed within the feces.

The feces of both freshwater and marine organisms settled to the bottom of

exposure beakers and tanks; the formation of larger aggregates containing high

concentrations of MPs suggests that benthic organisms (regardless of a size preference in

food materials) may be more susceptible to MPs (Long et al. 2016). My data suggest that

the ingestion of MPs by either pelagic or benthic organisms greatly influences the

transport and fate of MPs in both freshwater and marine ecosystems. In the present study,

MPs examined were present throughout the gut in exposed organisms and did not

aggregate in a specific section of the gut. There was also no evidence of MP

95

translocation out of the gut. Other studies have shown the tendency of MPs to aggregate

within (Cole et al. 2013; Murray and Cowie 2011) or translocate from the gut (Browne et

al. 2008) in invertebrates. As in other aquatic invertebrates, such as isopods (Hamer et al.

2014), and copepods (Cole et al. 2013), given enough depuration time, H. azteca and A.

tenuiremis were capable of egesting MPs. The different ingestion and egestion of MPs as

demonstrated by H. azteca in the present study, could account for the varying toxicity

seen with regards to growth and reproduction.

My results demonstrate that the polypropylene MP fibers used in the present study

were more toxic to H. azteca than the polyethylene MP particles, where toxicity to

polypropylene MP fibers followed a dose-dependent response, and toxicity to

polyethylene MP particles did not. During the acute exposure to polypropylene MP

fibers, as exposure concentrations increased, there was a corresponding significant

increase in MP ingestion and egestion, and a decrease in growth, whereas an acute

exposure to polyethylene MP particles did not result in significant reductions in growth

or egestion time in response to increasing polyethylene MP particle exposures. A chronic

28-d exposure to polyamide MP fibers resulted in a reduction in assimilation efficiency in

the freshwater amphipod Gammarus fossarum, suggesting that despite the ability to egest

MP fibers if provided a depuration period, the ability to attain enough nutrients may still

be affected (Blarer and Burkhardt-Holm 2016).

Increased toxicity due to polypropylene MP fiber exposures may be attributable to

the difference in shape, as it has been suggested that MP shape may have a significant

influence on MP fate and transport (Wright et al. 2013), thus potentially also impacting

96

the toxicity on exposed organisms. For example, the lobster, Nephrops norvegicus, was

shown to be unable to completely egest polypropylene fibers, leading to the retention of

fibers within the chitinous foregut of the animal, which resulted in decreased organism

growth (Murray and Cowie 2011). When crustaceans molt, they shed the chitinous lining

in their fore- and hindguts in addition to their exoskeleton. Fibers, possibly caught in

these less flexible structures, result in their retention within the gut. The aggregation of

fibers in the gut of the lobsters exposed to polypropylene fibers, and the consequential

formation of ball-like structures may have also influenced polypropylene fiber gut

retention time (Murray and Cowie 2011). In the present study, these structures were not

formed in H. azteca exposed to polypropylene MP fibers; however, the possibility that

smaller fibers, or other materials may cause similar results cannot be ruled out. Although

not seen in my exposures to polyethylene MP particles, polystyrene MP particles can be

retained within the foregut of crabs (Carcinus maenas) resulting in a longer retention

time in the gut (Watts et al. 2014). Fathead minnows and mummichogs were not exposed

to polypropylene fiber MPs in our present study; however, the formation of fiber

aggregates structures has also been observed to occur in Atlantic cod, suggesting that

MPs may block the gastrointestinal tract of fish as well as lobsters (Bråte et al. 2016;

Murray and Cowie 2011). Fiber and fragmented MPs are found in greater abundance in

the environment as compared to spherical MPs, and the sharper edges on MP surfaces

may contribute to a greater physical harm once ingested. It is likely that the shaper edges

on the polypropylene fiber MPs used in this study contributed to the longer egestion

times in exposed amphipods. Although gut physiological changes due to MP exposures

97

were not examined as an endpoint in my exposures, it is possible that the ingested fibrous

MPs resulted in the abrasion and tearing of tissue within the gut of exposed amphipods.

The growth rates of H. azteca depend greatly on food rations, and due to reduced

food uptake and growth (as measured based on the amount of time to the onset of

amplexus), significant reductions in reproduction have also been observed by others

(Moore and Farrer 1996). Specifically, the number of neonates produced by a single

female significantly correlates with the length of the female (Othman and Pascoe 2001).

In the present study, chronic exposures to polyethylene MP particles resulted in

significant reductions in growth and reproduction that may have resulted from reduced

food intake, or an overall interference in food processing caused by ingested MP

particles. Organisms in the same exposures that experienced significant reductions in

growth (as compared to the controls), also produced significantly less neonates per

female. Copepod growth was delayed due to microplastic exposure where nauplii took

27-40% longer to develop into copepodites, and copepodites subsequently took 11-33%

longer to develop into adults. Copepodites exposed to 10, 000 MPs/mL took twice the

amount of time to become adults than organisms that were not exposed to any

microplastics. Surprisingly, however, reproduction was not influenced by the presence of

microplastics. The physical toxicity of microplastics is likely an energetics issue, and the

amount of time it takes organisms to reach sexual maturity will influence brood

developments. Although growth was delayed in exposed copepods, the results showed

that when copepodites developed into adults, reproduction was not influenced at all.

Mating occurred within 24 hours of pairing random females and males in all treatments,

98

suggesting that as long as copepods reach sexual maturity, microplastic ingestion may not

immediately affect reproduction. Exposures to microplastics that go beyond the first two

broods may show some negative effects on brood size and viable nauplii; however,

reproduction past the first two broods was not investigated in this study. Further, the

chronic effects of microplastic exposures is likely to be species-specific, where exposures

of various shapes of microplastics to the marine isopod Idotea emarginata demonstrated

a lack in discriminatory feeding, and there were no significant effects on survival, growth

and intermolt duration (Hamer et al., 2014). The population level consequences of

reduced reproduction as a result of exposure to MPs need to be explored further.

Although ingestion rates were not calculated in our study, the ingestion of inert

particles in general has been shown to decrease the ingestion and processing of food

items (Cole et al. 2013; Ayukai 1987; Fernandez 1979). This trend has been

demonstrated for clay particles (Robinson et al. 2009), and carbon nanotubes (Roberts et

al. 2007; Edgington et al. 2010) using the freshwater filter feeding crustacean, Daphnia

magna. MP ingestion may cause a mechanical hazard, resulting in the blockage of food

passage (Tourinho et al. 2010) or may cause the organism to feel satiated, indirectly

resulting in a reduction in food intake (Thompson 2006). Many invertebrates have

minimal lipid reserves, and a reduction in uptake of natural food may result in decreased

growth, reproduction, and increased mortality (Ayukai 1987).

Several factors may have confounded the comparison of toxicity data in this study

between the polyethylene MP particles and polypropylene MP fibers. First, they are

composed of different polymers. Polyethylene and polypropylene are among the most

99

extensively used plastics (Arutchelvi et al. 2008), yet toxicity studies have yet to

specifically target the effects of exposure of MPs composed of these polymers. These

polymers are biochemically inert (Teuten et al. 2009), and aside from the additional

methyl functional group present on alternating carbons in polypropylene, the structural

characteristics do not differ greatly. Second, the polypropylene MP fibers used in the

present study were aged, whereas the polyethylene MP particles were purchased. It is

possible that some of the differences seen between the acute exposures to polyethylene

and polypropylene MPs could be due to the oxygen-containing functional groups present

on the surface of polypropylene MP fibers due to the various aging processes the marine

rope endured. Finally, these plastics had different shapes. Perhaps the shape of the MPs

used in this study may have had a great influence on the differences seen in acute

exposures to the freshwater amphipod, H. azteca, based on the potential for fibers to

aggregate and interact with the gut differently than spherical MPs upon being ingested.

There are many factors that make it challenging to definitively establish the

environmental concentrations of MPs in aquatic systems, one of which is the different

sampling techniques used to measure the presence of MPs. Sampling techniques used to

quantify the presence of smaller plastic fragments include beach combing, sediment

sampling, trawls, observational studies, and biological samplings (Cole et al. 2011).

Each technique has a different limit of detection and method of separating plastic

fragments from water or sediment, which present problems because many reports have

quantified MP presence in terms of an area or volume sampled, as opposed to a

concentration value. Another issue with establishing an environmentally-relevant MP

100

concentration stems from the different definitions that are used for the term

“microplastic”, where the size range varies greatly between studies (Cole et al. 2011).

MP abundances worldwide in both freshwater and marine ecosystems have been shown

to be greater than that of larger plastic debris (greater than 5 mm), representing more than

half of the total plastic samples (Moore et al. 2011; Lattin et al. 2004; Eriksen et al.

2014). However, as shown in Table 1.1, upwards of 2.7 billion MPs have been found in

in aquatic ecosystems; the highest number of MPs used in my study was 100,000

MPs/mL exposure treatments for fathead minnows and mummichogs (approximately 50

million MPs). Although no toxicity was observed in exposed fathead minnows and

mummichogs, this same concentration of 100,000 MPs/mL (approximately 20 million

MPs) resulted in significant mortality when amphipods were exposed to MPs.

Furthermore, the field studies cited in Table 1.1 only captured MPs using mesh nets

where the MPs captured would be at least 300 µm in diameter; it is likely that

environmental concentrations of MPs similar in size to those used in my experiments (10-

27 µm in diameter) are significantly higher. In addition, the potential for smaller MPs to

have a longer residence time within exposed organisms (Rosenkranz et al. 2009)

demonstrates the need for additional research regarding the biological impact of MPs on

aquatic organisms.

Little information is currently available on the chronic toxicity of microplastics,

and nutrient uptake may be affected by the presence of microplastics; therefore,

additional studies need to be conducted to quantify endpoints other than mortality (Cole

et al. 2013; Wegner et al., 2012). MP fibers may also result in a higher bioavailability of

101

adsorbed contaminants than MP spherical particles due to MP fibers having a longer gut

residence time. Despite the lack of significant ingestion of microplastics by exposed

nauplii, the negative impact on growth was seen in both the intermediate and highest

microplastic exposure treatments. Microplastic ingestion has been seen to be highly

variable and effects on biological function does not seem to solely depend on ingestion,

as microplastics have been observed to stick to the carapace and gills of exposed

organisms (Au et al. 2015). The generally hydrophobic surface characteristics and large

surface area to volume ratio that MPs have in comparison to larger plastic products also

present the potential for these MPs to not only be contaminated by persistent organic

pollutants (POPs), but to also act as a source for various types of additives, that may

prove toxic to exposed organisms (Cole et al. 2011; Teuten et al. 2009); thus, the

presence of contaminated MPs in aquatic systems will likely alter the bioavailability of

contaminants. This research demonstrates that MPs in the aquatic environment may

affect individual growth, mortality, and reproduction in H. azteca. These adverse effects

could translate into population and community level effects, and thus, should be

characterized further in future studies. While the present study only examined the

physical effects of MPs, it is important to consider other aspects of MP exposure,

including the effect of adsorbed contaminants and of plastic additives leaching from

plastics.

102

2.6 Limitations

The focus of Chapter Two was to examine whether MPs could elicit a

toxicological response in exposed aquatic organisms. When this study began

approximately four years ago, there was limited information available on not only what

types and quantities of MPs were most prevalently found in the environment, but also

how exposures to various types of MPs would affect organisms on various biological

levels. One limitation to this study is the use of MPs with both different shapes and

polymers. Unfortunately, I was unable to determine whether MP shapes or polymers

may play a larger role in having a toxicological effect on exposed organisms. In order to

determine whether a certain MP polymer type or shape was more toxic, it would have

been beneficial to use different concentrations of MPs composed of one type of polymer,

with additional treatments that differed based on MP shape. However, understanding

the driving factors of MP toxicity in terms of MP characteristics was not the goal of this

study; the goal was to determine whether a threshold of toxicity for one MP type could be

established for multiple organisms. Additionally, another limitation of this study is the

predominant use of primary MPs (spherical polyethylene MPs). Secondary MPs are

found more prevalently in the environment than primary MPs, and the use of the

polypropylene MP fibers during these experiments would have been more

environmentally relevant. However, the fibrous MPs would have not been able to be

used for copepod exposures, as the fibers were larger than the copepods themselves; the

use of smaller, well-characterized MPs allowed observations of whether multiple

organisms were capable of ingesting MPs.

103

2.7 Conclusions

To further our understanding of the effects of MPs on both aquatic invertebrates

and fish, the present study demonstrated that despite spherical polyethylene MPs being

easily egested by exposed organisms dose-dependent effects on MP uptake were

observed in all four organisms. Further, 10- and 7-day LC50s of approximately 4.6 x 104

and 7.9 x 104 MPs/mL for spherical polyethylene MPs were quantified for amphipods

and copepods, respectively. No mortality was observed for either fathead minnows or

mummichogs at MP exposures used during these bioassays. Interestingly, polypropylene

MP fibers were more toxic to exposed amphipods than the polyethylene MP spheres,

where mortality was observed at concentrations as low as > 22.5 MPs/mL. Chronic

exposures to polyethylene spherical MPs resulted in significant reductions in growth for

exposed amphipods and copepods; however, reproduction was only significantly affected

in MP exposure treatments where amphipods also experienced growth reductions. My

results suggest that MP exposures can result in mortality, and significant reductions in

growth and reproduction at environmentally relevant concentrations.

104

2.8 Figures

Figure 2.1. Mortality and polyethylene microplastic (MP) ingestion following 10-d

acute exposure

A) Mean percent mortality (SD = ± 7.86) and B) mean MP ingestion (SD = ± 0.32)

during the 10-day PE MP chronic exposure (0 and 100,000 MPs/ml) to Hyalella azteca.

The asterisk represents statistical differences (p < 0.05) in percent mortality and number

of MPs ingested/organism between exposure to MPs and no exposure to MPs.

105

Figure 2.2. Polypropylene MP fiber ingestion, mortality, growth and egestion

following 10-d acute exposure

A) MP fiber ingestion (SD = ± 0.69), B) mean percent mortality (SD = ± 6.61), C) Mean

growth (dry weight/organism (mg)) (SD = ± 0.01), and D) Mean egestion time (hr) (SD =

± 0.92) following the 10-day PP MP fiber particles (0 and 90 MPs/mL) acute exposure to

Hyalella azteca. The asterisk represents statistical differences in mean MP fiber

ingestion, percent mortality, growth and MP egestion time (p < 0.05, p < 0.001, p < 0.05,

and p < 0.001, respectively) between no exposure and exposure to MPs.

A B

C D

106

Figure 2.3: Polyethylene MP ingestion, growth and reproduction following 42-d

chronic exposure

A) Mean MP ingestion (average number of MPs ingested per organism) (SD = ±

0.45), B) Mean growth in terms of dry weight per organism (mg/organism (SD =

± 0.06), and C) Mean reproduction (number of neonates per female) (SD = ±

107

2.39) of Hyalella azteca, in response to exposure to PE MP spherical particles

exposures (0, 5,000, 10,000 and 20,000 MPs/mL) at days 10, 28 and 42. The

asterisk represents statistical differences in mean MP fiber ingestion, growth, and

reproduction (p < 0.05) between no exposure and exposure to MPs at each time

point.

108

Figure 2.4: Images of fluorescent MP spherical particles (10-27um in diameter)

ingested by H. azteca

Images were taken with a Leica SP8X Multiphoton spectral confocal microscope at

different destructive sampling time points after being exposed to 20,000 MPs/mL for (A)

10-d, (B) 28 days, and (C) 42-d. The presence of fluorescent MP spherical particles in

the hind gut of H. azteca (D and E) and a fecal pellet (F) after being exposed to 10,000

MPs/mL at day 28 (D and E).

109

Figure 2.5: Mortality following 42-d chronic exposure to polypropylene fibrous MPs

Mean percent mortality (SD = ± 20.55) during 42-day PP MP fiber particles (0 - 22.5

MPs/mL) chronic exposure to Hyalella azteca. Different letters represent statistical

differences (p < 0.05) in percent mortality among the MP exposure treatments at specific

time points.

05

101520253035404550

10 28 42

Perc

ent m

orta

ility

Day

0 MPs/mL 5.625 MPs/mL 11.25 MPs/mL 22.5 MPs/mL

A A B B AAA AAAAB

110

Figure 2.6: Growth following 42-d chronic exposure to polypropylene fibrous MPs

Mean growth in terms of dry weight per Hyalella azteca (mg/organism (SD = ± 0.11), in

response to exposure to PP MP fiber particles (between 0 and 22.5 MPs/mL) at days 10,

28 and 42. Different letters represent statistical differences (p < 0.05) in growth among

the MP exposure treatments at specific time points.

0.0000.1000.2000.3000.4000.5000.6000.7000.8000.9001.000

10 28 42

Dry

wei

ght (

mg)

/org

anism

Day


A A B B

BBA

BBBA

B

111

Figure 2.7: Reproduction following 42-d chronic exposure to polypropylene fibrous

MPs

Mean reproduction (number of neonates per female) (SD = ± 3.22) in Hyalella azteca in

response to exposure to PP MP fiber particles (between 0 and 22.5 MPs/mL) at days 28

and 42. Different letters represent statistical differences (p < 0.05) in reproduction

among the MP exposure treatments at specific time points.

02468

101214161820

28 42

Num

ber o

f neo

nate

spe

r fem

ale

Day


AA

B BBB

A

A

112

Figure 2.8: Gut residence time of polypropylene MP fibers during 42-d chronic

exposure

Mean egestion time (measured in gut residency time (hr)) (SD = ± 17.3)) during 42-day

PP MP fiber (0 - 22.5 MPs/mL) chronic exposure to Hyalella azteca. Different letters

represent statistical differences (p < 0.05) in egestion time among the MP exposure

treatments at specific time points.

0

50

100

150

200

250

300

350

10 28 42

Aver

age

gut r

esid

ency

tim

e(to

tal m

in p

er re

p)

Day


A A A BBAA

BBAAB

113

Figure 2.9: Mortality following 7-d acute exposure to polyethylene MPs

Mean percentage mortality (SD = ± 16.9) following the 7-d acute polyethylene

microplastic spherical particle exposure (0 - 80 000 microplastics/mL) exposure to

Amphiascus tenuiremis. The asterisk represents statistical differences (p < 0.05) in

percentage mortality between microplastic exposure treatments with and without

fluoranthene (FLU).

0102030405060708090

100

20 000 30 000 40 000 80 000

Aver

age

perc

ent

mor

talit

y

Microplastic concentrations (MPs/mL)

Microplastics only FLU-satured microplastics

*

114

Figure 2.10: Polyethylene MPs ingested throughout 32-d chronic exposure Response to 32-d chronic polyethylene microplastic particle exposure (0 - 10 000

microplastics/mL to Amphiascus tenuiremis. Mean number of ingested polyethylene

microplastics per organism (standard deviation (SD) = ± 1.94) in response to exposure to

polyethylene microplastic spherical particles (0 microplastics/mL, 2 500

microplastics/mL, 5 000 microplastics/mL, and 10 000 microplastics/mL). Different

letters represent statistical differences in the number of ingested microplastics between

different microplastic exposure treatments at time points specific to each developmental

stage (d6 for nauplii; d15 for copepodites; d25 for adults prior to being randomly paired

to quantify reproduction effects).

0

1

2

3

4

5

6

7

8

Nauplius Copepodite AdultNum

ber o

f pol

yeth

ylen

e m

icro

plas

tics

inge

sted

per

org

anism

Developmental stage

0 MPs/mL 2 500 MPs/mL 5 000 MPs/mL 10 000 MPs/mL

B B B

C

A

A

A A

A A A A

115

Figure 2.11: Images and growth resulting from the ingestion of fluorescent

polyethylene MPs by nauplii and adult copepods

Amphiascus tenuiremis chronically exposed to fluorescent polyethylene microplastics

(10-27 µm in diameter) for 32 days. Images taken with a Leica SP8X Multiphoton

spectral confocal microscope of fluorescent polyethylene microplastic spherical particles

ingested by A. tenuiremis A) nauplii after being exposed to 5 000 microplastics/mL for 8

days and C) adults after being exposed to 2 500 microplastics/mL for 24 days. The black

arrows indicate the presence of polyethylene microplastic spherical particles. Mean

growth is measured in terms of the amount of time (number of days) it takes to reach the

next development stage, B) nauplii to copepodite, and D) copepodite to adult. There is a

25 µm

200 µm

A B

C D

116

statistical difference (p<0.05) in mean growth at both development stages between

exposure to microplastics (5 000 and 10 000 MPs/mL) and no exposure to microplastics.

117

Figure 2.12: Reproduction following 32-d exposure to polyethylene MPs

Mean reproduction (number of nauplii per pair) (SD = ± 1.43) in Amphiascus tenuiremis,

in response to exposure to PE MP spherical particles (between 0 and 10,000 MPs/mL).

There was no statistical differences (p > 0.05) in reproduction among the MP exposure

treatments at any of the specific time points.

0

1

2

3

4

5

6

7

8

9

1 2

Aver

age

num

ber o

f nau

plii

per p

air

Brood

0 MPs/mL 2500 MPs/mL 5000 MPs/mL 10000 MPs/mL

118

Figure 2.13. Ingestion of polyethylene MPs following 4-d acute exposure to fathead

minnows

Mean number of polyethylene microplastics (MPs) ingested per Pimephales promelas

(SD = ± 754) during a 4-d acute exposure to 0 – 100,000 MPs/mL. Different letters

represent statistical differences (p < 0.001) in the number of ingested microplastics

between different microplastic exposure treatments.

0

2000

4000

6000

8000

10000

12000

14000

0 100 1000 10000 100000

# M

Ps in

gest

ed /

orga

nism

Microplastic exposure concentration (MPs/mL)

A A

B

CC

119

Figure 2.14. Ingestion of polyethylene MPs following 4-d acute exposure to

mummichogs

Mean number of polyethylene microplastics (MPs) ingested per Fundulus heteroclitus

(SD = ± 417) during a 4-d acute exposure to 0 – 100,000 MPs/mL. Different letters

represent statistical differences (p < 0.001) in the number of ingested microplastics

between different microplastic exposure treatments.

0

1000

2000

3000

4000

5000

6000

7000

8000

9000

0 100 1000 10000 100000

# M

Ps in

gest

ed /

orga

nism

Microplastic exposure concentration (MPs/mL)

A A

A

B

C

120

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Edgington, AJ, Roberts, AP, Taylor, LM, Alloy, MM, Reppert, J, Rao, AM, Mao, J, and Klaine, SJ. 2010. The influence of natural organic matter on the toxicity of multiwalled carbon nanotubes. Environ Sci Technol 29: 2511-2518.

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CHAPTER THREE

TOXICITY OF FLUORANTHENE-CONTAMINATED MICROPLASTICS TO FRESHWATER AND MARINE INVERTEBRATES AND FISH

3.1 Abstract

There is limited information on the ability for microplastics (MPs) to act as a

vector for other contaminants that may have either been incorporated in the original

plastic product (additives), or those that may have adsorbed onto and absorbed into the

plastic after being released into the environment. Both organic and inorganic

contaminants have been extracted from field-collected microplastics, suggesting that

microplastics are a source for environmental contaminants to exposed organism. This

study focuses on the investigation of whether polyethylene MPs can influence the

bioavailability of the polycyclic aromatic hydrocarbon (PAH), fluoranthene (FLU), to

exposed freshwater amphipods, Hyalella azteca, fathead minnows, Pimephales promelas,

and marine copepods, Amphiascus tenuiremis, and mummichogs, Fundulus heteroclitus.

All four organisms were acutely exposed to fluoranthene-contaminated polyethylene

microplastics (FLU-MPs) and the results suggest that FLU is bioavailable to organisms

upon ingestion of FLU-MPs, and that MPs can acts a vector for organic contaminants in

aquatic ecosystems.

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3.2 Introduction

Microplastics (MPs) can efficiently adsorb persistent organic pollutants (POPs),

and the high presence of MP debris has increased the concern related to the potential role

of MPs as a vector for POPs in aquatic ecosystems (Cole et al. 2011; Ivar do Sul and

Costa 2014; Koelmans et al. 2014). In addition to affecting energy production, oxidative

metabolism, and neurotransmission, exposure to contaminated MPs increased the

bioavailability of POPs in fish, mussels, and lugworms (Oliveira et al. 2013; Rochman et

al. 2013; Avio et al. 2015; Besseling et al. 2013). Further, in the presence of digestive

fluids that simulate cold and warm blooded organisms (different pH and temperature),

greater desorption of phenanthrene and dichlorodiphenyltrichloroethane (DDT) from

polyvinyl chloride MPs was observed (Bakir et al. 2014). Since more than 90% of plastic

debris can consist of MPs, it is likely that continuous ingestion of contaminated MPs will

result in the bioaccumulation of POPs and other contaminants in fish and mammals

(Eriksen et al. 2014; Desforges et al. 2015). However, many researchers have questioned

the validity and environmental relevance of contaminated MPs used in laboratory

experiments, claiming that other natural particulate matter (i.e. organic matter, sediment)

are more significant contaminant vectors (Koelmans et al. 2016). When exposed to

fluoranthene and polystyrene microplastics, fluoranthene uptake was attributed to algae

consumption by Mytilus spp.; however, the presence of MPs did significantly decrease

the overall amount of fluoranthene partitioning to algae, despite low ratios of

microplastic to algae (in terms of mass) during the 7-d exposure (1:289) (Paul-Pont et al.

2016). Additional information is needed regarding how MPs will influence the

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bioavailability of different contaminants, in the presence of other sorbents, as it is likely

that the different chemical and physical MP properties will ultimately decide contaminant

sorption onto and off of MP surfaces. Similarly, the source of organic matter and

environmental conditions will determine the structure and presence of functional groups

on organic matter, also influencing POP transport and fate in aquatic ecosystems. Both

chemical and physical characteristics of organic matter and MPs will determine the

sorption of environmental contaminants onto the surface.

Most natural organic matter has functional groups that contain oxygen, making

them polar to some extent, and the number and types of these functional groups will

impact how these macromolecules will interact with surrounding water molecules and

contaminants (Schwarzenbach et al. 2003). For instance, proteins have both polar and

non-polar portions in their structure that allow them to fold and unfold in certain ways

depending on the pH, temperature, and binding of certain ligands. There are also a

variety of proteins in different types of tissue, and the sorption of PAHs onto proteins

may differentiate based on small changes in protein types and differences in their

structures as the types of domains exposed to the aqueous environment depends heavily

on the pH of the system (Schwarzenbach et al. 2003). In addition, as lignin has a lower

oxygen to carbon ratio than cellulose or chitin (0.33, 0.84, and 0.64: 1, respectively),

lignin has a greater affinity for PAHs relative to the other biogenic molecules

(Schwarzenbach et al. 2003).

Likewise, the source of plastics is just as important in determining their structure

and resulting contaminant sorption capacity. The crystallinity of MPs (which determines

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the amount of surface area available for contaminant sorption), which will greatly

influence how plastics interact with pollutants and natural organic matter, depends on the

temperatures, solvents, and polymers involved in plastic production (Guo et al. 2012).

Three polymer structural properties that impact MP crystallinity are chain length, chain

branching, and inter-chain bonding (Gowariker et al. 1986). As polymer chain length

increases, there is an increase in polymer hardness and ductility; increased chain length

also allows for greater surface area for pollutants to sorb onto. Specifically, some planar

compounds like polycylic aromatic hydrocarbons (PAHs) are not very flexible (i.e.

fluoranthene), and although a plastic may have large amounts of surface area due to the

availability of micropores, the adsorption and absorption of inflexible planar PAHs may

not be proportional to the total surface area because these compounds will not be able to

physically get to or fit in the smaller pore spaces (Sperling 2006). A uniformed inner-

matrix with larger pore spaces will be more amenable to the sorption of larger chemicals,

although these plastics may technically have less surface area.

Chain branching in the polymer structure will impact PAH sorption (Guo et al.

2012), where the presence of hydrophobic domains will influence persistent organic

pollutant sorption onto plastic surfaces. Polypropylene has a methyl group that is present

on alternating carbons on the main backbone chain, and it generally has a greater surface

area than polyethylene. This will generally result in greater amounts of PAH adsorption

and absorption as there are more sorption sites on the surface as well as within the MP

inner matrix. Finally, inter-chain bonding present in the MP will also influence PAH

sorption as greater sorption will be noted in polymer chains that are not able to closely

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align themselves with one another. Similar to particulate organic matter (POM), MPs can

also contain functional groups that will make their surfaces more or less hydrophobic.

Plastics are ubiquitous because their production can be easily adjusted to their

intended purpose. Because of this versatility, the coatings and additives incorporated into

their structures will also greatly influence pollutant ad- and absorption. For example,

plastics are used to a great extent in supplies (i.e. nets, ropes, and clothing) used in

aquaculture and other fishery-related activity. The coatings on polypropylene marine rope

are different than those used on polyester clothing as the marine rope will need to repel

water to a greater extent than everyday clothing; marine rope has a greater capacity for

contaminant sorption than the fibers clothing may shed (unless one refers to a raincoat or

outdoor-type clothing) and are transported to the environment from washing machine

effluent (Napper and Thompson 2016).

The degradation of POM and MPs (or larger plastics to produce secondary MPs)

can also greatly influence contaminant sorption. In the environment, POM and MPs are

both subjected to physical, themo- and oxidative degradation. POM residues may change

due to diagenesis, which refers to the degradation and rearrangement of molecules from

the original macromolecule during the aging process (Schwarzenbach et al. 2003).

Mature organic matter that has had more time to experience diagenesis generally has a

lower oxygen to carbon ratio (Schwarzenbach et al. 2003), which may suggest that older

organic matter has a greater capacity for POP sorption. MPs also age in the environment,

and the addition of oxygen-containing functional groups (e.g. carbonyl, carboxyl)

decreases the hydrophobicity of the polymer, thereby reducing the sorption of

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hydrophobic compounds onto MP surfaces (Endo et al. 2005). Discolored microplastics,

or those containing a more yellow color were found to have higher concentrations of

PCBs than non-discolored pellets (Endo et al. 2005); however, their study was unable to

determine whether the yellow coloring was due to degradation pathways or was the

original color. The accumulation of biofilms and biological growth on the surface of

microplastics in the environment may result in an increased sorption of contaminants,

such as heavy metals and organic pollutants, as compared to virgin microplastics

(Holmes et al. 2012; Schorer and Eisele 1997). As MPs age, the cracking in the surfaces

also provides greater surface area for POP and environmental contaminant sorption. It is

likely that as MPs age, the potential for contaminant sorption will increase similarly to

POM.

Fluoranthene (FLU) was selected because it is a model polycyclic aromatic

hydrocarbon (PAH) that is one of the USEPA’s Priority Pollutants regulated in Clean

Water Act programs (USEPA 2014). FLU is a frequently monitored carcinogen

according to recommendations by the European Union Scientific Committee for Food

and the European Union as FLU is commonly found in the environment in mixtures with

other PAHs. Environmental concentrations of FLU in streams and wastewater effluent

can reach concentrations surpassing 10 and 500 µg FLU/L (Irwin 1997); variables

affecting FLU concentrations in aquatic ecosystems include season, climate, proximity to

urban and manufacturing plants, as well as materials surrounding water bodies (asphalt,

coal tar linings). The exposure of polyethylene MPs contaminated with the PAH, pyrene,

increased the bioavailability of pyrene to exposed fish compared to PAHs alone (Oliveira

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et al. 2013). It is likely that exposure to FLU in the presence of polyethylene

microplastics will also result in increased FLU bioavailability to both freshwater and

marine invertebrates and fish in situ.

Hyalella azteca, Amphiascus tenuiremis, Pimephales promelas, and Fundulus

heteroclitus were acutely exposed to fluoranthene-contaminated polyethylene

microplastics (FLU-MPs). The bioavailability of fluoranthene was quantified using

whole body burden analysis. Two acute bioassays per organism were conducted, one

involving the exposure to FLU-MPs, and the second involving the exposure to water-

borne fluoranthene concentrations that had been observed as desorbed from FLU-MPs

during the first bioassay. A 42-d chronic water-borne fluoranthene exposure to H. azteca

was also conducted in order to determine whether fluoranthene body burdens would

differ in adult amphipods, and influence reproduction.

3.3 METHOD

3.3.1 Characterization of fluoranthene adsorption to microplastics

Adsorption isotherm

Adsorption isotherms were developed to describe the interaction between

fluoranthene and polyethylene microplastics. The same polyethylene microplastics used

in bioassays in Chapter Two (microplastic characterization and manufacturing details

described in 2.3.2 Microplastic Exposure Water Preparation) were used in the following

bioassays that investigated fluoranthene bioavailability to amphipods, fathead minnows,

copepods, and mummichogs. Adsorption of fluoranthene to polyethylene microplastics

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was characterized using a full factorial experimental design with three concentrations of

microplastics and eight concentrations of fluoranthene for a total of 24 treatments. All

treatments were run in triplicate 20 mL glass scintillation vials, and adsorption isotherms

of fluoranthene to microplastics were quantified through fluorospectrometry. A stock

concentration of 1.0 g fluoranthene/mL was made in methanol, and stored at 0°C until

used. Fluoranthene treatments (0 – 200 µg FLU/L) were made in the USEPA recipe for

reconstituted hard water, and 30 ppt artificial seawater and controlled to 0.1% volume

fraction of methanol to avoid a co-solvent effect. Masses of 0.001, 0.01, and 0.1 g of

polyethylene microplastics were added to scintillation vials, sealed with aluminum-foil-

lined Teflon screw caps, and allowed to come to equilibrium on a rotary tumbler at room

temperature (23°C± 1°C) for 7 days. The water within the scintillation vials was analyzed

for fluoranthene in black polystyrene 96-well plates with a Molecular Devices Gemini

fluorescence microplate reader at 280/440 nm excitation/emission. Analysis of a

methanol rinse of the scintillation vials indicated negligible loss by fluoranthene sorption

to the scintillation vial walls; therefore, sorbed fluoranthene to polyethylene microplastics

could be calculated directly by analyzing the aqueous fluoranthene concentration in each

fluoranthene-microplastic suspension.

Isotherm modeling

Experimental data were transformed and fit with the Langmuir model. The

goodness of fit was analyzed by comparing the model constants and the correlation

coefficients (R2). The Langmuir model was used to quantify fluoranthene adsorption to

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PE MPs over 7 days, and determine whether all binding sites on PE MPs could be

saturated with fluoranthene.

Fluoranthene-saturated microplastic preparation

PE MPs were weighed using a Mettler Toledo AT201 Analytical Balance and

mixed with 200 µg FLU/L- spiked water. Microplastic-fluoranthene solutions were

placed on a rotary tumbler at room temperature (23°C± 1°C) for 48 hours. The

microplastic-fluoranthene solutions were then vacuum filtered using 0.45 µm nylon

filters, and rinsed with clean water. The fluoranthene-saturated microplastics were re-

suspended in clean water. Different exposure treatment solutions were made as needed

prior to every water renewal, and microplastic concentrations were quantified using a

hemocytometer.

Fluoranthene adsorption to the microplastic beads was further verified using

multiphoton microscopy. This was done using a Leica SP8X multiphoton spectral

confocal microscope (Leica Microsystems, Buffalo Grove, IL; Clemson Light Imaging

Facility) equipped with a 40X (HC PL APO; NA=1.1, water immersion) objective lens.

Multiphoton imaging was conducted using a Chameleon S, femto-second pulsed infrared

(IR) laser (Coherent, Inc., Santa Clara, CA) with an excitation of 710 nm. Fluorescent

emission from 406 to 445 nm was detected using a Leica HyD1, GaAsp detector. The

detection of adsorbed fluoranthene was not affected by the fluorescence of the

microplastic or the presence of different amounts of salts in any of the different water

compositions.

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3.3.2 Fluoranthene-contaminated microplastic bioassays

3.3.2.1 Freshwater amphipod Hyalella azteca

Fluoranthene-contaminated polyethylene microplastic exposure

Toxicity of fluoranthene-contaminated polyethylene MPs (FLU-MPs) to H.

azteca was determined using revised USEPA methods for conducting 10-day water-only

toxicity exposures (USEPA 2000). Acute bioassays were conducted in a static-renewal

mode, where both hard reconstituted water (HRW) renewals containing FLU-MPs and

the feeding of TetraMin and diatoms occurred daily (Ivey et al. 2016). Revisions to the

standard feeding methods (USEPA 2000) included altered feeding regimes. The

concentrations of TetraMin Tropical Fish Food Flakes and diatoms fed to exposed

organisms increased incrementally throughout the exposure (Ivey et al. 2016).

Preparations for diatom and TetraMin feeding regiments are described in 2.3.3 Toxicity

Bioassays: Diatom culture, due to these methods being different than those used in

standard USEPA methods (USEPA 2000). All organisms were exposed in 250 mL

beakers at 25°C and a photoperiod of 16L/8D in the same HRW in which organisms were

cultured.

To initiate the 10-d acute exposures, ten juvenile amphipods less than 7-d old

were randomly chosen and placed in 250 mL beakers. The 10-d acute exposures involved

6 replicate beakers per treatment to examine mortality and fluoranthene bioavailability.

Organisms were exposed to 0, 5,000, 10,000, and 20,000 FLU-MPs/mLs. These MP

concentrations are well below the LC20 of PE MPs observed in previously conducted MP

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acute bioassays (see Chapter Two), and were not predicted to cause significant mortality

in FLU-MP acute bioassays.

Following the 10-d acute exposure to FLU-MPs, amphipods were gently rinsed

three times in clean HRW. As observed in the previous 10-d acute MP toxicity bioassay,

polyethylene MPs are excreted by juvenile amphipods in approximately four hours, thus

following exposure to FLU-MPs, amphipods were placed in clean HRW, which was

statically renewed every 30 minutes, for four hours to ensure all ingested MPs has been

excreted. Amphipods were then placed in 1 mL amber glass vials. Fluoranthene whole

body burdens were quantified using methods described below in 3.3.3 Fluoranthene

bioavailability quantification.

Water-borne fluoranthene exposure (No microplastics)

Throughout the bioassay, fluoranthene desorbed from the polyethylene

microplastics between the daily water renewals of fluoranthene-saturated microplastics.

Measurements of the exposure water revealed that 0, 3.5, 10, and 18 µg FLU/L

corresponded with 0, 5,000, 10,000, and 20,000 FLU-MPs/mL (Table 3.1), respectively,

with no statistical difference in measured water concentrations of fluoranthene between

replicates. To differentiate between the bioavailable fluoranthene from ingested

microplastics or from exposure water, a subsequent USEPA standard 42-d bioassay was

conducted (feeding regiment revised according to Ivey et al. 2016) where amphipods

were exposed only to the water-borne concentrations of fluoranthene measured during the

fluoranthene-saturated microplastic bioassay (0-36 µg FLU/L). A 14-d depuration period

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was added to the end of the 42-d exposure to fluoranthene to investigate whether exposed

amphipods were able to eliminate fluoranthene that was taken up during the chronic

exposure. The same protocols were used as the fluoranthene-saturated microplastics 7-d

bioassay in regards to the exposure set up, daily water renewals, feedings, and

monitoring. Fluoranthene exposure solutions were also prepared as needed by using a

stock concentration of 1.0 g fluoranthene/mL (made in methanol, and stored at 0°C until

used) to spike HRW. Exposure solution was controlled to 0.1% volume fraction of

methanol to avoid a co-solvent effect.

The 42-d chronic exposures involved 19 replicate beakers per treatment, at which

3 replicates were destructively sampled after 10, and 42 days. The remaining 3 replicates

were evaluated after the 14-d depuration period. After 10, 42, and 56 (14 days post-

exposure) days, survival, growth, and fluoranthene body burden were quantified.

Additionally, reproduction was examined during the fluoranthene exposure, and

depuration period. Amphipods were then placed in 1 mL amber glass vials. Fluoranthene

whole body burdens were quantified using methods described below in 3.3.3

Fluoranthene bioavailability quantification.

Table 3.1: Hyalella azteca acute exposure to fluoranthene-contaminated microplastics (FLU-MPs), and chronic exposure to only water-borne fluoranthene (FLU)

Acute 10-d FLU-MP Exposure (#FLU-MPs/mL)

Chronic 42-d FLU Exposure: associated water

bioavailable fraction of FLU (µg FLU/L)

5,000 3.5 (n/a) 5

10,000 10 20,000 18 (n/a) 36

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3.3.2.2 Marine copepod Amphiascus tenuiremis


There are no standard toxicity bioassays that use adult copepods, and to ensure

that enough biomass could be used for quantifying fluoranthene body burden, adult

copepods (rather than juveniles as previously used in chronic exposures) used in a 7-d

exposure to FLU-MPs to investigate whether fluoranthene adsorbed to ingested

microplastics is bioavailable to exposed adult copepods. One hundred adult copepods

(50:50 male:female) were exposed in petri dishes with 20 mL of 30 ppt artificial seawater

(same water used for copepod culture) and a 16:8-h light:dark photoperiod.

Fluoranthene-saturated microplastic exposure solutions (FLU-MPs) were prepared in

fresh renewal seawater every day over the 7d bioassay. FLU-MP exposure concentrations

(0 – 80,000 microplastics/mL) were measured twice using a hemocytometer. All

copepods were observed every 24h at 40x total magnification with a Meiji EMT Stereo

Microscope to monitor microplastic ingestion, egestion, and survival. During daily water

renewals, glass Pasteur pipettes were used to slowly remove live copepods from the

exposure petri dishes while minimizing the amount of exposure water taken up. The

copepods were placed in clean petri dishes containing freshly prepared FLU-MP

solutions, and petri dishes were covered with glass petri dish covers to minimize water

loss. Following the water renewals, water samples were filtered (to remove

microplastics) and analyzed for fluoranthene using black polystyrene 96-well plates with

a Molecular Devices Gemini fluorescence microplate reader at 280/440 nm

excitation/emission. Copepods were fed approximately 20,000 live algal cells (D.

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tertiolecta) per copepod every day following the daily water renewals. After the 7-d

exposure, surviving copepods were then placed in 1 mL amber glass vials. Fluoranthene

whole body burdens were quantified using methods described below in Section 3.3.3.



microplastics between the daily water renewals of FLU-MPs. Exposure water samples

revealed that 11.2, 22.2, 59.0, and 138.6 µg FLU/L corresponded with 20,000, 30,000,

40,000, and 80,000 MPs/mL, respectively, with no statistical difference in measured

water concentrations of fluoranthene between replicates (Table 3.2). To differentiate

between the bioavailable fluoranthene from ingested microplastics or from exposure

water, a subsequent 7-d bioassay was conducted where adult copepods were exposed

only to the water-borne concentrations of fluoranthene measured during the fluoranthene-

saturated microplastic bioassay. The same protocols were used as the fluoranthene-

saturated microplastics 7-d bioassay in regards to the exposure set up, daily water

renewals, feedings, and monitoring. Fluoranthene exposure solutions were also prepared

as needed by using a stock concentration of 1.0 g fluoranthene/mL (made in methanol,

and stored at 0°C until used) to spike 30 ppt artificial seawater. Exposure solution was

controlled to 0.1% volume fraction of methanol to avoid a co-solvent effect. After the 7-

d exposure, surviving copepods were then placed in 1 mL amber glass vials.

Fluoranthene whole body burdens were quantified using methods described below in

Section 3.3.3.

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Table 3.2: Amphiascus tenuiremis acute exposure to fluoranthene-contaminated microplastics (FLU-MPs), and water-borne fluroanthene (FLU)

Acute 7-d FLU-MP Exposure (#FLU-MPs/mL)

Acute 7-d FLU Exposure: associated water

bioavailable fraction of FLU (µg FLU/L)

20,000 11.2 30,000 22.2 40,000 59.0 80,000 138.6

3.3.2.3 Freshwater fish Pimephales promelas


Standard US EPA 4-d acute fathead minnow bioassay methods were used to

investigate whether fluoranthene adsorbed to polyethylene microplastics is bioavailable

to exposed fathead minnows (USEPA 2002). Four-day old fathead minnows were also

used in an acute exposure to 100,000 MPs/mL (no fluoranthene) or 100,000 FLU-

MPs/mL to ensure that fathead minnows were capable of ingesting the MPs (USEPA

1996). Previous results demonstrated that fathead minnows exposed to 100,000 MPs/mL

did not result in significant mortality, thus any mortality that would be observed during

this exposure would likely be due to fluoranthene exposure. Ten juvenile fathead

minnows were randomly selected and placed in 250 mL beakers with MHW under a

16:8-h light:dark photoperiod. Three replicates per treatment (100,000 MPs/mL, and

100,000 FLU-MPs/mL) were used. Fluoranthene-saturated microplastic exposure

solutions (FLU-MPs) were prepared in fresh MHW every day over the 4-d bioassay (MPs

were exposed to fluoranthene for 48 hours prior to being filtered from spiked

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fluoranthene solutions and put in clean MHW for daily renewals). Solutions of MPs

without fluoranthene were also prepared daily prior to the daily water renewals. Both

MP exposure treatments were measured twice using a hemocytometer. Following the

water renewals, water samples were filtered (to remove microplastics) and analyzed for

fluoranthene using black polystyrene 96-well plates with a Molecular Devices Gemini

fluorescence microplate reader at 280/440 nm excitation/emission. Fathead minnows

were also fed brine shrimp every day following the water renewal. After the 4-d

exposure, surviving fathead minnows were euthanized using pH buffered 1.0 g/mL MS-

222, and placed in 1 mL amber glass vials. Fluoranthene whole body burdens were

quantified using methods described below in Section 3.3.3.



microplastics between the daily water renewals of the fluoranthene-saturated

microplastics. Exposure water samples revealed that 162.5 µg FLU/L corresponded with

100,000 FLU-MPs/mL, with no statistical difference in measured water concentrations of

fluoranthene between replicates. To differentiate between the bioavailable fluoranthene

from the ingested microplastics or from exposure water, a subsequent 4-d bioassay was

conducted where fathead minnows were exposed to only the water-borne concentration

of fluoranthene measured during the fluoranthene-saturated microplastic bioassay, as well

as two concentrations of fluoranthene less and greater than 162.5 µg FLU/L. The same

protocols were used as the fluoranthene-saturated microplastics 4-d bioassay in regards to

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the exposure set up, daily water renewals, feedings, and monitoring. Fluoranthene

exposure solutions were also prepared as needed by using a stock concentration of 1.0 g

fluoranthene/mL (made in methanol, and stored at 0°C until used) to spike MHW.

Exposure solution was controlled to 0.1% volume fraction of methanol to avoid a co-

solvent effect. After the 4-d exposure, surviving fathead minnows were euthanized using

pH buffered 1.0 g/mL MS-222, and placed in 1 mL amber glass vials. Fluoranthene

whole body burdens were quantified using methods described below in Section 3.3.3.

3.3.2.4 Marine fish: Fundulus heteroclitus


There are no standardized acute or chronic toxicity bioassays established for

Fundulus heteroclitus, thus similar to the protocols used in the previous chapter to

examine the acute toxicity of polyethylene microplastics to mummichogs, standard

USEPA 4-d acute fathead minnow bioassay methods were used to investigate whether

fluoranthene adsorbed to polyethylene microplastics was bioavailable to exposed

mummichogs (USEPA 2002; 1996). Four day old mummichogs were also used in this

acute exposure to 100,000 MPs/mL (no fluoranthene) or 100,000 FLU-MPs/mL to ensure

that mummichogs were capable of ingesting the MPs (USFWS 1985). Previous results

demonstrated that mummichogs exposed to 100,000 MPs/mL did not result in significant

mortality, thus any mortality that would be observed during this exposure would likely be

due to the presence of fluoranthene. Ten juvenile mummichogs were randomly selected

and placed in 250 mL beakers with 18 ppt artificially prepared sea water (same as that

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used in culture protocols) under a 16:8-h light:dark photoperiod. Three replicates per

treatment (100,000 MPs/mL, and 100,000 FLU-MPs/mL) were used. Solutions for

fluoranthene-saturated microplastic exposure (FLU-MPs) were prepared in fresh 18 ppt

sea water every day over the 4-d bioassay (MPs were exposed to fluoranthene for 48

hours prior to being filtered from spiked fluoranthene solutions and put in clean 18 ppt

sea water for daily renewals). Solutions for MPs without fluoranthene exposure were also

prepared daily prior to the daily water renewals. Both MP exposure treatments were

measured twice using a hemocytometer. Following the water renewals, water samples

were filtered (to remove microplastics) and analyzed for fluoranthene using black

polystyrene 96-well plates with a Molecular Devices Gemini fluorescence microplate

reader at 280/440 nm excitation/emission. Mummichogs were also fed brine shrimp every

day following the water renewal. After the 4-d exposure, surviving mummichogs were

euthanized using pH buffered 1.0 g/mL MS-222, and placed in 1 mL amber glass vials.

Fluoranthene whole body burdens were quantified using methods described below in

Section 3.3.3.



microplastics between the daily water renewals of fluoranthene-saturated microplastics.

Exposure water samples revealed that 200 µg FLU/L corresponded with 100,000 FLU-

MPs/mL, with no statistical difference in measured water concentrations of fluoranthene

between replicates. To differentiate between the bioavailable fluoranthene from ingested

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microplastics or from exposure water, a subsequent 4-d bioassay was conducted where

mummichogs were exposed to clean 18 ppt sea water (no fluoranthene or microplastics),

and only the water-borne concentration of fluoranthene measured during the

fluoranthene-saturated microplastic bioassay (200 µg FLU/L); no other concentrations of

fluoranthene were used due to limited number of mummichogs available. The same

protocols were used as the fluoranthene-saturated microplastics 4-d bioassay in regards to

the exposure set up, daily water renewals, feedings, and monitoring. Fluoranthene

exposure solutions were also prepared as needed by using a stock concentration of 1.0 g

fluoranthene/mL (made in methanol, and stored at 0°C until used) to spike 18 ppt

artificial seawater. Exposure solution was controlled to 0.1% volume fraction of

methanol to avoid a co-solvent effect. After the 4-d exposure, surviving mummichogs

were euthanized using pH buffered 1.0 g/mL MS-222, and placed in 1 mL amber glass

vials. Fluoranthene whole body burdens were quantified using methods described below

in Section 3.3.3.

3.3.3 Fluoranthene bioavailability quantification

Whole body burdens were quantified in order to evaluate fluoranthene

bioavailability due to FLU-MP and water-borne fluoranthene exposures. Due to the

small size (i.e. limited tissue samples) of the organisms used in the bioassays described

above, methods from Chandler et al. (2012) and Klosterhaus et al. (2002) were used to

quantify fluoranthene bioavailability. Following the exposures to FLU-MPs and water-

borne fluoranthene, H. azteca and A. tenuiremis were placed in 1 mL amber vials

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containing 0.5 mL HRW and 30 ppt sea water, respectively. The vials containing

invertebrates were then placed in -20°C for five minutes (the water within the vials did

not freeze); decreasing the temperature of the water within the vials reduces animal

movement and results in their sinking to the bottom of the vial, making it easier to

remove the overlying water within the vials. Following the exposures to FLU-MPs and

waterborne fluoranthene, P. promelas, and F. heteroclitus were euthanized with MS-222

and placed in 1 mL amber vials containing MHW, and 18 ppt sea water, respectively.

After immobilizing the invertebrates and euthanizing the fish, the overlying water

in each of the vials could then be easily removed. Fifty µL of isooctane, 10 µL methanol,

and 40 µL of 4M sodium hydroxide, and 15-30 (0.5 mm diameter) solvent cleaned

zirconia beads (Biospec11079105z Zirconia/Silica Beads) were added to each vial.

Organisms were homogenized for 100 seconds at 3,000 oscillations/min using a Mini-

BeadBeater (Biospec Products, Bartlesville, OK, USA), vortexed for 10 s, and

centrifuged for 10 minutes at 3,500 g (Klosterhause et al. 2002). The top layer of each

vial, which contained fluoranthene and iso-octane, was removed and transferred to a

clean vial and stored at – 20°C until analysis. Fluoranthene in the iso-octane was

quantified using black polystyrene 96-well plates with a Molecular Devices Gemini

fluorescence microplate reader at 280/440 nm excitation/emission.

3.3.4 Data analysis

The following endpoints: 1) fluoranthene adsorption to MPs, 2) fluoranthene body

burden due to MP ingestion, and 3) fluoranthene body burden due to external water

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exposure were analyzed using a series of one-way analysis of variance (ANOVA) tests to

determine whether there were significant differences between treatments due to MP

concentration and fluoranthene concentrations. In regards to bioassays that evaluated

whether fluoranthene was bioavailable through exposure to fluoranthene-contaminated

MPs, one-way ANOVA tests were followed by a series of contrasts evaluated with

Student’s t-tests to determine mean differences between treatments exposed to

polyethylene MP particles with or without fluoranthene. In regards to bioassays that

evaluated whether fluoranthene was bioavailable through the exposure to waterborne

fluoranthene, one-way ANOVA tests were followed by a series of contrasts evaluated

with student’s t-tests to determine mean differences between treatments exposed to

different concentrations of fluoranthene (no MPs). These calculations were performed

using JMP Pro 11 Statistical Discovery software (SAS, Cary, NC, USA).

3.4 Results

3.4.1. Fluoranthene adsorption to polyethylene microplastics

All concentrations of polyethylene MPs were saturated with FLU after 48 hours in

concentrations of FLU greater than 5 µg FLU/L for both HRW and 30 ppt artificially

prepared sea water (Figure 3.1). Concentrations of FLU greater than 40 µg FLU/L did not

result in significantly higher amounts of FLU adsorbed to polyethylene MPs at any time

points (data not shown). Confocal images of polyethylene MPs in 30 ppt artificial sea

water (Figure 3.2) were taken to ensure the presence of FLU on MPs used in the 7-d

acute bioassays. Images taken of polyethylene MPs with FLU demonstrated uneven

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adsorption of FLU on PE MP surfaces, suggesting that FLU adsorption to the PE MPs

used for the bioassays may not have been the same for each MP. Figure 3.2A serves as a

negative control for the presence of FLU on MPs. Image analysis was not able to be used

to quantify how much fluoranthene adsorbed to MPs, thus the images were used

qualitatively to confirm the presence of fluoranthene on microplastics used in the

bioassays.

3.4.2 Freshwater amphipod Hyalella azteca

Less than 7-day old Hyalella azteca were exposed to polyethylene microplastics

(5,000, 10,000, and 20,000 MPs/mL) that either were or were not saturated with

fluoranthene in a standard USEPA 10-d acute bioassay. The three concentrations of

microplastics were well below the 10-d polyethylene microplastic LC50 (4.6 x 104

MPs/mL) for H. azteca, as to ensure sufficient survival for fluoranthene body burden

analysis. As a result of the 10-d exposure, mortality was only observed at the highest

microplastic concentration (20,000 MPs/mL), where exposures to microplastics without

fluoranthene resulted in 20% mortality in exposed organisms. Exposure to 20,000 FLU-

MPs/mL resulted in 70% mortality in exposed organisms, and no significant mortality

was observed in the exposure to 18 µg FLU/L (waterborne FLU bioavailable in

exposures to 20,000 FLU-saturated MPs/mL). Although the data is not shown, no

significant mortality was observed in in either the low or intermediate FLU-saturated MP

exposures (5,000 and 10,000 FLU-MPs/mL, respectively) or exposures to water-borne

fluoranthene (0-36 µg FLU/L; Figure 3.3). The additional toxicity due to the presence of

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fluoranthene adsorbed to the polyethylene microplastics suggests that despite the gut

residence time of the microplastics not being significantly longer than that of natural food

materials, it is enough time for the desorption of fluoranthene from the microplastics

during a 10-d acute exposure. Further, the toxicity resulting from the combination of

both contaminants suggests that there is a greater than additive toxicity effect when

amphipods are exposed to contaminated MPs. In exposures to just the waterborne

fluoranthene, there was not a dose-dependent effect on mortality, despite the highest

concentration of fluoranthene (36 µg FLU/L) being double that of the amount of

fluoranthene that had desorbed off the highest concentration of fluoranthene-

contaminated microplastic exposures. The results suggest that in the ranges of

microplastics, fluoranthene, and fluoranthene-contaminated microplastics used, acute

exposures to fluoranthene-contaminated microplastics resulted in significantly higher

mortality than either microplastic or waterborne fluoranthene exposures alone.

Interestingly, significant mortality (p<0.05) was observed in all fluoranthene exposures at

day 56, after the two week depurations period (Figure 3.3). Mortality was not the goal of

the chronic exposure to fluoranthene, and throughout the 42-day chronic exposure, less

than 5% mortality was observed in all exposure treatments. However, at the end of the

depuration period, mortality significantly increased in all fluoranthene exposure

treatments except 18 µg FLU/L. Aside from that exposure, there is a dose-dependent

effect on mortality. Exposures to 5 and 10 µg FLU/L resulted in 20-25% mortality that

increased to 32% at exposures to the highest fluoranthene concentration (36 µg FLU/L).

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Treatments in which amphipods were exposed to FLU-MPs, resulted in a dose-

dependent increase in fluoranthene body burden (Figure 3.4). Specifically, exposures to

20,000 FLU-MPs/mL resulted in significantly higher fluoranthene body burdens (357 µg

FLU/mg dry weight) than exposures to 5,000 and 10,000 fluoranthene-contaminated

MPs/mL (230 and 260 µg FLU/mg dry weight, respectively). Water samples from the 10-

d exposure revealed that 3.5, 10, and 18 µg FLU/L, desorbed from the 5,000, 10,000, and

20,000 fluoranthene-contaminated MPs/mL exposures. In order to differentiate the

amount of fluoranthene that was taken up due to microplastic ingestion from the

fluoranthene that was taken up from the external water exposure, a second 10-d bioassay

was conducted where amphipods were exposed to a range of waterborne fluoranthene (0-

36 µg FLU/L). As a result of the waterborne fluoranthene 10-d bioassay, there was also a

dose-dependent fluoranthene body burden, where exposures to the highest fluoranthene

water borne concentration (36 µg FLU/L) resulted in a body burden of 84 µg FLU/mg

dry weight (Figure 3.5).

Another standard USEPA 42-d chronic toxicity test was conducted to understand

the influence of long term exposure to the same amount of waterborne fluoranthene on

sublethal endpoints such as fluoranthene body burden, and reproduction (Figure 3.6). A

depuration period of 14 days followed the 42-d exposure to fluoranthene; and the same

endpoints were quantified at day 56 as well. At day 42, the fluoranthene body burden

was quantified, and was dose-dependent. Significantly greater fluoranthene body burdens

(p<0.0001) were quantified in amphipods exposed to 18 and 36 µg FLU/L (76.0, and

73.6 µg FLU/L, respectively) as compared to organisms exposed to 0-10 µg FLU/L.

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Organisms exposed to either 18 or 36 µg FLU/L did not have significantly different

fluoranthene body burdens. Interestingly, after a 14-d depuration period, fluoranthene

body burden reduced significantly in all exposure concentrations, where there was no

significant difference in fluoranthene body burdens across any of the fluoranthene water-

borne exposure treatments, although significant mortality was observed.

In regards to reproduction, neonate production was only significantly lower

(p<0.05) than that of the control at the highest fluoranthene exposure (36 µg FLU/L),

where there was 50% less neonates produced by the present females (Figure 3.7). On day

56, reproduction increased by at least 60% in all treatments except for those organisms

that had originally been exposed to 36 µg FLU/L; reproduction did not significantly

change for organisms exposed to 36 µg FLU/L, where neonate production remained at

approximately 2 neonates per female. As compared with the previous study (Chapter

Two), growth was not quantified due to the objective of investigating body burden. Nine

replicate beakers with ten organisms per replicate were used, and there were

consequently not enough neonates to extend the experimental design. However, based on

observation alone, the organisms exposed to the highest fluoranthene concentration did

appear to physically be smaller than those in the other five treatments.

3.4.3 Marine copepod Amphiascus tenuiremis

Seven-day exposures of FLU-saturated polyethylene MPs to adult copepods

resulted in significantly greater mortality (100% mortality) in the highest MP exposure

concentration (80,000 MPs/mL; data not shown). The FLU body burden was only

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quantified in polyethylene MP exposure concentrations between 20,000 and 40,000

MPs/mL because there were no live organisms in the treatment of 80,000 FLU-saturated

MPs/mL at the end of the 7-d exposure. Exposure to FLU-saturated MPs resulted in a

dose-dependent FLU body burden, with 1.28, 2.10, and 3.41 µg FLU /organism

corresponding to exposures to 20,000, 30,000 and 40,000 MPs/mL, respectively (Figure

3.8). FLU body burdens were dose-dependent across all FLU- MP treatments. As

mentioned above, 11.2, 22.2, 59.0, and 138.6 µg FLU/L desorbed from the exposures of

20,000, 30,000, 40,000, and 80,000 FLU-saturated MPs/mL, respectively (Figure 3.8).

Seven-day exposures to the water bioavailable fraction of FLU also resulted in a dose-

dependent FLU body burden. FLU body burdens of 0.06, 0.10, and 0.13 µg FLU/

organism resulted from exposures to 11.2, 22.2, and 59.0 µg FLU/L (which represent the

amount of FLU that desorbed from FLU-saturated MP treatments of 20,000, 30,000, and

40,000 MPs/mL, respectively). However, there was no significant difference between the

FLU body burdens of copepods exposed to different concentrations of water bioavailable

FLU. Seven-day exposures to the water bioavailable fraction of FLU also included two

additional FLU exposure treatments; FLU body burden was only significantly greater

than the control (0 µg FLU/L) at the highest FLU exposure (118.08 µg FLU/L) (Figure

3.9). Interestingly, when comparing the fluoranthene body burdens of copepods and

amphipods exposed to 20,000 FLU-MPs/mL (normalized to dry weight), the body

burdens are not significantly different (Figure 3.10).

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3.4.3 Freshwater fish Pimephales promelas

Following a 96-hr exposure to fluoranthene-contaminated polyethylene MPs

(FLU-MPs), the results demonstrated that the ingestion of FLU-MPs increases the

bioavailability of fluoranthene to exposed fathead minnows. Exposures to 100,000 FLU-

MPs/mL resulted in fluoranthene body burdens of approximately 540 µg FLU per

organism (Figure 3.11). Approximately 162.5 µg FLU/L desorbed from the FLU-

saturated MPs, thus a 96-hr exposure to a range of waterborne fluoranthene was also

conducted (0- 162 µg FLU/L) in order to differentiate the fluoranthene body burden that

resulted from microplastic ingestion, from that of external fluoranthene exposure.

Exposures to water-borne fluoranthene resulted in a dose-dependent fluoranthene body

burden, with body burdens of 96.4, and 368.5 µg FLU/organism, corresponding with 96-

hr exposures to 40.6 and 162.5 µg FLU/L, respectively. Unfortunately, the replicates

where fathead minnows were exposed to 81.3 µg FLU/L were unable to be analyzed due

to mishandling of the beakers during the experiment, resulting in organism death prior to

the end of the 96-hr exposure. After comparing the fathead minnow body burdens

resulting from exposures to 100,000 FLU-saturated MPs/mL and 162.5 µg FLU/L (the

corresponding amount of fluoranthene that desorbed from 100,000 FLU-saturated

MPs/mL), the results showed that 70% of the fluoranthene body burden from exposures

to 100,000 FLU-saturated MPs/mL can be attributed to the external water-borne

bioavailable fluoranthene. No mortality was observed during exposures to water-borne

fluoranthene, or fluoranthene-contaminated microplastics (data not shown).

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3.4.4 Marine fish Fundulus heteroclitus

Mummichogs exposed to 100,000 fluoranthene-contaminated microplastics/mL

(FLU-MPs) for 96 hours did not result in a significantly different fluoranthene body

burden than that of mummichogs exposed to 200 µg fluoranthene/L (the amount of

fluoranthene that desorbed from 100,000 FLU-MPs/mL) (Figure 3.12). There was also

no mortality observed in either 96-hr bioassay to FLU-MPs or water-borne fluoranthene.

3.5 Discussion

Plastic debris can concentrate organic contaminants up to 500 times the

concentration of contaminants in the water column (Wurl and Obbard 2004); although

plastic itself may be biochemically inert, it is capable of enhancing absorption of soluble

contaminants to various tissues. The influence of MPs on the bioavailability of

contaminants in aquatic environments has not been widely investigated, however the

exposure to PAHs and MPs has been shown to increase the bioavailability of PAHs to

exposed invertebrates and fish (Paul-Pont et al. 2016; Oliveira et al. 2013). Thus far,

studies have demonstrated the capacity of MPs to sorb both metals and organic

contaminants, and act as a source of contaminants to aquatic organisms (Voparil and

Mayer 2000; Voparil et al. 2004; Bakir et al. 2012; Bakir et al. 2014; Khan et al. 2015).

Amphipods and copepods exposed to FLU-MPs in my study had a dose-dependent

fluoranthene body burden, where over 90% of the fluoranthene body burden was due to

microplastic ingestion and not from the fluoranthene that desorbed from the MPs into the

exposure water. However, almost the opposite was observed for fathead minnows, where

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only about 30% of the fluoranthene body burden was due to FLU-MP ingestion. One

other study has investigated whether polystyrene microplastics (2-6 µm) influenced the

bioavailability of fluoranthene to exposed mussels Mytilus spp. (Paul-Pont et al. 2016).

The results suggested that microplastic exposure may increase fluoranthene

bioavailability by inhibiting detoxication pathways; mussels exposed to polystyrene

microplastics and fluoranthene had higher fluoranthene body burdens following the 7-d

depuration period in comparison to those only exposed to fluoranthene (no

microplastics). In my study, amphipods chronically exposed to fluoranthene were able to

quickly excrete ingested fluoranthene, as there was no significant difference in

fluoranthene body burdens between the control (no fluoranthene) and any of the

fluoranthene exposures, following the 14-d depuration period. No chronic FLU-MP

exposure was conducted due to limited MP supply; however, it would be interesting to

investigate whether my results would have differed if the polyethylene MPs used would

have had a similar effect on fluoranthene detoxication pathways in exposed amphipods,

as seen in exposed mussels exposed to polystryene and fluoranthene (Paul-Pont et al.

2016). After the 14-d depuration period, mortality also significantly increased in

fluoranthene exposure treatments (5, 10, and 36 µg FLU/L); the data suggested that a

chronic exposure to concentrations of fluoranthene associated with contaminated

microplastics may ultimately lead to decreased survival, despite maintaining the ability to

excrete fluoranthene.

Although no studies have used marine copepods, studies have investigated the

bioavailability of contaminants in aquatic ecosystems to the benthic marine lugworm,

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Arenicola marina, where lugworms exposed to sand with 5% contaminated polyvinyl

chloride microplastics resulted in the uptake of nonylphenol, phenanthrene,

polybrominated diethyl ether-47, and triclosan (Browne et al. 2013). However, in

comparison to exposures to contaminated sand particles alone, there was a greater

bioavailability of PBDE-47 and triclosan when lugworms were exposed to contaminated

microplastics (Browne et al. 2013). It should be noted that in comparison to natural

organic matter and microplastics, sand is a poor sorbent for organic contaminants due to

differences in available surface area, surface charge, and generally less potential for

binding hydrophobic compounds. When mussels, Mytilus galloprovincialis, were

exposed to pyrene-contaminated polyethylene and polystyrene MPs, the bioaccumulation

of pyrene was observed, along with additional negative cellular effects including DNA

damage and a reduction in various enzymes related to antioxidant and immune responses

(Avio et al. 2015). Interestingly, despite a greater concentration of pyrene adsorbed to

the polyethylene MPs, there was no significant difference between the concentration of

pyrene found in the gills and digestive glands due to polymer type (Avio et al. 2015).

Contaminant bioavailability may also be influenced by MP characteristics such as

shape, size and polymer type. Various studies have shown that fibrous MPs are

predominantly found in the environment over other MP shapes (i.e. fragments, pellets,

shavings, beads) (Graham and Thompson 2009; Nor and Obbard 2014). Further, the

analysis of gut contents in field-collected marine decapods, Nephrops norvegicus,

demonstrated the formation of ball-shaped aggregates, comprised of fibrous MPs, in 50%

of the organisms. Interestingly, intermolt phase and organism size seemed to influence

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the type and quantity of MPs found in the gut, respectively, where organisms with a

softer carapace, and those larger in size had fewer plastic balls in the guts (Murray and

Cowie 2011). The results from Murray and Cowie (2011) suggest that the formation of

MP aggregates may be age-specific, and if it is difficult for exposed organisms to egest

MPs via excretion, molting may be one of the only ways in which certain invertebrates

are able to eliminate ingested MPs (Murray and Cowie 2011). Longer gut residence

times of contaminated MP fibers between molting periods may result in greater

desorption of adsorbed contaminants. However, MP shape effects may also be species-

specific, as Hamer et al. (2014) demonstrated that there was no significant effect on MP

ingestion based on the MP shapes tested (microbeads, fragments, fibers) on exposed

marine isopods, Idotea emarginata. MP size may also influence contaminant

bioavailability, where 3.0 µm spherical polystyrene MPs ingested by mussels were able

to translocate from the gut to the hemolymph in greater concentrations as compared to

9.6 µm polystyrene MPs (Browne et al. 2008). Few studies have shown the translocation

of MPs from the digestive tract into other internal organs, but Browne et al. (2008)

demonstrated that polystyrene MPs in both size classes remained in the hemolymph for

48 days, a residence time that far surpassed the 3-hr exposure.

Not as much research is available on whether MPs act as a vector for

environmental contaminants to fish or other higher trophic level organisms. A few

studies have shown MPs act as vectors for both POPs and silver to marine and freshwater

fish (Oliveira et al. 2013; Rochman et al. 2014a; Rochman et al. 2014b; Khan et al.

2015). However, the uptake of polychlorinated biphenyls, DDT, and polybrominated

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diethylether was not attributed to the amount of these contaminants extracted from

microplastics found in the stomachs of northern fulmars (Fulmarus glacialis); POP

ingestion was largely thought to result from the ingestion of POPs through prey

consumption by fulmars (Herzhe et al. 2016). In my work, MPs did increase the

bioavailability of FLU to exposed juvenile fathead minnows, while the same was not

observed in mummichogs, despite mummichogs having a FLU body burden

approximately 4x higher than that of fathead minnows. No mortality was observed in

exposures to FLU-MPs or water-borne FLU for bioassays using fathead minnows and

mummichogs, despite the relatively high concentrations of FLU (162.5 and 200 µg

FLU/L, respectively). Other studies have demonstrated that median effect and lethal

concentrations for freshwater species are 7.7 and 187 µg FLU/L, respectively, and 400

and 500 µg FLU/L for marine species (Horne and Oblad 1983; Gendusa 1990; Suedel et

al. 1993; US EPA 1978; Rossi and Neff 1978). Further, the higher limits of FLU toxicity

are very close or over the solubility of FLU in fresh- and saltwater, 206 and 127 µg

FLU/L, respectively (May 1980).

My experiments, similar to most laboratory tests, were conducted under

fluorescent light, although it has been demonstrated that ultraviolet (UV) light increases

the toxicity of fluoranthene to freshwater and saltwater organisms (Ankley et al. 1995;

Boese et al. 1997; Pelletier et al. 1997). Exposure to fluoranthene is generally associated

with phototoxicity due to the production of reactive oxygenated species; sensitivity to

fluoranthene in the presence of UV radiation decreases with age (Finch and Subblefield

2015; Weinstein et al. 1997). It is possible that little mortality was observed in my results

158

(except in the 7-d 80,000 FLU-MPs/mL exposure to copepods) due to the lack of reactive

oxygenated species during the exposure. A 96-hr exposure to 44 and 212 µg FLU/L

under fluorescent lighting, resulted in 50% mortality in exposed H. azteca and P.

promelas, both concentrations greater than those used in my fluoranthene exposures for

amphipods and fathead minnows (Spehar et al. 1999). Fathead minnows exposed to 12.2

µg FLU/L fluoranthene in the presence of UV light for 96 hours experienced 50%

mortality (Spehar et al. 1999). In the environment, it is likely that exposure to MPs and

fluoranthene, as well as other phototoxic contaminants, may result in greater toxicity

compared to my laboratory results, as juvenile fish, invertebrates, and embryos (typically

laid in shallow areas) are susceptible to high exposures to UV light due to their

translucent bodies, even after MPs are ingested. Further, the aggregation of MPs on the

gills of exposed organisms in the presence of UV light and phototoxic contaminants may

result in further oxidative damage to gill tissue, thereby reducing respiratory capabilities

(Weinstein et al. 1997; Watts et al. 2014; Farrell and Nelson 2013).

Although total body burden analysis was the only method used to quantify

fluoranthene bioaccumulation in my bioassays, other studies have demonstrated that

higher concentrations of contaminants partition into the gut lining, as opposed to other

tissues that may come into contact with contaminated microplastics (i.e. body wall, and

gills; Browne et al. 2013). Uptake routes of contaminants via the presence of

microplastics in the environment have not been well documented; however, results to

date suggest microplastic ingestion as compared to water-borne only exposure leads to

greater bioaccumulation, and demonstrates the complexity in quantifying the transport

159

and fate of contaminants associated with microplastics. The presence of microplastics in

aquatic ecosystems can act as a greater source of organic contaminants than the water

bioavailable fraction; however, polymer characteristics (i.e. shape, size, polymer type,

surface characteristics) and water chemistry will also largely influence the sorption of

contaminants onto microplastic surfaces. Although not widely investigated in regards to

contaminant interactions with microplastics, the influence of the presence of organic

matter and other sorbents in aquatic ecosystems is indisputably important to

understanding the dynamics of microplastic presence and exposures in the environment.

3.6 Limitations

One of the main limitations to the bioassays conducted in Chapter Three is the

saturation of binding sites on the surface of polyethylene MPs with fluoranthene in order

to determine whether adsorbed fluoranthene is bioavailable to exposed organisms upon

the MPs being ingested. Although saturation the MPs with fluoranthene was necessary to

ensure sufficient fluoranthene would be present on the MPs during the exposure, in terms

of environmental relevance, it is unlikely that any MPs would only have one type of

contaminant adsorbed to their surfaces or that the MPs would be at saturation. The

binding affinity of different contaminants to different MPs will largely determine whether

a given contaminant will be bioavailable to an exposed organism. It is also likely that

greater sorption of contaminants onto MPs in the environment will increase as MPs age

and fragment, developing crevices in the surface and increasing the overall available

surface area for contaminant sorption. Compared to imaging of fragmented MPs

160

purchased from Polygroup Inc. (data not shown), the binding of fluoranthene onto the

spherical MPs purchased from Cospherix Inc. (the MPs used throughout my dissertation)

was not as uniformly distributed on the MP surface. It is likely that the bioavailability of

fluoranthene due to exposure to the primary polyethylene MPs used in this study will

differ greatly from that which may result from exposure to other types of MPs. Finally

another limitation to this study was the inability to renew the exposures to both

fluoranthene and fluoranthene-contaminated MPs more than once every 24 hours during

the exposures. Fluoranthene is volatile, and the amount of fluoranthene quantified in the

exposure water during the fluoranthene-contaminated MP and waterborne fluoranthene

exposures likely underestimated the amount of fluoranthene actually present in the

exposure water.

3.7 Conclusions

The MPs used in this study were able to become completely saturated with

fluoranthene within 48 hours, which suggests that the release of MPs into the

environment is likely creating a different sink for environmental contaminants.

Furthermore, the results from Chapter Three demonstrates that contaminants like

fluoranthene that are present on MPs found in the environment are likely to be

bioavailable to exposed organisms upon being ingested. Specifically, with both the

amphipods and copepods, the fluoranthene associated with MP ingestion resulted in over

95% of the fluoranthene body burden in all exposure treatments. On the other hand,

fluoranthene associated with MP ingestion contributed to less than 5% of the resulting

161

body burden. The toxicity associated with MPs and the adsorbed contaminants is likely

to be species-specific.

162

3.8 Figures

Figure 3.1: Adsorption isotherm of fluoranthene onto polyethylene microplastics Adsorption isotherm of fluoranthene onto fluorescent polyethylene microplastic spherical

particles suspended in 30 ppt artificial sea water (blue; SD = ± 1.27), and hard

reconstituted freshwater (orange; SD = ± 0.78); n=3 for both isotherms.

0

5

10

15

20

25

30

35

0 10 20 30 40 50

qe (µ

g/m

g)

Ce (µg FLU/L)

30 ppt HRW

163

Figure 3.2: Images taken of polyethylene microplastics with and without

fluoranthene

Images taken with a Leica SP8X Multiphoton spectral confocal microscope of

fluorescent polyethylene microplastic spherical particles A) suspended in 30 ppt artificial

sea water and B) suspended in 50 µg fluoranthene/L-spiked 30 ppt artificial sea water for

48 hours. The white arrow represents the presence of fluorescent polyethylene

microplastics spherical particles (A and B), and the orange arrow represents the presence

of adsorbed fluoranthene on the surface of fluorescent polyethylene microplastic

spherical particles (B).

A B

164

Figure 3.3: Mortality following 42-d chronic exposure to fluoranthene and 14-d

depuration period

Mean percent mortality for Hyalella azteca exposed to fluoranthene at day 10 (SD = ±

0.16), and day 42 (SD = ±1.44) during the 42-d exposure to fluoranthene. Day 56

(recovery) represents the percent mortality resulting from exposure to clean water (no

FLU) during the 14-d depuration period (day 56; SD = ±1.51). The asterisk represents

statistical differences (p < 0.05) in percent mortality between the different time points

within a treatment group.

-10

0

10

20

30

40

0 3.5 5 10 18 36

Perc

ent m

orta

lity

µg FLU/L

Day 10 Day 42 Day 56 (recovery)

* *

* *

*

165

Figure 3.4: Fluoranthene body burden following 10-d acute exposure to

fluoranthene-contaminated microplastics and water-borne fluoranthene

Mean fluoranthene body burden of exposed Hyalella azteca (µg FLU/organism) during

an acute 10-d exposure to fluoranthene-contaminated polyethylene microplastic spherical

particles (5,000, 10,000 and 20,000 fluoranthene-contaminated polyethylene

microplastics/mL) (SD = ± 63.09) and fluoranthene (3.5, 10 and 18 µg FLU/L) (SD = ±

2.17), normalized to dry weight. The upper-case letters represent statistical differences (p

< 0.0001) in fluoranthene body burden between fluoranthene water-borne exposure

treatments, and the lower-case letters represent statistical differences (p<0.0001) in

0

50

100

150

200

250

300

350

400

450

3.5 µg FLU/L 5,000 MPs/mL 10 µg FLU/L 10,000MPs/mL

18 µg FLU/L 20,000MPs/mL

Fluo

rant

hene

bod

y bu

rden

(µg

FLU

) per

mg

dry

wei

ght

Treatment: Exposure to aqueous fluoranthene (µg FLU/L) and fluoranthene-saturated microplastics (MPs/mL)

B

AA

BAA

MP FLU exposure

Water-only FLU exposure

aab

b

166

fluoranthene body burden between fluoranthene-contaminated (MPs/mL) microplastic

exposure treatments.

167

Figure 3.5: Fluoranthene body burden following 10-d acute exposure to water-borne

fluoranthene

Mean fluoranthene body burden of Hyalella azteca (µg FLU/organism) (SD = ± 3.60),

normalized to dry weight, during an acute 10-d exposure to fluoranthene (0-36 µg

FLU/L). The letters represent statistical differences (p < 0.0001) in fluoranthene body

burden between fluoranthene exposure treatments.

0

20

40

60

80

100

120

0 3.5 5 10 18 36

Fluo

rant

hene

bod

y bu

rden

(µg

FLU

) per

m

g dr

y w

eigh

t

Fluoranthene exposure treatment (µg FLU/L)

A A AA

B

C

168

Figure 3.6: Fluoranthene body burden following 42-d chronic exposure to

fluoranthene and 14-d depuration period

Mean fluoranthene body burden of Hyalella azteca (µg FLU/organism) (SD = ± 3.60

(day 10); SD = ± 13.07 (day 42); SD = ± 3.44 (day 52) ) , normalized to dry weight,

during a chronic 42-d exposure to fluoranthene (3.5, 10 and 18 µg FLU/L), and 14-d

depuration period (day 56). The letters represent statistical differences (p < 0.0001) in

fluoranthene body burden between fluoranthene exposure treatments evaluated on day

10. The asterisk represents statistical differences (p < 0.001) in fluoranthene body burden

between fluoranthene exposure treatments evaluated on day 42. There was no significant

difference between fluoranthene body burdens between fluoranthene exposure treatments

evaluated on day 52 (depuration).

0

20

40

60

80

100

120

140

160

0 3.5 5 10 18 36

Aver

age

FLU

bod

y bu

rden

(µg

FLU

) per

m

g dr

y w

eigh

t

Fluoranthene exposure concentration (µg FLU/L

Day 10 Day 42 Day 56 (recovery)

*

*

B

C

AAAA

169

Figure 3.7: Reproduction following 42-d chronic exposure to fluoranthene and 14-d

depuration period

Mean reproduction (number of neonates per female) (SD = ± 2.39) of exposed Hyalella

azteca, in response to exposure to water-borne fluoranthene (0-36 µg/L) at days 42 and

56. The asterisk represents statistical differences in mean reproduction (p < 0.05)

between fluoranthene exposure concentrations within a time point.

0

5

10

15

20

25

30

0 3.5 5 10 18 36

Neo

nate

s/fe

mal

e

Fluoranthene exposure concentration (µg FLU/L)

Day 42 Day 56 (recovery)

* *

170



Mean fluoranthene body burden of exposed Amphiascus tenuiremis (SD = ± 0.26) during

the 7-d acute exposure to water-only available fraction of fluoranthene (no microplastics),

and fluoranthene-saturated polyethylene microplastics. Different letters represent

statistical differences (p < 0.05) in fluoranthene body burden per organism between

treatments of fluoranthene-saturated polyethylene microplastics. There is no statistical

difference (p > 0.05) in fluoranthene body burden per organism between treatments of

water-only available fractions of fluoranthene (no microplastics).

0

0.5

1

1.5

2

2.5

3

3.5

4

4.5

11.16 µgFLU/L

20,000MPs/mL

22.2 µg FLU/L 30,000MPs/mL

59.04 µgFLU/L

40,000MPs/mL

Fluo

rant

hene

bod

y bu

rden

(µg

FLU

) pe

r org

anism

Treatment: Exposure to aqueous fluoranthene (µg FLU/L) and fluoranthene-saturated microplastics (MPs/mL)

B

C

A

MP FLU exposure

Water-only FLU

171

Figure 3.9: Fluoranthene body burden following 7-d acute exposure to water-borne

fluoranthene

Mean fluoranthene body burden of exposed Amphiascus tenuiremis (SD = ± 0.10) during

the 7-d acute exposure to water-only available fraction of fluoranthene (no microplastics).

Asterisks represent statistical differences (p < 0.05) in fluoranthene body burden per

organism between treatments of water-only available fractions of fluoranthene (no

microplastics).

0

0.2

0.4

0.6

0.8

1

1.2

1.4

0 5.58 11.16 22.2 59.04 118.08

Fluo

rant

hene

bod

y bu

rden

(µg

/L)

per o

rgan

ism

Fluoranthene exposure concentration (µg FLU/L)

*

172

Figure 3.10: Comparison of fluoranthene body burden between copepods and

amphipods

Mean fluoranthene body burden of exposed Amphiascus tenuiremis and Hyalella azteca

(SD= ± 121.84) during the 7-d copepod, and 10-d amphipod acute explores to 20,000

fluoranthene-saturated polyethylene microplastics/mL. There is no significant difference

between the fluoranthene body burdens between invertebrates exposed to the same

number of fluoranthene-saturated polyethylene microplastics.

0

100

200

300

400

500

600

Copepod AmphipodFluo

rant

hene

bod

y bu

rden

(µg

FLU

) per

m

g dr

y w

eigh

t

Exposure to 20,000 FLU-contaminated MPs/mL

173



Mean fluoranthene body burden per organism after a 4-d Pimephales promelas exposure

to 100,000 fluoranthene saturated microplastics/mL (orange) (SD= ± 99.04), and water

bioavailable fluoranthene (blue) (SD= ± 12.29), with no microplastics present. Different

letters represent statistical differences (p < 0.0001) between each exposure treatment.

0

100

200

300

400

500

600

700

0 40.625 162.5 162.5 + MPs

Fluo

rant

hene

bod

y bu

rden

(µ

g FL

U/o

rgan

ism)

Treatment: Fluoranthene water bioavailable fraction (ug FLU/L); Microplastic exposure concentration (MPs/mL)

Water bioavailable FLU FLU saturated MPs

A

B

C

D

174



Mean fluoranthene body burden per organism after a 4-d Fundulus heteroclitus exposure

to 100,000 fluoranthene saturated microplastics/mL, and 200 µg FLU/L water

bioavailable fluoranthene (SD= ±305.33). There is no significant difference in

fluoranthene body burden (p > 0.05) between organisms exposed to microplastics with

and without fluoranthene.

0

500

1000

1500

2000

2500

3000

200 200 + 100 000 MPs/mL

Fluo

rant

hene

bod

y bu

rden

(u

g FL

U/o

rgan

ism)

Fluoranthene water bioavailable fraction (µg FLU/L); Microplastic exposure concentration (MPs/mL)

175

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https://www.epa.gov/sites/production/files/2015-09/documents/priority-pollutant-list-epa.pdf

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CHAPTER FOUR

MICROPLASTIC TROPHIC TRANSFER

4.1 Abstract

To evaluate the process of trophic transfer of microplastics, it is important to

consider various abiotic and biotic factors involved in their ingestion, egestion,

bioaccumulation, and biomagnification. Current research regarding microplastic presence

in the environment suggests that regardless of feeding mechanism or habitat, all

organisms in aquatic habitats are susceptible to microplastic exposure. Limited field

evidence from higher trophic level organisms in a variety of habitats suggests that trophic

transfer of microplastics may be a common phenomenon and occurs concurrently with

direct ingestion. Thus far in situ results have demonstrated that microplastics imbedded in

food materials can be ingested by crabs, and that microplastics can undergo trophic

transfer within a zooplankton food web. However, current uncertainties regarding actual

environmental microplastic concentrations and different reporting methods for

microplastic quantification and characterization processes make it difficult to ascertain

whether the trophic transfer of microplastics is a significant route in which multiple

levels of biological organization may be exposed.

To investigate microplastic trophic transfer between freshwater and marine

invertebrates to fish, Pimephales promelas and Fundulus heteroclitus were fed Hyalella

azteca and Amphiascus tenuiremis that had been previously exposed to microplastics

with and without fluoranthene (FLU), respectively. In the previous chapter, my results

indicated that the FLU adsorbed to microplastics was significantly more bioavailable to

181

exposed fathead minnows, but not necessarily mummichogs. However, fluoranthene was

bioavailable to fathead minnows and mummichogs that were indirectly exposed to both

FLU and microplastics through the consumption of prey (i.e. amphipods and copepods,

respectively). Another experiment was conducted to evaluate whether microplastics and

FLU could undergo trophic transfer between Pimephales promelas and Morone chrysops

x Morone saxatilis. Hybrid striped bass that were fed fathead minnows that had

previously ingested microplastics with and without FLU showed the presence of FLU in

the bile, suggesting that microplastics may act as a vector for organic contaminants in

aquatic food chains between fish species as well. Microplastics indirectly ingested

through prey will likely serve as a vector for organic contaminants, and may contribute to

the bioaccumulation and biomagnification of contaminants in the environment.

4.2. Introduction

Limited research is available regarding the trophic transfer of microplastics,

however, the discovery of microplastics in the scat and tissues of organisms that represent

a diverse array of habitats suggest the strong potential for trophic transfer occurring

concurrently with the direct ingestion of microplastics (Fry et al.1987; Eriksson and

Burton 2003; Goldsworthy et al. 1997; Lusher et al. 2012). Microplastics are in the size

range of many plankton species and other types of particles that are normally ingested by

aquatic invertebrates, thus posing the potential for microplastics and associated

contaminants will undergo trophic transfer to higher biologically classified organisms

(Desforges et al. 2015; Browne et al. 2008). Invertebrates and fish with different feeding

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mechanisms and habitats have also been observed to easily consume microplastics in vivo

and in vitro (Browne et al. 2008; Thompson et al. 2004; Graham and Thompson 2009;

Setälä et al. 2014; Jabeen et al. 2017).

Approximately 80% of plastic debris is attributed to land-based sources (i.e.

industrial discharges, garbage management, litter), thus it is likely that terrestrial

organisms will also be exposed to microplastics due to the application of microplastic-

containing sludge, and the potential release of microplastic fibers from concrete cracking

(Andrady 2011; Rillig 2012; Zubris and Richards 2005; Pelisser et al. 2010).

Sandhoppers exposed to polyethylene spherical microplastics (10-45 µm) for 24 hours

were able to excrete all microplastics within 24 hours after the initial exposure (Ugolini et

al. 2013). Categorizing sandhoppers as strictly a terrestrial organism would be inaccurate

as they dwell below the wet sand on beach coastlines, making them susceptible to plastic

debris of marine as well as terrestrial origin, yet no other studies have investigated the

toxicity of microplastics to terrestrial organisms. The incorporation of surface

microplastics into the burrows of the common earthworm, Lumbricus terrestris,

demonstrates that there may be indirect toxicological effects (i.e. altering microplastic

bioavailability to sediment organisms) associated with microplastic exposures (Lwanga et

al. 2017). Organisms that inhabit such transient and dynamic ecosystems throughout their

lifetime or at different developmental stages will be exposed to microplastics that are

specific to certain environmental conditions, thus making it imperative that additional

research be conducted to best determine how terrestrial and aquatic exposures to

microplastics should be studied.

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Microplastics have been found in all aquatic ecosystems sampled, but it is

premature to state definitely whether the sediment or surface water will be the larger sink

of microplastics due to the different methods in which microplastics are collected and

quantified (Claessens et al. 2011; Law et al. 2010; Moore et al. 2001; Thompson et al.

2004). However, due to upwards of 300 thousand microplastics/m3 found in urbanized

rivers, and larger quantities of microplastics found in fish from urban rivers, it is very

likely that organisms that either live in, or frequent urbanized coastal areas are at great

risk for microplastic exposures (McCormick et al. 2016; Dris et al. 2015; Sanchez et al.

2014). Therefore, coastal food webs may be more susceptible to the trophic transfer than

those in other habitats. It is also likely that remote regions will have microplastic trophic

transfer; it is likely that the low density and slow degradability of plastic in general

contribute to the fast transportation of plastic to areas where plastic use is low (Free et al.

2014).

Zooplankton plays a large role in aquatic food webs, and the presence of

microplastics and zooplankton in both the sediment and water column makes it inevitable

that microplastics will be consumed by both zooplankton and potential predators. The

trophic transfer of polystyrene microplastics to macrozooplankton occurred after only

three hours of exposure to mesozooplankton that had previously ingested polystyrene

microplastics (Setälä et al. 2014). Although the amount of microplastics ingested per

zooplankton individual may not be high in the environment (less than one particle per

organism examined), the continuous consumption of high numbers of zooplankton by

predators like humpback whales and juvenile fish make it likely for microplastics to

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bioaccumulate in larger fish and mammals (Desforges et al. 2015). The presence of

microplastics and plastic additives in whale feeding grounds also makes it likely that

whales ingest high concentrations of microplastics and persistent organic pollutants and

contaminated prey (Fossi et al. 2016). The ingestion of microplastics by organisms

throughout the water column in addition to the benthos indicates the ease at which

microplastics may be transferred throughout the food web and water column (Setälä et al.

2014).

The residence time of microplastics in the tissues and gut of biota are a function

of several factors. Microplastics were found in the hemolymph of crabs (Carcinus

maenas) 21 days after being provided with mussels (Mytilus edulis) that had consumed

polystyrene microplastics (Farrell and Nelson 2013). The residence time of the

microplastics found within the crabs varied among different organs (i.e. digestive tract,

hepatopancreas, ovary, gills), but were less than that of the hemolymph (Farrell and

Nelson 2013). The translocation of polystyrene microplastics, <10 and 0.5µm, into the

hemolymph of invertebrates for up to 48 and 21 days, respectively, suggests that perhaps

the longer residence time of microplastics due to translocation may depend on polymer

composition (Farrell and Nelson 2013; Browne et al. 2008). Body residence time may

also be influenced by uptake routes, where it was observed that the ventilation of

microplastics through gills resulted in a longer microplastic residence time, than that

from microplastic ingestion, when crabs were exposed to 8-10 µm polystyrene

microplastics (Watts et al. 2014). Microplastic shape is another characteristic that may

influence body residence time, as the ingestion of fiber-shaped microplastics by the crab,

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C. maenas, and lobster, Nephrops, resulted in the formation of an microplastic aggregate

within the gut and a reduction in microplastic egestion times (Murray and Cowie 2011;

Watts et al. 2015). Longer body residence times of microplastics will increase the chance

that microplastics within a prey can be consumed by a higher level organism, thus

providing another route of exposure to microplastics in the environment. Whether this

contribution is sufficient to be toxic at environmentally relevant concentrations is

uncertain as there is a high variability in microplastic consumption by exposed organisms

(Farrell and Nelson 2013; Au et al. 2015).

Numerous chemicals such as plasticizers and the antibacterial triclosan are often

incorporated into plastic products during their production (i.e. additives) and can also

adsorb onto plastics once released into the environment (i.e. persistent organic

pollutants). Concerns have been voiced about the trophic transfer of chemicals

associated with microplastics, and the toxicological ramifications of microplastics acting

as a source and vector for a mixture of contaminants. It is also likely that the uptake of

microplastic-associated contaminants will be greater for higher trophic organisms as

desorption rates of persistent organic contaminants from microplastics were faster in

treatments that simulated gut conditions of warm blooded organisms (i.e. higher body

temperature, lower pH, and the presence of gut surfactants) (Bakir et al. 2014). Only one

published study thus far has demonstrated the trophic transfer of polyethylene

microplastics and adsorbed benzo[a]pyrene from Artemia sp. to zebrafish, where the

desorption of benzo[a]pyrene was observed in the intestinal epithelium and resulted in an

induction of cytochrome P450 1A (CYP1A) in the liver of the exposed zebrafish (Batel et

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al. 2016). Plastic ingestion has also been related to trace metals and PBDEs present in

sea bird feathers and abdominal adipose tissue, respectively (Lavers and Bond 2016;

Tanaka et al. 2013), demonstrating how plastic ingestion can contribute to toxicant

uptake in larger marine organisms. However, when persistent organic pollutant (POP)

uptake from plastic ingestion by fulmars was compared to that from the ingestion of prey,

it was found that the bioaccumulation of POPs due to plastic ingestion is negligible in

comparison to other POP uptake routes (Herzke et al. 2016). The limited evidence to date

indicates that it is likely that microplastics may act as a source of contaminants to

exposed organisms through their direct and indirect (trophic transfer) consumption;

however, the studies mentioned have exposed contaminated microplastics to clean test

organisms (where the transfer of contaminants to the organism is favored) (Koelmans

2015). In environments where contaminants are already present, microplastic ingestion

may act as a sorbent for contaminants already present within the exposed organism,

resulting in an overall depuration effect (Koelmans 2015). Rainbow trout first exposed to

polychlorinated biphenyls (PCB) and then polyethylene microplastics through their feed

for up to 9 weeks after the initial PCB exposure demonstrated that there was no

significant effect on PCB elimination rates, indicating that in this experiment, the

ingestion of uncontaminated MPs did not result in an expedited depuration of PCBs from

exposed fish (Rummel et al. 2016). Further investigations on whether microplastics will

act as a source of or depuration method for environmental contaminants are needed.

The presence of microplastics in the wild as well as areas designed for

aquaculture indicates the potential for microplastics to be present in organisms commonly

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ingested by humans (Davidson and Dudas 2016; Castro et al. 2016; De Witte et al. 2014).

Microplastics found in both wild and cultured Venerupis philippinarum, Manila clams

(0.07-5.47 particles/g), in British Columbia indicates that microplastics are in seafood

consumed by humans (Davidson and Dudas 2016). Microplastic fibers between 200 and

1500 µm were detected in blue mussels, Mytilus edulis, purchased from a store,

demonstrating that microplastics ingested by higher trophic organisms through trophic

transfer will also be specific to certain microplastic characteristics, as no other

microplastic shapes were found in mussels sampled (De Witte et al. 2014). Microplastics

were found in higher concentrations in supermarket-purchased clam, Crassostrea gigas,

in comparison to mussel M. edulis, attained from a farm further showing microplastics

are present in human consumption-ready shellfish (Van Cauwenberghe and Janssen

2014).

There are no published studies thus far that have investigated the trophic transfer

of microplastics between invertebrates and fish, and smaller prey fish to a larger

predatory fish. Fathead minnows, mummichogs, and hybrid striped bass were trained to

consume amphipods, copepods, and fathead minnows, respectively, that have been

previously exposed to microplastics with and without adsorbed fluoranthene to

investigate whether the indirect ingestion of microplastics will influence the

bioavailability of fluoranthene and microplastics to a predator organism. In my study the

use of five organisms commonly found in both freshwater and marine ecosystems in

trophic transfer bioassays, using the same contaminants and methods, provides results

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that will contribute to cross-species comparisons in terms of microplastic toxicity and

trophic transfer.

4.3 Methods

4.3.1 Microplastic exposure preparations

The same polyethylene spherical microplastics purchased from Cospherix used in

previous studies (see Chapters Two and Three) were also used in the trophic transfer

bioassays, where the microplastics were saturated with fluoranthene using methods

described in 3.3.1. Briefly, polyethylene microplastics were placed in amber vials

containing 800 µg fluoranthene/L for 48 hours under constant rotation (30 rpm).

Concentrations of polyethylene microplastics prepared for each specific prey organism,

and the number of prey organisms fed per predator organism are described in Table 4.1.

High concentrations of fluoranthene were used to ensure saturation of fluoranthene on

microplastics. After 48 hours, the microplastics were vacuum filtered through 0.22 µm

nylon filters and rinsed with clean water three times prior to being re-suspended in clean

reconstituted hard water (RHW), 30 ppt artificially prepared sea water, and moderately

hard water (MHW) for amphipod, copepod, and fathead minnow exposures, respectively.

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Table 4.1: Microplastic exposure concentrations (#MPs/mL) and number of prey organisms fed to prey and predator organisms, respectively.

Microplastic (MP) Exposure to Prey

Organism: #MPs/mL

Prey Organism Number of Prey Organisms fed per Predator Organism

Predator Organism

10,000 Hyalella azteca 20 Pimephales promelas

40,000 Amphiascus tenuiremis

20 Fundulus heteroclitus

100,000 Pimephales promelas

15 Morone chrysops x Morone saxatilis

4.3.2 Microplastic trophic transfer from Hyalella azteca to Pimephales promelas

As described in Section 2.3.1, known-age juvenile amphipods for the 3-d feeding

bioassay were attained. Briefly, mature amphipods were placed in 1.0-L glass mason jars

for one week; and less than 7-d old neonates were placed in an aerated holding chamber

prior to being used during the fathead minnow trainings and feeding bioassays.

Group and Individual Training Sessions

Three-month old fathead minnows, Pimephales promelas, were trained to

consume ≤ 7-day old amphipods over the course of nine days. During both the group and

individual training sessions, fathead minnows were fed juvenile amphipods that were not

fed microplastics with or without fluoranthene. Tanks used for both group and individual

training sessions were aerated, kept at 25 ± 1°C, and the USEPA moderately hard water

used for fathead minnow culture and experiments were statically renewed daily (USEPA

1989).

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Three fathead minnows were placed within each 5-L tank for the group training

sessions. The fish were stocked so as to not exceed 1.0 g wet weight biomass/ L of water;

and dissolved oxygen, and ammonia concentrations were monitored during the training

and exposure to ensure levels did not drop below 5 mg/L or exceed 0.5 mg/L,

respectively. Food was withheld from all fathead minnows for the first 48 hours, to

provide incentive for the fish to consume juvenile amphipods. During the four days of

group training, each tank was given approximately 50 amphipods per day, and the

consumption of amphipods by each fish was observed and noted. Very few fathead

minnows consumed amphipods within 30 minutes on the first few days, thus Tetramin

was provided to the fathead minnows to ensure fathead minnow survival. However, by

the second or third day, at least one fish per group training tank had consumed a few

amphipods. It was important that the fish learned to consume the amphipods quickly,

because during the exposure after 30 minutes the water within the tanks would be

renewed to avoid the possible excretion of microplastics by the amphipods, and the

potential direct ingestion of microplastics (and fluoranthene) by the fathead minnow.

After the group training, the fathead minnows that were capable of ingesting 15

amphipods within 30 minutes were removed from the group training tanks, and placed in

individual tanks for the remaining three days to ensure the fish would consume

amphipods alone (without competition). Each fish was then provided with 20 amphipods

each day for the remaining three days, and their ability to consume amphipods were

observed. Twenty fathead minnows were capable of consuming 20 amphipods quickly

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(within five minutes) after the group and individual training sessions and, therefore, were

used in the 3-d trophic transfer experiment.

Hyalella azteca exposure to polyethylene microplastics with and without

fluoranthene

Two days prior to the start of the 3-d trophic transfer experiment (the second day

of the group training), juvenile amphipods were exposed to 10,000 fluoranthene-

contaminated polyethylene microplastics (FLU-MPs)/mL and 10,000 polyethylene

microplastics/mL in 250 mL glass beakers containing USEPA hard reconstituted water

(HRW) amended as described in Borgmann (1996) (2.3.1. Organism Culture:

Freshwater amphipod Hyalella azteca) at 25 ± 1°C. The amphipods were exposed to

either FLU-MPs or MPs without FLU for a period of 48 hours prior to being used in the

3-d trophic transfer experiment; two more groups of amphipods were counted and

exposed to either 10,000 FLU-MPs or MPs without FLU (MPs/mL). The last set of

amphipods exposed to MPs with or without FLU on the first day of the 3-d trophic

transfer experiment were exposed to fathead minnows on day 3 of the trophic transfer

experiment). The exposure water containing either FLU-MPs or MPs without FLU were

statically renewed daily until being used in the 3-d trophic transfer experiment. Daily

water renewals ensured that sufficient oxygen was present in the water, and that no

aeration was needed while the amphipods were being exposed to MPs with or without

FLU.

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3-d Trophic transfer experiment

Each day, amphipods were removed from the exposure beakers containing MPs

with or without FLU, and gently rinsed with clean HRW three times to ensure all MPs

present on the cuticle of exposed organisms were removed. Twenty amphipods were

pipetted into tanks containing individual fathead minnows and the consumption of

amphipods was observed to occur within 30 minutes. Three hours after the feeding of

amphipods, the water within the tanks holding fathead minnows was statically renewed to

ensure the fathead minnows would not re-ingest fecal matter containing MPs or FLU.

On day three of the experiment, exposed fathead minnows were euthanized (pH

buffered 1.0 g Tricaine/L) two hours after being fed amphipods that had previously

ingested either MPs with or without FLU. The wet weight and length of the fathead

minnows were measured. The gallbladders were removed from exposed fathead

minnows, placed in individual amber 1.0 mL centrifuge tubes and stored in -80°C until

analysis.

Bile Analysis

Methods to analyze fluoranthene metabolites in the bile of exposed fathead

minnows follows those described in Linard et al. (2014). Briefly, after allowed to thaw,

the bile was drained from the gallbladder and diluted with 140 µL Nanopure water. The

diluted bile was then vortexed and centrifuged for 5 minutes at 14,000 rpm. Twenty-five

µL of diluted bile was then collected for a protein quantification assay, and 85 µL of

diluted bile was further diluted with 185 µL 50:50 MeOH to H2O solution. The

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fluorescence of fluoranthene metabolites in 250 µL of the diluted bile was analyzed in a

black 96-well plate at the excitation/emission wavelengths of 280/440 nm using a BioTek

Synergy H1 Hybrid Reader. GEN5 2.07 was the program used in tandem with the

BioTek Synergy H1 Hybrid Reader to attain the fluorescence readings. Total protein

content was quantified with the PierceTM bicinchoninic acid protein assay, using a

BioTek Synergy H1 Hybrid Reader to measure absorbance at 562 nm of the samples

together with a bovine serum albumin standard curve. After initial analysis of the data,

the bile fluorescence data were not normalized to protein concentrations because protein

samples were compromised during storage.

4.3.3 Microplastic trophic transfer from Pimephales promelas to Morone chrysops x

Morone saxatilis

Morone chrysops x Morone saxatilis culture

Morone chrysops x Morone saxatilis, hybrid striped bass, were chosen to be the

predator fish species for this study because our lab has previously conducted research

investigating whether the exposure to antidepressants would influence the predatory

behavior (i.e. ability to catch prey) of a predatory fish (hybrid striped bass) towards a

prey fish (fathead minnow) (Sweet et al. 2016). The ability to maintain a culture and

quantify and characterize predatory behavior of hybrid striped bass has been previously

demonstrated to occur successfully at the Clemson University Cherry Farm Aquatic

Laboratory facility. Juvenile hybrid striped bass fingerlings were purchased from Keo

Fish Farms, located in Keo, Arkansas, and maintained at the Clemson University Cherry

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Farm Aquatic Laboratory facility in a flow through system where water was pumped

from Lake Hartwell, and underwent filtering and ultraviolet treatments prior to going into

individual tanks. Fingerling hybrid striped bass were fed Zeigler’s trout food crumble #3

for the first two weeks, and then 2mm, 3mm and 5mm sized Zeigler’s trout food as the

fish reached sizes of 3-4, 4-5, and > 5 inches, respectively. As soon as the fingerling

hybrid striped bass reached a size between 4 and 5 inches, they were placed in 5-L tanks

for both the training sessions and 1-d trophic transfer experiment.

Pimephales promelas preparation

Four-day old Pimephales promelas were cultured at the Clemson University

Institute of Environmental Toxicology (CU-ENTOX) using methods described in Section

2.3.1. Juvenile fathead minnows were placed in a holding tank for four days, and then

used in either the training sessions or 1-d trophic transfer experiment. Newly hatched

fathead minnows were placed in new holding tanks at 25 ± 1°C with USEPA moderately

hard water that was statically renewed every day, for 8 days (7 days for the training, and

1 day for the trophic transfer experiment), to ensure sufficient numbers of fathead

minnows were available (USEPA 1989).

Group and individual training sessions

Hybrid striped bass were conditioned to consume pellet food, thus the fish had to

be trained to chase and consume live fathead minnows for the trophic transfer

experiment. Groups of three hybrid striped bass were placed in each tank that had black

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lining surrounding three of the four glass panels to limit stress that the hybrid striped bass

may experience by outside movement. The fish were stocked so as to not exceed 1.0 g

wet weight biomass/ L of water; and dissolved oxygen, and ammonia concentrations

were monitored during the training and exposure to ensure levels did not drop below 5

mg/L or exceed 0.5 mg/L, respectively. The same flow through system and temperature

ranges were maintained throughout the training sessions and trophic transfer experiment.

After the hybrid striped bass were placed in tanks, food was withheld from hybrid

striped bass for two days to increase the chance that some would feel inclined to feed

upon 4-d old fathead minnows. For the first four days following the two days in which

food was withheld, groups of 20 4-d old fathead minnows were placed in the tanks for a

period of 30 minutes. The fathead minnows used for the group and individual trainings

were fed less than 48-hour old brine shrimp; no microplastics were provided to the

fathead minnows used in the training sessions. After the four days of group training, 20

hybrid striped bass were able to consume 15 4-d old fathead minnows within 30 minutes.

The 20 hybrid striped bass were then placed in individual clean 5-L tanks, where their

ability to consume 15 fathead minnows was monitored over the next three days to ensure

the hybrid striped bass would consume fathead minnows alone (without competition)

Pimephales promelas exposure to polyethylene microplastics with and without

fluoranthene

Four-day old fathead minnows were exposed to 100,000 polyethylene MPs/mL

either with or without FLU for two days prior to being fed to hybrid striped bass during

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the 1-d trophic transfer experiment. Groups of fifteen fathead minnows were exposed to

MPs with or without FLU in 250 mL glass beakers, where the MP exposures were

statically renewed daily. Since the hybrid striped bass would only be fed once, and then

their bile would be analyzed for FLU metabolites following the one-time feeding

exposure, fathead minnows were exposed to MPs either with or without fluoranthene two

days prior to the 1-d trophic transfer experiment (starting from the second day of the

individual training session).


Fathead minnows were removed from the exposure beakers containing MPs with

or without FLU, and gently rinsed with clean moderately hard water three times to ensure

all MPs present on the outside of exposed organisms were removed. Fifteen fathead

minnows were pipetted into tanks containing individual hybrid striped bass and the

consumption of fathead minnows was observed to occur within 30 minutes. The amount

of time it takes for hybrid striped bass is variable, thus five hybrid striped bass that had

either consumed fathead minnows previously exposed to MPs without FLU (control), and

with FLU were destructively sampled at five and ten hours after being fed the fathead

minnows.

Hybrid striped bass were euthanized (pH buffered 1.0 g Tricaine/L) five and ten

hours after being fed fathead minnows that had previously ingested either MPs with or

without FLU. The wet weight and lengths of the hybrid striped bass were measured. The

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gallbladders were removed from exposed hybrid striped bass, placed in individual amber

1.0 mL centrifuge tubes and stored in -80°C until analysis.

Bile analysis

Methods to analyze fluoranthene metabolites in the bile of exposed hybrid striped

bass also followed those described in Linard et al. (2014) and described above. After

initial analysis of the data, the bile fluorescence data were normalized to protein

concentrations.

4.3.4 Microplastic trophic transfer from Amphiascus tenuiremis to Fundulus

heteroclitus

As described in Section 2.3.1, adult copepods were collected and counted to use

in the microplastic trophic transfer experiment (Chandler 2004). Briefly, mature

copepods were placed glass petri dishes containing 30 ppt artificially prepared salt water

prior to being used for training sessions and the 3-d trophic transfer experiment.

Group and individual training sessions

Three-week old mummichogs, Fundulus heteroclitus, were trained to consume

adult copepods over the course of nine days. During both the group and individual

training sessions, mummichogs were fed copepods that were not fed any microplastics or

fluoranthene; copepods were fed Dunaliella tertiolecta, as described in Section 2.3.1.

(Chandler 2004). The 250 mL glass beakers used for both group and individual training

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sessions were kept at 25 ± 1°C; and the 18 ppt salt water was statically renewed every

day.

Three mummichogs were placed within each beaker for the group training

sessions. The fish were stocked to not exceed 1.0 g wet weight biomass/ L of water; and

dissolved oxygen, and ammonia concentrations were monitored during the training and

exposure to ensure levels did not drop below 5 mg/L or exceed 0.5 mg/L, respectively.

Food was withheld from all mummichogs for the first 48 hours, to provide incentive for

the fish to consume copepods. During the four days of group training, each beaker was

given approximately 100 copepods each day, and the consumption of copepods by each

fish was observed and noted. Very few mummichogs consumed copepods within 30

minutes in the first few days, because in comparison to the brine shrimp that are fed to

juvenile fathead minnows and mummichogs, the copepods are transluscent, and are

harder to see. However, by the second or third day, at least one fish per group training

beaker had consumed a few copepods. It was important that the fish learned to associate

the pipette with food (i.e. copepods) quickly, because during the exposure, after 30

minutes, the water within the beakers was renewed to avoid the possible excretion of

microplastics by the copepods, and the potential direct ingestion of microplastics (and

fluoranthene) by the mummichog.

After the group training, the mummichogs capable of ingesting 20 copepods

within 30 minutes were removed from the group training beakers, and placed in

individual beakers for the remaining three days to ensure the fish would consume

copepods alone (without competition). Each fish was then provided with 20 copepods

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each day for the remaining three days, and their ability to consume copepods was

observed. Ten mummichogs were capable of consuming 20 amphipods quickly (within

five minutes) after the group and individual training sessions, and were used in the 3-d

trophic transfer experiment.

Amphiascus tenuiremis exposure to polyethylene microplastics with and without

fluoranthene

Two days prior to the start of the 3-d trophic transfer experiment (the second day

of the group training), juvenile copepods were exposed to 40,000 fluoranthene-

contaminated polyethylene microplastics (FLU-MPs)/mL and 40,000 polyethylene

microplastics/mL (no FLU) in 250 mL glass beakers containing 30 ppt artificially

prepared salt water 25 ± 1°C. Two more groups of copepods were counted and exposed

to either 40,000 FLU-MPs or MPs without FLU (MPs/mL). The last set of copepods

exposed to MPs with or without FLU on the first day of the 3-d trophic transfer

experiment were exposed to mummichogs on day 3 of the trophic transfer experiment.

The exposure water containing either FLU-MPs or MPs without FLU was statically

renewed daily until used in the 3-d trophic transfer experiment. Daily water renewals

ensured that sufficient oxygen was present in the water, and that no aeration was needed

while the copepods were exposed to MPs with or without FLU.

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Each day, copepods were removed from the exposure beakers containing MPs

with or without FLU, and gently rinsed with clean water three times to ensure all MPs

present on the cuticle of exposed organisms were removed. Twenty copepods were

pipetted into tanks containing individual mummichogs and the consumption of copepods

was monitored over 30 minutes. Two hours after the feeding of copepods to

mummichogs, the water within the tanks holding mummichogs was statically renewed to

prevent mummichogs from re-ingesting fecal matter containing MPs or FLU.

On day three of the experiment, exposed mummichogs were euthanized (pH

buffered 1.0 g Tricaine/L) two hours after being fed copepods that had previously

ingested either MPs with or without FLU. Juvenile mummichogs were still too young to

have developed large enough gall bladders, thus whole body burdens of fluoranthene

were quantified. Each mummichog was placed in an individual amber 1.0 mL centrifuge

tubes and stored in -80°C until analysis.

Fluoranthene body burden analysis

Methods to quantify fluoranthene that desorbed from microplastics indirectly

ingested through prey (i.e. copepods) were the same as those described in Section 3.3.3.

Briefly, following the removal of all overlying water that may have been transferred with

the exposed mummichog, 50 µL of isooctane, 10 µL methanol, and 40 µL of 4M sodium

hydroxide, and 15-30 (0.5 mm diameter) solvent cleaned zirconia beads

(Biospec11079105z Zirconia/Silica Beads) were added to each vial. Organisms were

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homogenized for 100 seconds at 3,000 oscillations/min using a Mini-BeadBeater

(Biospec Products, Bartlesville, OK, USA), vortexed for 10 s, and then centrifuged for 10

minutes at 3,500 g (Klosterhause et al. 2002). The top layer of each vial, which

contained fluoranthene and iso-octane, was removed and transferred to a clean vial and

stored at – 20°C until analysis. Fluoranthene in the iso-octane was quantified using black

polystyrene 96-well plates with a Molecular Devices Gemini fluorescence microplate

reader at 280/440 nm excitation/emission.

4.3.5 Data analysis

Determining whether MPs and the associated fluoranthene can undergo trophic

transfer was analyzed by using student’s t-tests to quantify whether there was a

significant difference in fluoranthene bioavailability between predator organisms

provided with prey organisms that were pre-exposed to MPs with or without

fluoranthene. Specifically, fluoranthene body burdens of the predator organisms

(juvenile mummichogs) were compared using student’s t-tests to determine if the indirect

exposure to fluoranthene-contaminated MPs through prey (i.e. adult copepods)

consumption resulted in significant fluoranthene body burdens in mummichogs, as

compared to mummichogs that were fed prey that were exposed to MPs without

fluoranthene. Similarly, fluoranthene bioavailability through prey consumption was also

quantified by analyzing the fluoranthene present in the bile of fathead minnows and

hybrid striped bass that consumed amphipods and fathead minnows, respectively, that

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were either pre-exposed to MPs with or without fluoranthene. These calculations were

performed using JMP Pro 11 Statistical Discovery software (SAS, Cary, NC, USA).

4.4 Results

4.4.1 Microplastic trophic transfer from Hyalella azteca to Pimephales promelas

Fathead minnows fed amphipods that were pre-exposed to 10,000 FLU-MPs/mL

had an average of 1.74 ± 1.68 mg of fluoranthene metabolites present in the bile (Figure

4.1). The ingestion of amphipods that were pre-exposed to FLU-saturated MPs by

fathead minnows resulted in a significant amount of fluoranthene (approximately 1.5

mg/organism) in the bile of fathead minnows. The fluoranthene indirectly ingested

through trophic transfer was bioavailable to exposed fathead minnows, and although not

quantified, MPs were found in the fecal matter of predator organisms.

4.4.2 Microplastic trophic transfer from Pimephales promelas to Morone chrysops x

Morone saxatilis

Hybrid striped bass fed fathead minnows that were pre-exposed to 100,000 FLU-

MPs/mL had an average of 0.42 ± 0.21 µg of fluoranthene metabolites present in the bile

(Figure 4.2). The fluoranthene indirectly ingested through trophic transfer was

bioavailable to exposed hybrid striped bass five hours after the feeding (approximately

1.3 mg/organism); there were no fluoranthene metabolites in the hybrid striped bile ten

hours after the feeding. The ingestion of fathead minnows that were pre-exposed to

FLU-saturated MPs by hybrid striped bass resulted in a significant amount of

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fluoranthene metabolites in the bile of hybrid striped bass, and although not quantified,

MPs were found in the fecal matter of predator organisms.

4.4.3 Microplastic trophic transfer from Amphiascus tenuiremis to Fundulus

heteroclitus

Mummichogs fed copepods that were pre-exposed to 40,000 FLU-MPs/mL had

an average body burden of 1.50 ± 1.32 mg fluoranthene metabolites (Figure 4.3). The

fluoranthene indirectly ingested through trophic transfer was bioavailable to exposed

mummichogs (approximately 1.5 mg/organism). The ingestion of copepods that were

pre-exposed to FLU-saturated MPs by mummichogs resulted in a significant body burden

of fluoranthene metabolites, and although not quantified, MPs were found in the fecal

matter of predator organisms.

4.5 Discussion

Microplastics can be introduced into a food web through their direct or indirect

ingestion (i.e. trophic transfer). Fathead minnows and mummichogs that consumed

amphipods and copepods that were previously exposed to microplastics, respectively,

resulted in the trophic transfer of microplastics to both fish. Although microplastics

within predator organisms were not quantified, microplastics were found within the

digestive tracts of all three predatory fish, after being fed prey that had been pre-exposed

to microplastics with and without fluoranthene. Further, the indirect exposure to

fluoranthene through the consumption of prey that had been pre-exposed to fluoranthene-

204

contaminated microplastics also resulted in the presence of fluoranthene metabolites in

mummichogs, as well as the bile of fathead minnows and hybrid striped bass.

Zooplankton and other small invertebrates are common prey for juvenile fish; and the

ingestion of microplastics by both pelagic and benthic invertebrates suggests that juvenile

fish will likely consume large amounts of prey that have been pre-exposed to

microplastics (Cole et al. 2013; Setälä et al. 2013; Au et al. 2015). Trophic transfer is a

likely pathway for microplastic exposure in aquatic ecosystems. Both H. azteca and A.

tenuiremis are epibenthic invertebrates, and are likely to be more susceptible to

microplastics that could settle through the water column; however, the ease at which

invertebrates are consumed by pelagic fish demonstrated how microplastic transport and

fate within the water column and sediment will be greatly influenced by prey and

predatory dynamics as well as the continuously changing physical and chemical

properties of microplastics in the environment.

Organisms are likely to preferentially consume microplastics that are a specific

size, shape or color. Amberstripe scad, Decapterus muroadsi, collected along the coast

of Easter Island had consumed higher amounts of blue polyethylene microplastics than

those of other colors or polymers; the preferred natural prey are blue copepods that are

similar in size and color as the blue microplastics consumed by the scad (Ory et al. 2017).

Microplastics can also be ingested through the consumption of aggregates or fecal matter

containing phytoplankton, and other organic matter in addition to microplastics (Long et

al. 2015; Long et al. 2016; Cole et al. 2013). Smaller microplastics are likely to be

consumed in high concentrations as microplastics often aggregate together, thus

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organisms that may have a larger food size preference may also inadvertently consume

smaller microplastics that formed larger aggregates. The adsorption of microplastics on

the external cuticle and gills of exposed organisms may also result in the trophic transfer

of microplastics (Watts et al. 2016). It is important to note that the consumption of

microplastics through prey capture is not the only way in which higher trophic organism

will be exposed to microplastics.

Spherical polyethyelene microplastics between 10 and 27 µm were used in all

three trophic transfer experiments; and, from my previous studies, their body residence

time in exposed predators did not exceed those of normally consumed food materials

(Chapter Two). Other studies have demonstrated that fibrous microplastics and those that

are smaller (< 0.5 mm) have longer residence times than larger spherical and fibrous

microplastics in exposed organisms (Farrell and Nelson 2013; Browne et al. 2008; Watts

et al. 2015). More information is needed on how different microplastic properties will

influence their uptake into and excretion from exposed organisms. Although it is

unlikely that microplastics consumed by prey, and then their predators will have

difficulties in being excreted in a much larger gastrointestinal tract, if small enough, the

microplastics may be able to translocate into other organs thereby potentially having

longer residence times in higher trophic organisms as well. Longer microplastic

residence times within prey organisms will result in greater amounts of time at which

associated contaminants can desorb from microplastics, and become bioavailable, thereby

also resulting in a greater possibility that predator organisms may be exposed to higher

concentrations of pollutants.

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Fluoranthene adsorbed to the consumed polyethylene microplastics was

bioavailable to the predator fish (fathead minnows, hybrid striped bass, and

mummichogs), suggesting that microplastics may be a significant vector of contaminants

to organisms through prey as well as through direct consumption of microplastics by the

predator fish. The amount of polycyclic aromatic hydrocarbon (PAH; 386 to 1996 ng/g

microplastics) found on microplastics collected from Santos Harbor, an important

industrialized port in Brazil, varied based on water depth (Fisner et al. 2013). The

highest concentration of PAHs was associated with microplastics found in the first 10 cm

of the sediment; the variability in PAHs found at different depths was attributed to matrix

differences, and polymer characteristics (Fisner et al. 2013; Barnes 2005; Morét-

Ferguson et al. 2010). Although the authors suggested the low concentrations of PAHs

found on the MPs are unlikely to cause harm, due to PAHs having a higher binding

affinity for plastic than sediment, the bioavailability of PAHs to aquatic organisms will

depend on physiological characteristics of exposed organisms and the amount of

contaminants present on MPs (Teuten et al. 2007). Like many other anthropogenic

pollutants, microplastic pollution is associated with more urban areas (McCormick et al.,

2016); once released into the environment, microplastics in urban areas are likely to sorb

higher concentrations of various pollutants, thus making organisms inhabiting such

regions more susceptible to a mixture of pollutants through microplastic exposure.

Microplastic trophic transfer studies have been limited thus far to zooplankton

food webs, and the mixture of microplastics in the feed of a predator organism; here I

demonstrated that although hybrid striped bass were able to eliminate ingested

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microplastics and fluoranthene within ten hours of being indirectly ingested through

fathead minnow consumption, the fact that the fluoranthene was bioavailable five hours

post-consumption demonstrates that humans may be susceptible to additional

contaminant exposure through fish consumption. Although the exposure through fish

consumption may be minimal as the internal organs are normally removed prior to

cooking the fish, human exposure to microplastics and associated contaminants through

bivalve and shellfish consumption is likely (Davidson and Dudas 2016; Van

Cauwenberghe and Janssen 2014).

No other study has investigated the toxicity of a single type of MP as a function of

acute and chronic exposures, and ability to influence the bioavailability of organic

pollutants in both freshwater and marine invertebrates and fish. Unfortunately, with the

versatility of plastic products resulting in an infinite variety of microplastics that may be

produced, it will be difficult to conduct studies that encompass a wide range of

toxicological endpoints. However, understanding how polymers will differentially

degrade in specific environmental conditions will not only provide information on

microplastic transport and fate, but also shed light on which organisms will be susceptible

to different types of microplastics. More progress is occurring for producing

biodegradable plastic products, yet more information is needed on the additives being

used and whether the quick release of those chemicals upon the degradation of the

polymer backbone will be toxic as well.

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4.6 Limitations

Trophic transfer studies are normally rare because feeding studies have many

variations due to organism differences in terms feeding abilities and timeliness. After

training the mummichogs, fathead minnows, and hybrid striped bass, approximately 75%

of the organisms could successfully catch and consume their respective prey, thereby

limiting the number of exposure treatments that could be conducted. Further, as the

amount of prey needed per predator organism was high throughout the training process

and bioassays, limited numbers of prey had to be taken into consideration when

determining the number of maximum prey that could be used during the exposure.

Unfortunately, it could not be determined whether the bioavailability of MPs or

fluoranthene is dose-dependent because only two treatments could be conducted in all

three trophic transfer experiments: predator organisms were fed prey that were either

exposed to MPs with or without fluoranthene. This objective’s goal was to ultimately

answer the question of whether or not trophic transfer is a realistic route of MP exposure

to predatory organisms. Another limitation to these experiments was the variations

associated with MP ingestion by the prey organisms. The results from Chapter One

demonstrated the large variability associated with MP ingestion by copepods and

amphipods. Practically, I could not quantify the number of ingested MPs by each prey

organism prior to feeding the prey organisms to a predator organisms, thereby making it

difficult to ascertain the number of actual MPs that were indirectly ingested by the

predator organisms. Another limitation of the experimental design of these trophic

transfer bioassays is the uncertainty associated with the metabolism of the predatory fish;

209

when quantifying the amount of fluoranthene present in either the whole body burdens of

mummichogs or bile of fathead minnows or hybrid striped bass, if the fluoranthene had

been excreted prior to the end of the experiment, then larger variations in fluoranthene

bioavailability would occur. For example, in hybrid striped bass trophic transfer

experiments, the bile from hybrid striped bass was removed after five and ten hours of the

initial feeding of prey. At ten hours, there was no significant difference between the bile

fluorescence of hybrid striped bass that were fed fathead minnows, previously exposed to

MPs with or without fluoranthene, whereas a significant difference was observed in

hybrid striped bass bile that was examined at hour five. Further, as we were unable to

distinguish the difference between the parent fluoranthene compound and the various

fluoranthene metabolites in the bile or whole body burdens, there are limitations in terms

of quantifying the ratio of bioavailable parent fluoranthene to fluoranthene metabolites.

4.7 Conclusions

The results from Chapter Four demonstrates that both MPs and their associated

contaminants can undergo trophic transfer in aquatic ecosystems. Not only did the

polyethylene MPs and adsorbed fluoranthene undergo trophic transfer between

invertebrates and juvenile fish, but they also were able to undergo trophic transfer

between two fish species. Trophic transfer serves as a realistic route of exposure to both

MPs and associated contaminants. Although amphipods and copepods are classified as

benthic and pelagic invertebrates, respectively, both realistically spend portions of their

life cycle in both regions of the water columns. Fathead minnows, hybrid striped bass,

210

and mummichogs are all pelagic fish, and as they consume prey that dwell in both pelagic

and benthic habitats, food webs encompassing the entire water columns are likely to be

susceptible to MPs that either remain buoyant in or sink through the water column. No

other study has demonstrated whether organisms in the higher levels of biological

organization can be susceptible to MPs indirectly through prey consumption. Further, as

hybrid striped bass are commonly consumed by humans and other terrestrial organisms, it

is possible for MPs and associated contaminants to be transferred from aquatic to

terrestrial ecosystems.

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4.8 Figures

Figure 4.1: Fluoranthene in bile of fathead minnows due to trophic transfer

Mean fluoranthene concentration (SD = ± 864.8) present in bile in fathead minnows

exposed to fluoranthene-saturated microplastics through prey (20 amphipods/fathead

minnow). Amphipods were exposed to 10,000 fluoranthene-saturated microplastics/mL.

Compared to the control (fathead minnows consumed amphipods that had consumed

10,000 microplastics/mL without fluoranthene), there is a significant difference

(symbolized by the asterisk) between the two treatments (p < 0.05).

0

500

1000

1500

2000

2500

3000

3500

4000

Control FLU-MPs

FLU

(µg)

/org

anism

Treatments

*

212

Figure 4.2: Fluoranthene in bile of hybrid striped bass due to trophic transfer

Mean fluoranthene concentration (SD = ± 0.09) present in bile of hybrid striped bass

exposed to fluoranthene-saturated microplastics through prey (15 fathead

minnows/hybrid striped bass). Fathead minnows were exposed to 100,000 fluoranthene-

saturated microplastics/mL. Different letters represent significant differences in

fluoranthene metabolite concentrations in the bile of hybrid striped bass exposed to

fathead minnows that were exposed to 100,000 MPs/mL without FLU, and 100,000

MPs/mL with FLU (MP+FLU), 5 and 10 hours after ingestion.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

5hr MP 5hr MP+FLU 10hr MP 10hr MP+FLU

µg F

LU/m

g pr

otei

n

Treatment

a a

b

ab

213

Figure 4.3: Fluoranthene whole body burden of mummichogs due to trophic

transfer

Mean fluoranthene body burden of mummichogs (SD = ± 720.3) exposed to

microplastics without (MP-only) and with fluoranthene (MP+FLU) through prey (20

copepods/mummichog). Copepods were exposed to 40,000 fluoranthene-saturated

microplastics/mL. Compared to the control (mumichogs fed copepods that had consumed

40,000 microplastics/mL without fluoranthene), there is a significant difference

(symbolized by the asterisk) between the two treatments (p < 0.05).

0

500

1000

1500

2000

2500

3000

MP (No FLU) MP+FLU

Fluo

rant

hene

bod

y bu

rden

(µ

g FL

U/o

rgan

ism)

Treatments: 100,000 MPs/mL without FLU (No FLU) and with FLU (MP+FLU)

*

214

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CHAPTER FIVE

CONCLUSIONS The questions surrounding the toxicological implications of microplastic pollution

in the environment are just beginning to be answered; my research attempts to bridge

some of the early research gaps regarding different organism responses to microplastic

exposure, and whether microplastics can act as a source and vector for persistent organic

pollutants as exemplified by the polycyclic aromatic hydrocarbon, fluoranthene.

The first objective for my dissertation focused on the investigation of microplastic

toxicity to aquatic organisms. There is little information available on the chronic toxicity

of microplastics to aquatic organisms, thus chronic microplastic exposures were

conducted using the freshwater amphipod, Hyalella azteca, and the marine copepod,

Amphiascus tenuiremis, after acute lethal thresholds were established for these two

invertebrates. The 10- and 7-d LC50s for spherical polyethylene microplastics for

exposed H. azteca and A. tenuiremis were 4.64 x 104 and 7.90 x 104 microplastics/mL,

respectively. H. azteca was also acutely exposed to fibrous polypropylene microplastics;

the LC50 was 71 microplastics/mL, several orders of magnitude less than the 10-d LC50

for spherical polyethylene microplastics. However, from the limited acute microplastic

exposures to H. azteca, I am unable to conclude whether microplastic shape or polymer

plays a more influential role in determining microplastic toxicity to exposed amphipods.

My results do demonstrate that different microplastics will have toxicological effects at

different concentrations.

220

Chronic exposures to spherical polyethylene and fibrous polypropylene

microplastics resulted in significant reductions in growth and reproduction in exposed H.

azteca. Body residence times of the fibrous microplastics were significantly higher than

those of the spherical microplastics, suggesting that greater microplastic body residence

times may reduce nutrient uptake, thereby exerting sublethal effects on overall organism

growth and neonate production. Although reproduction was not affected during chronic

spherical polyethylene microplastics exposures to A. tenuiremis, growth was significantly

affected, where developmental time was dose-dependent, resulting in slower growth.

For the vertebrate studies, fathead minnows, Pimephales promelas, and

mummichogs, Fundulus heteroclitus, were acutely exposed to polyethylene

microplastics. There was no mortality for either fish within the exposure concentrations

used (0-100,000 microplastics/mL). Despite body residence times not being significantly

different between food items and spherical microplastics in all exposed organisms

(invertebrates and vertebrate), microplastic ingestion was dose-dependent. Acute

exposures to microplastics did not have any negative effect on exposed fish in these

experiments.

The second objective for my dissertation investigated whether microplastics can

influence the bioavailability of fluoranthene to exposed H. azteca, A. tenuiremis, P.

promelas, and F. heteroclitus. Acute exposures to fluoranthene-contaminated spherical

polyethylene microplastics resulted in fluoranthene being bioavailable to exposed

amphipods and copepods in a dose-dependent manner. The fluoranthene body burden in

exposed invertebrates was primarily due to the ingestion of fluoranthene-contaminated

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microplastics, where only 5% of the fluoranthene body burden was due to the

fluoranthene that had desorbed from the microplastics during the acute exposure. When

fathead minnows and mummichogs were exposed to fluoranthene-contaminated

microplastics, fluoranthene was also bioavailable; however, a majority of the body

burden was due to the fluoranthene that had desorbed into the exposure water.

The third objective for my dissertation evaluated whether microplastics and

adsorbed fluoranthene could undergo trophic transfer. H. azteca and A. tenuuiremis were

exposed to fluoranthene-contaminated polyethylene microplastics for two days prior to

being fed to P. promelas and F. heteroclitus, respectively, resulting in the trophic transfer

of both microplastics and fluoranthene to both predatory fish. Interestingly, there was

approximately 1.5 mg fluoranthene per fathead minnow and mummichog that fed upon

prey that had been pre-exposed to fluoranthene-contaminated microplastics. Hybrid

striped bass that were fed fathead minnows that had been pre-exposed to fluoranthene-

saturated microplastics also resulted in the presence of 1.3 mg fluoranthene in the bile of

hybrid striped bass. The results demonstrate that the indirect ingestion of microplastics

and adsorbed contaminants through prey consumption is a pathway in which

microplastics can act as a vector of environmental contaminants not only through direct

exposure, but through trophic transfer.

My research is the most comprehensive study to date on mircoplastic toxicity

because it focused on both freshwater and marine invertebrates and fish. The production

of data that used similar exposure methods allows the scientific community to make

cross-species comparisons that are difficult to determine with the currently available

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microplastic toxicity data. As previously mentioned, there are many different types of

plastic products, and the varying field collection, sample preservation, and analysis

methods make it difficult to determine realistically what environmental microplastic

concentrations actually exist; current research underestimates environmental microplastic

concentrations that are less than 5 mm in diameter, and the vast array of toxicological

effects that both direct and indirect microplastic exposures have on wildlife. My results

demonstrate that microplastics can have both acute and chronic toxicological effects on

aquatic organisms, and that the presence of microplastics in the environment needs to be

taken into consideration when evaluating the exposure to environmental contaminants.

5.1 Future research needs

Critical research needs include standardizing methods of field characterization of

microplastics; quantifying uptake and depuration rates in organisms at different trophic

levels; quantifying the influence that microplastics have on the uptake and/or depuration

of environmental contaminants among different trophic levels; and investigating the

potential for biomagnification of microplastic-associated chemicals. More integrated

approaches involving computational modeling are required to fully assess trophic transfer

of microplastics. Suggestions that arise from my results include the following for future

research (Au et al., in press).

1. Standardize microplastic collection, preservation, and characterization

methods:

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It is difficult to compare the effects of concentrations of microplastics found in

the environment and those used in laboratory settings in toxicity tests. Methods

may need to be polymer- and media-based (i.e. water, sediment samples).

Polymers have different properties, and some of the harsh solvents and

temperatures used to eliminate organic materials found on plastic debris may alter

or even dissolve certain polymers, resulting in an over- or underestimation of

microplastics found in different field samples. For laboratory experiments,

researchers need to provide a concentration of microplastics in terms of both mass

and total number of microplastics per unit of volume. Using one or the other

makes it difficult to compare concentrations of microplastics used in different

exposure treatments as the mass of microplastics (which is most often reported)

will be affected by the density and size of the microplastics used.

2. Quantify microplastic uptake and depuration rates (body retention times) in

exposed organisms of different trophic levels, and determine what factors

influence body retention times:

To understand how microplastics and their associated contaminants may undergo

bioaccumulation and biomagnification, it is necessary to understand whether

exposure to microplastics with certain characteristics will result in an increased

uptake rate and/or decreased depuration rate. Some studies have demonstrated

that spheres ≤10 µm and fibers have longer body residence times, suggesting that

these microplastics will have a greater chance at being indirectly ingested by a

predatory organism (Farrell and Nelson 2013; Watts et al. 2014; Au et al. 2015).

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Very little is known about the behavior of microplastics in organisms of higher

trophic levels after ingestion of prey species that had accumulated microplastics.

3. Determine whether microplastics can act as a significant source or

depuration pathway for environmental contaminants:

Many studies that use thermodynamic modeling to justify whether microplastics

can act as a significant source of persistent organic pollutants and other

contaminants assume equilibrium within the time span of the exposure. However,

some studies have demonstrated that considerable time may be needed for smaller

microplastics to reach equilibrium with the surrounding seawater, suggesting that

organisms in urban areas that are exposed to large numbers of microplastics with

high concentrations of adsorbed contaminants may be at greater risk for

microplastics acting as a source of contaminants, where the kinetics may favor the

desorption of contaminants from the ingested microplastics. The reverse could

also be true; microplastics that have not had time to reach equilibrium with their

environment may be ingested by organisms that have higher concentrations of

pollutants, thus favoring sorption of pollutants onto ingested microplastics.

4. Investigate the biomagnification effect of different classes of microplastic-

associated chemicals at different trophic levels:

Different classes of chemicals associated with microplastics, and that as such are

transferred through food chains, may have different effects in organisms from

different trophic levels. For example, it is known that PAHs are poorly

biotransformed in invertebrates, but well metabolized in vertebrates. The potential

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for toxic effects at higher trophic levels after biomagnification of microplastics

and their associated environmental contaminants needs investigation.

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5.2 References

Au, SY, Bruce, TF, Bridges, WC, Klaine, SJ. 2015. Responses of Hyalella azteca to acute and chronic microplastic exposures. Environ Toxicol Chem 34: 2564-2572. Au, SY, Lee, CM, Weinstein, JE, van den Hurk, P, and Klaine, SJ. 2017. Trophic transfer of Microplastics in aquatic ecosystems: identifying critical research needs. Integr Enviro Assess Manage, in press. Farrell, P, Nelson, K. 2013. Trophic level transfer of microplastic: Mytilus edulis (L.) to Carcinus maenas (L.). Environ Pollut 177: 1-3. Watts, AJR, Lewis, C, Goodhead, RM, Beckett, SJ, Moger, J, Tyler, CR, Galloway, TS. 2014. Uptake and retention of microplastics by the shore crab Carcinus maenas. Environ Sci Technol 48: 8823-8830.

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Toxicity of Microplastics to Aquatic Organisms