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Fate of Nanoplastics in Marine Larvae: A Case Study Using Barnacles, Amphibalanus amphitrite Samarth Bhargava, Serina Siew Chen Lee, Lynette Shu Min Ying, Mei Lin Neo, Serena Lay-Ming Teo, and Suresh Valiyaveettil ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.8b00766 • Publication Date (Web): 21 Mar 2018 Downloaded from http://pubs.acs.org on March 22, 2018

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ACS Sustainable Chemistry & Engineering

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Fate of Nanoplastics in Marine Larvae: A Case Study Using

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Barnacles, Amphibalanus amphitrite

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Samarth Bhargavaa, Serina Siew Chen Leeb, Lynette Shu Min Yingb, Mei Lin Neob, Serena

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Lay-Ming Teob*, Suresh Valiyaveettila,*

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a

Materials Research Laboratory, Department of Chemistry, National University of Singapore, 3 Science Drive 3, Singapore 117543 b

St. John's Island National Marine Laboratory, Tropical Marine Science Institute, National

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University of Singapore, 18 Kent Ridge Road, Singapore 119227

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*Email: [email protected], [email protected]

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Abstract:

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The exposure of nanoplastics was investigated by observing their interaction with

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Amphibalanus amphitrite (commonly known as acorn barnacles). Poly(methyl methacrylate)

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(PMMA) and fluorescent perylene tetraester (PTE) dye were used to prepare highly

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fluorescent nanoplastic particles. At concentrations of 25 ppm, the PMMA particles showed

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no detrimental impact on barnacle larvae and their microalgae feed, Tetraselmis suecica and

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Chaetoceros muelleri. PMMA nanoplastics were ingested and translocated inside the body of

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the barnacle nauplii within the first 3 hours of incubation. The fluorescent PMMA particles

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inside the transparent nauplius were tracked using confocal fluorescence microscopy.

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Subsequently, the nanoplastics were fed to the barnacles under two conditions – acute and

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chronic exposure. The results from acute exposure show that nanoplastics persist in the body

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throughout stages of growth and development – from nauplius to cyprid and juvenile barnacle.

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Some egestion of nanoplastics was observed through moulting and faecal excrement. In

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comparison, chronic exposure demonstrates bioaccumulation of the nanoplastics even at low

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concentrations of the plastics. The impacts of our study using PMMA nanoparticles exceeds

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current knowledge, where most studies stop at uptake and ingestion. Here we demonstrate

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that uptake of nanoparticles during planktonic larval stages may persists to the adult stages,

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indicating potential for the long term impacts of nanoplastics on sessile invertebrate

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communities.

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Keywords: Microplastics, Nanoplastics, Marine Pollution, Barnacles, Fluorescent particles,

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Poly(methyl methacrylate)

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Introduction

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Globally, plastic waste materials have caused significant pollution in both lands and oceans.1

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Plastics with sizes smaller than 5 mm used as exfoliants in cosmetics and healthcare products

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are usually referred to as primary microplastics. Common plastic articles undergo degradation

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or weathering over time in nature to produce smaller sized particles, referred to as secondary

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microplastics by the United Nations Environment Programme (UNEP).2 Such plastic

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particles and microbeads have been accumulating in our environment and starting to show

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adverse health effects on different marine animals.3-5 Recently, microplastics have been

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found in various marine organisms ranging from barnacles, bivalves to fish.6

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Bioaccumulation of these plastic pollutants in such animals led to a transfusion and buildup

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in the food chain.7 Attempts to understand the source and fate of microplastic particles inside

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living organisms have gained significant attention owing to the potential impacts on human

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health, contamination of food supply and the environment.5-6, 8-10

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Few groups have investigated the presence of microplastics with a size range of 1 µm – 5

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mm,2, 5-6, 11-16 but even fewer data are available on the influence of smaller nanoplastics (1 nm

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– 1 µm)17-19 on marine animals. There is mounting evidence to show that numerous marine

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organisms such as plankton, fish and seabirds are readily consuming microplastic particles. A

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study in 2016 along the eastern coast of Brazil demonstrated and highlighted that edible fish

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were ingesting microplastics at rapid rates: 33% by the Brazilian sharpnose shark and 62.5%

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by the king mackerel.20 As many as 22% of the marine fish studied had plastic pellets (size 1

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to 5 mm) in their stomachs. Researchers have found up to 9,200 particles of microplastics per

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cubic meter of seawater in the coastal seas of Vancouver, Canada.21 While the focus of

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studies has mostly been on microplastics, few have investigated the effects of nanoplastics on

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marine organisms. The handful of studies focused on the toxicity and lethal effects of metal-

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based nanoparticles on the physiology, survivorship, and growth of various organisms such as

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zebrafish, barnacles, bivalves, and human cell line.

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potential effects of polymer-based nanoparticles on marine organisms. Nanoparticles, owing

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to their smaller size, are easily absorbed into cells and tissues. For this study, we focused on

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examining the ingestion, translocation and egestion of polymer-nanoparticles (ca < 200 nm in

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diameter) by the acorn barnacle, Amphibalanus amphitrite, larvae model under controlled

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environmental conditions.

22-26


Few studies have explored the

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The acorn barnacle larvae were selected for our studies owing to their transparent body, a

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short developmental period of seven days from larval stage to settlement onto substrata, and a

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well-established culture protocol.27-29 In this study, we used poly(methyl methacrylate)

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(PMMA), which is widely used in the manufacturing of paints, contact lenses, resins and in

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acrylic glasses,30 for preparing nanoplastic particles. The ease of processing and low toxicity

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makes PMMA an ideal polymer for testing the impact of nanoplastic particles inside the

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larvae.31 The present study investigates the potential toxicity on the larvae and, its microalgae

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feeds, as well as the ingestion, translocation and egestion of fluorescent PMMA nanoparticles

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through various stages of larval development of barnacle, A. amphitrite.

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Materials and Methods

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Preparation of Perylene Tetrabutyl Ester (PTE) Loaded PMMA Nanoplastics:

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Nanoplastics were prepared by using an established nanoprecipitation technique.32-35 The

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fluorescent dye, PTE was synthesized according to a reported procedure.36 To synthesize

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nanoparticles, a stock solution of PMMA (400 mg, MW: 15,000), SDS (7.7 mg, 26.6 µmol)

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and PTE (17.4 mg, 26.6 µmol) was prepared in acetone (100 mL). Appropriate amounts of

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this solution (5 mL) were diluted with ultra-pure water (50 mL) and stirred for 18 hours to

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remove excess acetone. The solution was filtered to obtain the nanoparticles and

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characterized using a range of techniques. The hydrodynamic size in DI water and

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morphology of the nanoplastics were determined using dynamic light scattering (DLS) and

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scanning electron microscope (SEM), respectively. The photophysical properties were

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characterized by ultra-violet and photoluminescence spectroscopy. Zeta potential and

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conductivity of the microplastic dispersion were measured using a zetasizer. The dispersed

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nanoplastic solutions were stable in DI water for 6 months. On addition of nanoplastics to

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seawater, the change in ionic strength led to flocculation of nanoplastic particles at

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concentrations above 50 ppm.

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Collection and Spawning of Barnacle Larvae: Acorn barnacles, A. amphitrite were

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collected during a low tide from intertidal mudflat at Kranji, Singapore (N1°26′18″,

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E103°45′25″). Under laboratory conditions, the barnacles released stage II nauplii. The

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phototactic stage II nauplii were concentrated by shining light through a transparent window

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in the spawning container and were collected using a pipette.28 The collected nauplii were

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transferred to 1 µm filtered seawater (FSW) at 27 Practical Salinity Unit (PSU) and counted

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using a Bogorov tray to determine the density of larvae in seawater.37 3 ACS Paragon Plus Environment


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Biological Studies: All assays were done in triplicates and average values are reported.

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Toxicity Assays of Nanoplastics Using Barnacle Larvae: The aim of the toxicity assays

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was to ascertain a sub-lethal concentration range of nanoplastics in the water column to

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enable further growth experiments. To test the potential toxicity of nanoplastics on barnacle

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larvae, stage II nauplii, were exposed to PMMA particles at concentrations of 5, 10 and 25

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ppm over a period of 24 hours. For each treatment level, 30 nauplii were added to every 2

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mL-glass vials (n = 3), and the appropriate volumes of PMMA nanoparticle stock solution

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(50 – 250 µL, 100 ppm) were added and topped up to a final test volume of 1 mL using clean

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FSW. The addition of PMMA nanoparticles had altered the salinities of solutions, hence,

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respective controls for each concentration were included in the experiments. Control

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solutions were prepared with comparable salinities but contained no PMMA. The vials were

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gently swirled, maintained at room temperature, and incubated for 24 hours. At the end of

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experiments, test solutions were transferred onto a Bogorov tray, followed by counting of

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living and dead nauplii using a stereomicroscope. Mortality counts were used to determine

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the relative toxicity of the nanoplastics to nauplii.

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Interaction of Nanoplastics with Microalgae: The growth and development of barnacle

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larvae require addition of microalgae as a feed.38-39 Even if microalgae do not assimilate the

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PMMA nanoparticles, the presence of such light-absorbing particles may influence the

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growth and proliferation of the algal cells in the medium. Thus, it is necessary to understand

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the effects of the nanoplastic particles on microalgae. The growth of the algal cultures,

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Tetraselmis suecica (CSIRO CS-187) and Chaetoceros muelleri (CSIRO CS-176), in f/2

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media, was monitored over 24 hours by exposing them to different concentrations of PMMA

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nanoparticle solutions.40

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Each microalgal stock solution was diluted to an approximate concentration of ca 100,000

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cells/mL. Five 1 mL aliquots of microalgal stock solutions were exposed to each nanoplastic

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concentrations of 0, 5, 10 and 25 ppm in 24 well plates. Control experiments comprised the

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respective microalgae stock solution with no PMMA particles and were used to determine the

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actual initial microalgal count by averaging the counts present in a 1 µL volume, using a

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Sedgwick Rafter. Samples were grown and incubated in f/2 media at 24°C and 33 PSU under

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a 12:12 hours light-dark cycle. The algal cells were observed under a confocal microscope

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(Olympus FV1000) to observe intake and interaction between algae and nanoplastics.


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Rapid Uptake of Nanoplastic Particles by Barnacle Nauplii: The continuous uptake of

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nanoplastics over time was studied at regular intervals of 45 minutes. Freshly spawned stage

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II nauplii were collected in a beaker containing clean FSW (200 mL) and stocked at a density

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of 38 nauplii/mL. PMMA nanoparticle solution (400 ppm) was added to the media with

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nauplii and the resultant solution was topped up with FSW to a volume of 250 mL to achieve

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a nanoplastic concentration of 25 ppm with a stocking density of 30 nauplii/mL. The solution

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was incubated at room temperature (25 °C). Samples (50 mL) of nauplii suspension were

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collected at four-time intervals (45, 90, 135 and 180 minutes), filtered, washed, and re-

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suspended in 50 mL FSW. The washed nauplii were fixed using 1% glutaraldehyde solution

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and imaged using a confocal microscope to visualize the presence and distribution of PMMA

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nanoplastics.

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Translocation of Nanoplastics within Barnacle Nauplii: To find out if the barnacle nauplii

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retain the nanoplastic particles within their bodies, 7,500 larvae were incubated with PMMA

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nanoplastics (25 ppm) for 3 hours. Nauplii were washed thoroughly with FSW to remove the

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remaining nanoplastic particles and maintained in clean FSW for further observations. The

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presence and distribution of nanoplastics inside the nauplii were monitored at 30, 75, 105,

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135, 165 and 180 minutes post-washing using a confocal microscope. Subsequently, the

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movement of nanoplastics inside the nauplii was continuously monitored at every 3 hours for

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a period of 24 hours (SI5). At 24 hours, a z-stack confocal image was also recorded to obtain

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a three-dimensional distribution of nanoplastics inside the nauplii body (AVI, Supporting

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Information).

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Effect of Acute and Chronic Exposure of Nanoplastics to Barnacle Larval Development:

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The effect of PMMA nanoplastics exposure time on barnacle nauplii was investigated under

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two conditions: acute and chronic exposure. Acute exposure is defined as exposure to a pulse

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of high concentration of nanoplastics over a short time interval, whereas chronic exposure is

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defined as a continuous exposure to a low dose concentration of nanoplastic solution

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throughout the entire life cycle of the organism. All experiments were run concurrently with a

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control (i.e. the absence of nanoplastics) under identical growth conditions.

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In a typical experiment, freshly spawned stage II nauplii were collected and divided into three

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batches for each treatment. For the controls, nauplii were stocked at 3 larvae/mL in 800 mL

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FSW and reared on a diet of an algal mixture of 1: 1 v/v of Tetraselmis suecica and

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Chaetoceros muelleri (5 x 105 cells/mL) at 28 °C with gentle aeration. For acute exposure, 5 ACS Paragon Plus Environment


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nauplii were concentrated at a density of 30 larvae/mL in 500 mL FSW and mixed with 25

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ppm of PMMA nanoparticle solution (400 ppm) and incubated for 3 hours at room

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temperature. After the incubation period, nauplii were filtered and washed thoroughly with

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excess FSW. The washed nauplii were restocked at 3 larvae/mL in 800 mL FSW and reared

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with the same feed regime as controls described earlier. For chronic exposure, nauplii were

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prepared and reared under the same conditions as controls, in presence of PMMA

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nanoplastics dosed at 1 ppm every two days.

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Nauplii of respective treatments were individually filtered through 50 µm mesh to remove

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faeces and uneaten feed, then re-suspended in clean FSW (800 mL) containing fresh

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microalgae feed once every two days until the end of the feeding experiments (i.e. 5 days).

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Fresh nanoparticle solutions of 1 ppm were added only to the chronic exposure experiments.

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On this feed regime, nauplii larvae metamorphosed into cyprids within five days and into

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juveniles within seven days.

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Samples were collected at the following time points, T = 0 (before exposure to nanoplastics),

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T = 2 days (Stage III nauplii), T = 5 days (cyprids), T = 7 days (settled barnacle juveniles).

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During collection of cyprids, moults and faecal matter were also collected from three

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treatments (control, acute and chronic conditions). Cyprids collected on day 5 were filtered

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through 50 µm mesh, washed with clean FSW prior to being distributed into 35 mm petri

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dishes, allowed to settle and metamorphosed into juveniles over the next two days. During

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the settlement period all the juveniles were maintained in FSW in the absence of feed and

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nanoplastics.

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Imaging of Specimens

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Sample Preparation

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To examine the effects of PMMA nanoplastics on the growth and development of barnacle

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larvae, various life stages of barnacle life cycle were recorded; nauplii stage III (2 days),

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cyprid (5 days) and juvenile (7 days). Faecal matter and moults were also collected and

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analyzed from the same experiment. All samples were sieved through a 50 µm mesh, washed

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with fresh FSW, re-suspended in FSW, fixed with 1% v/v glutaraldehyde solution and stored

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at 4°C before imaging with confocal microscopy. Prior to imaging, samples were brought to

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room temperature and transferred into Petri dishes. All samples were imaged with an

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Olympus FV1000 Confocal Microscope. 6 ACS Paragon Plus Environment

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Imaging Methods: Imaging of the PMMA nanoplastic distribution and retention inside the

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nauplii (Supporting Information, Figure S5) was conducted using an inverted microscope

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(Olympus BX51) with a mercury lamp as light source. Samples were focused on bright field

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mode and images were recorded using Differential Interference Contrast (DIC) mode with

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green fluorescence filter (filter cube: U-MGFPHQ, Ex: 460 – 480 nm, Em: 495 – 540 nm).

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MetaMorph 9.0 software was used for recording the images. The exposure for all samples

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was kept constant. Overlays of the images were also made to visualize the nanoplastic

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particle distribution.

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The collected samples of various larval stages of nauplii were analyzed using a confocal

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microscope (Olympus FV1000). All samples were imaged with reference to a control batch

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which was not exposed to nanoplastic particles. Laser power, magnification and brightness of

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the samples were preset to ensure minimum background noise. Z-stacks of samples were

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recorded to obtain a layer by layer profile of nanoplastics ingested-nauplii. For each larval

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stage, at least 10 unique z-stacks were recorded. Images were recorded in DIC and FITC

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mode. The excitation wavelength of the laser used was at 473 nm and emission signals were

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processed with a (Sharp-cut Dichroic Mirror) SDM560 filter.

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Analysis of Images: Images from the confocal microscope were qualitatively analyzed to

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determine relative fluorescence from the nauplii. To enhance the fluorescence intensity of

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images, the z-stack images were processed using Olympus Fluoview Ver. 4.2a Viewer. The

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signal saturation of all samples was adjusted in an identical manner to obtain a clear

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qualitative analysis of nanoplastic localization. FluoView and MetaMorph were also used to

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automatically introduce scale bars into the recorded images.

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Statistical Analyses: To determine the toxicity of PMMA nanoparticles to barnacle nauplii,

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the mean percent survivorship from each PMMA treatment were compared with their

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corresponding salinity adjusted controls using one-way analysis of variance (ANOVA). The

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toxicity of PMMA across different concentrations was also evaluated separately using one-

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way ANOVA. Similarly, the effects of PMMA nanoparticles on the mean cell count of each

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microalgae species were compared using one-way ANOVA.

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Results and Discussion:

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Preparation and Characterization of Nanoplastic Particles: The nanoparticles of PMMA

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incorporated with PTE dye were synthesized using nanoprecipitation. PMMA nanoplastic

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solutions (in DI water) with an average particle size, zeta potential and conductivity of 185 ± 7 ACS Paragon Plus Environment


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3 nm, -35.1 ± 0.4 mV and 0.0188 ± 0.0010 µScm-1, respectively, were obtained in large

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quantities. The fluorescent hydrophobic PTE dye was stabilized inside the PMMA particle.

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The photophysical properties of the PMMA particles were determined using Uv-Vis and

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photoluminescence spectroscopy (Figure 1A). The absorption maxima of the PTE dye

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molecules inside the PMMA particles were at 450 and 470 nm, and emission maximum was

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at 565 nm. The shape of absorption maxima and intensity of emission from the encapsulated

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PTE (Figure 1A) indicates no aggregation of dye molecules inside the particles. Precipitation

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of PTE alone in the absence of polymer led to the formation of rod-shaped particles, which

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showed weak fluorescence. This is expected due to the aggregation induced quenching of

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fluorescence from the dye molecules. For comparison, the spectra of a dilute solution of PTE

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in acetone and solid precipitated from solution are provided in the Supporting Information,

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Figure S1. The particle size distribution of the nanoplastics in DI water and sea water were

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also measured after dilution and incubation for 24 hours at a concentration of 25 ppm (Figure

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1C, 1D). The particles dispersed in DI water exhibited an average particle of 160 nm whereas

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the size of particles dispersed in FSW was measured to be ~600 nm.

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Figure 1: (A) Absorption (-●-) and emission (-▲-) spectra and (B) zeta potential of PMMA

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nanoparticle solution. DLS curves of PMMA nanoplastics measured at 25 ppm in DI water (C) and

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filtered sea water (D).


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Toxicity of PMMA Nanoplastics on Barnacle Nauplii: To assess the toxicity, around 30

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nauplii were placed in separate glass vials were exposed to PMMA particles at 5, 10 and 25

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ppm concentrations and incubated for 24 hours. For each concentration, triplicate

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measurements were done and the average value is reported. Appropriate controls were also

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carried out to validate our data. The mortality count was used to determine the relative

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toxicity of the polymer nanoparticles. No significant differences in toxicity were detected for

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all concentrations of PMMA nanoplastics investigated, as compared to their corresponding

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salinity-adjusted controls (Table S1). The proportions of larvae that survived were relatively

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high, i.e. >80% (Figure 2), suggesting that PMMA had no obvious toxicity on the nauplii. In

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addition, when only PMMA concentrations were compared, there were no significant

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differences in mean percent survivorship (ANOVA, p > 0.05, Table S1). PMMA is also

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known to be non-toxic and widely used in many commercial products such as medical

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implants.31 In a recent study, the toxicity of PMMA nanoparticles tagged with fluorophores

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was tested on Daphnia magna larvae.19 At concentrations as high as 1000 ppm, PMMA

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nanoparticles exhibited no significant toxic effects on D. magna larvae.18 This is consistent

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with the concentrations tested on the barnacle nauplii in our studies. Similar toxicity assays

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using polystyrene nanoplastics were also conducted by other groups on A. amphitrite and

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observed relatively low toxicity.29, 41

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When visualizing the nanoparticles-exposed larvae, the low concentrations (i.e. 5 and 10 ppm)

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showed lower fluorescence intensities during imaging, as compared to the larvae exposed to

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the concentration of 25 ppm. Given that PMMA showed no negative impacts on nauplii, and

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no flocculation and sedimentation were observed at a concentration of 25 ppm, subsequent

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experiments were done at this concentration.



1 2

Figure 2: Percentage survival of larvae after exposure to PMMA nanoplastics at concentrations of 5,

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10 and 25 ppm compared to respective salinity-adjusted controls, for 24 hours.

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Interaction of Polymer Nanoplastics with Microalgae: The established laboratory culture

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protocols use a mixture of two different microalgae, Tetraselmis suecica and Chaetoceros

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muelleri, as a feed to provide the necessary nutrients for normal growth and development of

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the A. amphitrite larvae.42 If the polymer particles are able to enter the algal cells, it offers a

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secondary route of entry of PMMA nanoparticles into the nauplii. Hence, it was necessary to

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ascertain the type of interaction between the individual algal species and the PMMA

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nanoparticles. This was carried out by exposing the algal cells with PMMA nanoparticles,

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incubating them for 24 hours and collecting the cells for imaging. The results were compared

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with untreated algal cells.

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The initial cell counts of Tetraselmis suecica and Chaetoceros muelleri used were (11.8 ± 1.3)

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× 104 and (10.3 ± 1.5) × 104 cells/mL, respectively. After 24 hours of incubation, no

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significant differences (Table S2) were observed between the mean final cell counts of

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Tetraselmis suecica and Chaetoceros muelleri for all three tested PMMA concentrations

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against the controls (ANOVA, p > 0.05, Figure 3). If PMMA particles had adhered onto the

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cell walls of the microalgae, it may have been possible to observe the adverse effect on the

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growth of microalgal cells in the media. Since we observed an almost similar number of

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microalgal cells in all samples (Figures 3A, 3B), irrespective of the PMMA particle

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concentration, it is likely that the polymer particles do not cause toxicity to algal cells. Algal

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cells exposed to nanoplastics showed no fluorescent polymer particles inside the cells (data 10 ACS Paragon Plus Environment

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not shown), thus implying that the addition of PMMA nanoparticles up to 25 ppm to

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microalgae culture had no significant impact on the proliferation, absorption of nutrients and

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photosynthetic functions of the microalgal cells. This may be explained by the characteristics

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of plant cells, where they are more selective in taking up materials from their surroundings

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due to the presence of a rigid cellulose cell wall.43 While there is evidence of small

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nanoparticles (< 50 nm) being able to permeate the cell wall of microalgae and plants, larger

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nanoplastic particles seem to be barricaded efficiently owing to their size and

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hydrophobicity.44-45

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Figure 3: Mean microalgae count observed after 24 hours of nanoplastic treatments of (A)

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Tetraselmis suecica and (B) Chaetoceros muelleri. The initial cell counts of the microalgal

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cells were (11.8 ± 1.3) × 104 and (10.3 ± 1.5) × 104 cells/mL respectively.

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Uptake and Translocation of PMMA Nanoplastics in Barnacle Nauplii: Barnacle nauplii

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are suspension filter feeders that can scavenge the water column for planktonic feed. The

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uptake of nanoplastics by the stage II nauplii was monitored at 45-minute intervals for an

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initial period of 3 hours after exposure to the nanoplastics at a concentration of 25 ppm

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(Figure S2). After the first 45 minutes interval, more than half of the nauplii (~50 – 60%) had

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ingested the nanoplastics with weak florescence signals appearing in the gut region. The

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signals were quantitatively compared with the control samples. The fluorescence intensity

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was 1200% higher than that of the control. After 90 minutes, the relative intensity of

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fluorescence from the nauplii gut had increased by a factor of 290%, as compared to the 45

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minute samples, and a larger proportion of nauplii (~75%) had ingested the nanoplastics. The

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fluorescence was seen to be localized in two major regions of the alimentary canal: the

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midgut and the hindgut of the nauplii body (Figure S4). A further 6% increase in relative

25

fluorescence intensity was observed after prolonged exposure to PMMA nanoplastics for 135 11 ACS Paragon Plus Environment


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minutes. All nauplii imaged exhibited prominent green fluorescence in the mid- and hindgut

2

regions of the body after 135 minutes of nanoplastic exposure. To ensure all nauplii had taken

3

up the nanoplastics, the nauplii were incubated with PMMA nanoparticles for an additional

4

45 minutes and images were collected.

5

In a study by Cole et al., the uptake of polystyrene micro- and nanoplastics was monitored in

6

oyster larvae post-fertilization.44 Their results showed that depending on the size of the

7

plastics, the larvae selectively ingest the plastic particles. Plastics smaller than 7.3 µm were

8

ingested by oyster larvae at all life stages whereas larger plastics (20 µm) were only ingested

9

by larvae 24 days post fertilization.46 Daphnia magna larvae were also exposed to fluorescent

10

microplastics and the uptake was monitored after 48 hours.19 Blue fluorescence from the D.

11

magna gut confirmed the uptake of the microplastics. Subsequent depuration studies were

12

conducted by introducing the exposed larvae into fresh media for 24 hours, and the changes

13

in larval fluorescence were further analyzed. While the results indicated that the plastics were

14

completely removed via defecation, it was impossible to confirm the complete removal of

15

microplastic particles due to the presence of autofluorescence from the larvae.18 Many such

16

short term studies on the uptake of microplastics in marine larvae highlight the presence of

17

plastic in the larval body, however to the best of our knowledge long term growth

18

experiments on such larvae have not been studied and analyzed.

19

When examining the fate of nanoplastics after ingestion by the nauplii, the distribution of

20

fluorescence signals inside the body was qualitatively monitored using fluorescence

21

microscopy (Figure 4). After 30, 75, and 105 minutes of initial exposure, the PMMA

22

nanoplastics were localized in the hindgut and midgut (Figure S5, S6, Figure 4A - D).

23

Interestingly, the ingested nanoplastics were distributed throughout the body of nauplii after

24

135 minutes of exposure (Figure 4E), suggesting that the nanoplastics had been absorbed and

25

retained in other parts of the body and tissues (Figure S6).


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Figure 4: Translocation and retention of nanoplastic particles at 0 (A), 30 (B), 75 (C), 105 (D), 135 (E) and 165 (F) minutes after initial uptake. Uptake exposure conditions: 25 ppm nanoplastics, 30 nauplii/mL, 3 hour exposure. Refer to Figure S5 for DIC and fluorescence only image.

6 7

The most striking result from this study was that even after 24 hours of incubation in clean

8

FSW, the PMMA particles were observed inside the nauplius (Figure S6, AVI). Further

9

evidence of the distribution of the nanoparticles inside the body and not just the surface of the

10

nauplius is shown in the AVI file (Supporting Information). In parallel, cellular studies

11

carried out using mammalian kidney cells (data not shown), PMMA nanoplastics were found

12

to permeate the cellular membrane in a span of 5 hours. Currently, we are establishing the

13

mechanism of entry of such particles across the cells and tissues. 13 ACS Paragon Plus Environment


1

Effects of Acute and Chronic Exposure of PMMA Nanoplastics on Barnacle Larvae

2

Development

3

The growth of barnacle larvae consists of three stages,47-48 and the duration in which the

4

larvae remain in each of these stages is mainly dependent on temperature and nutrients

5

availability.49 In order to understand the impact of nanoplastics on different life stages,

6

investigations on the distribution of nanoplastics inside barnacle larvae were conducted at

7

three-time points, stage III nauplii (2 days old), cyprid (5 days old) and juvenile (7 days old).

8

Three treatment conditions were analyzed at each time point: acute, chronic and control.

9

Acute exposure experiments were carried out to understand the persistence of nanoplastics

10

inside barnacles whereas chronic exposure experiments were conducted to simulate the

11

effects of environmentally significant concentrations of plastics over long term exposure.

12

Nauplii were exposed to 25 ppm nanoplastics solution for the duration of 3 hours as a one-off

13

acute exposure, followed by washing and culturing of the nauplii. The chronic treatment

14

involved continuous exposure of nanoplastics dosed at 1 ppm throughout the entire growth

15

cycle. The control batches of nauplii were reared in the absence of PMMA particles and the

16

data collected were used for comparison.

17

After 2 days, we found variations in the relative intensities of fluorescence among stage III

18

nauplii in the control, acute and chronic exposure treatment conditions (Figures 5A – C). The

19

larvae in control experiments with no nanoplastics, exhibited a low auto-fluorescence signal,

20

which could be attributed to the use of glutaraldehyde as fixing agent that lead to the

21

formation of Schiff’s base structure.50-51 However, the nauplii exposed to particles in acute

22

and chronic conditions exhibited significantly larger fluorescence signals, when compared to

23

controls (Table S3, S4).

24

After 5 days, the nauplii metamorphosed into the non-feeding cyprid form. Fluorescence

25

intensities from cyprids were the least in control, followed by acute and chronic exposures

26

(Figures 5D – F). In the control samples, fluorescence signals were detected only from the

27

outer carapace (shell). The distribution of fluorescence signals inside the cyprids differed

28

between the acute and chronic exposures. Cyprids with acute exposure exhibited

29

homogeneously distributed fluorescence across their bodies, with a high contrast in the

30

compound eye and internal appendages compared to its carapace (Figure 5E). Cyprids in the

31

chronic exposure showed similar distribution but stronger contrast defining the internal

32

organs, compound eyes and limbs (Figure 5F). The observed high contrast is the consequence 14 ACS Paragon Plus Environment

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1

of increased residence time of the nanoplastics inside the cyprid’s body. The fluorescent

2

intensity histograms (Figures 5D – F) and average fluorescence intensity values (Table S3)

3

also supported the presence of PMMA nanoplastics inside the cyprids. The relative

4

fluorescence intensities of control samples was observed to be significantly different than the

5

acute and chronic exposure (Table S4, ANOVA, p < 0.05).

6

After 7 days, cyprids suspended in clear FSW metamorphosed into settled juveniles. Again,

7

the intensity of observed fluorescence from juveniles was low for control and higher for those

8

exposed to PMMA particles under acute and chronic exposure conditions (Figures 5G – I,).

9

In the control sample, minimum amounts of auto-fluorescence were observed from localized

10

regions of the barnacle (Figure 5G). The mean fluorescence intensity of the samples indicated

11

that relative intensity of signal from chronic was the highest and control was the lowest

12

(Table S3). The intensity of fluorescence for the control samples was significantly different

13

from that of the acute and chronic exposed juvenile (Table S4, ANOVA, p < 0.05).

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16 15 ACS Paragon Plus Environment


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Figure 5: Confocal microscope images (10× magnification with a 2.2× zoom) of nauplius

2

after 2 days (A – C), cyprids after 5 days (D – F) and settled juvenile after 7 days (G – I)

3

imaged as controls (A, D, G), acute (B, E, H) and chronic (C, F, I) exposure. Scalebar: 100

4

µm, Inset: Relative intensity distribution of the green fluorescence. Exposure conditions:

5

Control – no nanoplastics, Acute – 25 ppm, 3 hour exposure, Chronic – 1 ppm, dosed every

6

two days.

7

Results showed that PMMA nanoplastics, specially during acute exposure (25 ppm, 3 hours),

8

remained within the bodies of barnacle larvae throughout their developmental cycle (Figure

9

5). Metamorphosis of barnacle larvae appeared to be unaffected by the presence of

10

nanoplastics, and that the particles became redistributed throughout the body. Even though,

11

the concentration of PMMA nanoparticles under chronic exposure conditions was twenty-

12

five-fold lower than the acute exposure, we found a significant build-up of nanoplastics

13

inside the larvae from chronic exposure, which yielded higher fluorescence signals. In

14

addition, common polyalkanes such as polystyrene, PMMA, polyethylene and polypropylene

15

have no known biodegradation pathways under in vivo conditions. This suggests that long-

16

term chronic exposure of such polymer nanoparticles results in bioaccumulation, which

17

potentially could cause more harm to tissues and the organisms, as compared to short-term

18

acute exposure.

19

We also found that the exposed larvae were able to remove some of the PMMA nanoplastics

20

via two methods: moulting and excretion of faecal matter throughout its lifecycle. Nauplii

21

samples treated with chronic exposure of PMMA nanoplastics were not examined in this

22

study. The continual exposure to nanoparticles makes it difficult to distinguish the

23

fluorescence particles found only in the moults and faecal matter (i.e. indication of ingestion

24

and egestion by barnacles) versus particles adsorbed onto these materials from the general

25

media. In the case of cyprid moults from nauplii exposed to acute conditions, PMMA

26

particles were found in both faecal matter and moults. The 3D z-stack micrographs of the

27

moult from acute exposure showed the presence of fluorescent nanoplastics in both the

28

nauplii and cyprid moults (Figures 6A – D). Naupliar moults of the acute exposed treatment

29

showed relatively higher fluorescence than their corresponding controls (Figure 6G). Similar

30

trends were also observed in the faecal matter of the acute and control (Figure 6I). The

31

relative fluorescence intensity of the control and acute exposed cyprids (Figure 6H) were

32

almost similar. This suggests that the nanoplastics excretion through moulting in the latter


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1

stages of the life cycle is less efficient. This could be attributed to the absorption and

2

retention of the plastics inside the barnacle body (as seen in the acute juvenile).

3

The more common route for excretion and removal of the nanoplastics is through defecation.

4

Optical images of the faecal matter collected from the larvae from the control showed no

5

fluorescence (Figure 6E) whereas after acute exposure, a distinct fluorescence was observed

6

(Figure 6F). The presence of PMMA nanoparticles in faecal matter implies that some of the

7

ingested nanoparticles are excreted without bioabsorption.

8 9 10

Figure 6: Possible nanoplastic egestion route via naupliar moult (A: control, B: acute), cyprid

11

moult (C: control, D: acute) and faecal matter (E: control, F: acute). (Scalebar 20×

12

magnification: 100 µm). The relative fluorescence intensity of the naupliar moult (G), cyprid

13

moult (H) and faecal matter (I) are also plotted as a function of position.

14

Our initial investigations showed that the particles in the faecal matter are intact, but further

15

studies are currently underway to evaluate changes in shape and size of the ingested particles,



1

if any. Results from this study suggest the possible natural pathways of excreting the

2

nanoplastics from the larvae’s bodies, but the nanoparticles are remained and bioaccumulated

3

inside the organism.

4

Conclusions

5

PMMA particles encapsulated with fluorescent PTE dye molecules are used for investigating

6

the intake, translocation and egestion by acorn barnacle larvae. Our experimental data

7

revealed that PMMA particles are non-toxic at concentrations as high as 25 ppm to barnacle

8

nauplii. The nanoplastics also had no significant interactions with algal cells, which are used

9

as feeds for barnacle larvae and did not affect their proliferation profile. Within 3 hours of

10

exposure, stage II nauplii had ingested nanoplastics and were retained inside the body

11

throughout all stages of larval development, which included metamorphosis and settlement

12

into the juvenile form. The ingested nanoplastics had also redistributed within the nauplii

13

body owing to its open circulatory system. Two modes of excretion of the nanoplastics were

14

also observed: via moulting and defecation.

15

Acute exposure at high concentrations (25 ppm, 3 hours exposure), and chronic exposure at

16

low concentration (1 ppm, continuous exposure), of nanoplastics to barnacle larvae led to an

17

accumulation of nanoplastics inside their bodies. Fluorescence signals from the ingested

18

polymer nanoparticles in the naupliar stage were also found to be persistent through the

19

subsequent cyprid and juvenile growth stages. The persistence of nanoplastics is expected in

20

the adult forms of barnacles, and this has wider implications on their growth and reproduction.

21

New studies focused on the interactions of nanoplastics with the later life history of barnacles

22

are currently underway to examine potential consequences.

23

ASSOCIATED CONTENT

24

Supporting Information

25 26

The Supporting Information is available free of charge on the ACS Publications website at DOI:

27 28

Absorbance and emission spectra of PTE dye, Confocal images of nauplii under different conditions, statistical analysis of the data, fluorescent intensity data.

29

■ AUTHOR INFORMATION

30

Corresponding Author

31

*E-mail: [email protected] (S. Valiyaveettil).

32

ORCID 18 ACS Paragon Plus Environment

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1

Suresh Valiyaveettil: 0000-0001-6990-660X

2

Notes

3

The authors declare no competing financial interest.

4 5

Acknowledgements: The authors acknowledge the funding support from the National

6

Research Foundation Singapore for research conducted under the Marine Science R&D

7

Program at NUS (R-143-000-676-281) and St. John’s Island National Marine Laboratory and

8

SB acknowledge Graduate Research Scholarship from the National University of Singapore.

9

References

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44

1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

11.

12.

13.

14.

15.

Jambeck, J. R.; Geyer, R.; Wilcox, C.; Siegler, T. R.; Perryman, M.; Andrady, A.; Narayan, R.; Law, K. L., Plastic waste inputs from land into the ocean. Science, 2015, 347 (6223), 768-771. UNEP Marine plastic debris and microplastics – Global lessons and research to inspire action and guide policy change. ; United Nations Environment Programme: Nairobi, 2016. Tanaka, K.; Takada, H., Microplastic fragments and microbeads in digestive tracts of planktivorous fish from urban coastal waters. Sci. Rep. - UK, 2016, 6, 34351. Mani, T.; Hauk, A.; Walter, U.; Burkhardt-Holm, P., Microplastics profile along the Rhine River. Sci. Rep. - UK, 2015, 5, 17988. Cole, M.; Lindeque, P.; Halsband, C.; Galloway, T. S., Microplastics as contaminants in the marine environment: A review. Mar. Pollut. Bull., 2011, 62 (12), 2588-2597. Wright, S. L.; Thompson, R. C.; Galloway, T. S., The physical impacts of microplastics on marine organisms: A review. Environ. Pollut., 2013, 178, 483-492. Rochman, C. M.; Hoh, E.; Kurobe, T.; Teh, S. J., Ingested plastic transfers hazardous chemicals to fish and induces hepatic stress. Sci. Rep. - UK, 2013, 3. Galloway, T. S.; Cole, M.; Lewis, C., Interactions of microplastic debris throughout the marine ecosystem. Nat. Ecol. Evol., 2017, 1, 0116. Steer, M.; Cole, M.; Thompson, R. C.; Lindeque, P. K., Microplastic ingestion in fish larvae in the western English Channel. Environ. Pollut., 2017, 226 (Supplement C), 250-259. Auta, H. S.; Emenike, C. U.; Fauziah, S. H., Distribution and importance of microplastics in the marine environment: A review of the sources, fate, effects, and potential solutions. Environ. Int., 2017, 102 (Supplement C), 165-176. Mizraji, R.; Ahrendt, C.; Perez-Venegas, D.; Vargas, J.; Pulgar, J.; Aldana, M.; Patricio Ojeda, F.; Duarte, C.; Galbán-Malagón, C., Is the feeding type related with the content of microplastics in intertidal fish gut? Mar. Pollut. Bull., 2017, 116 (1), 498-500. Karlsson, T. M.; Vethaak, A. D.; Almroth, B. C.; Ariese, F.; van Velzen, M.; Hassellöv, M.; Leslie, H. A., Screening for microplastics in sediment, water, marine invertebrates and fish: Method development and microplastic accumulation. Mar. Pollut. Bull., 2017, 122 (1), 403-408. Kanhai, L. D. K.; Officer, R.; Lyashevska, O.; Thompson, R. C.; O'Connor, I., Microplastic abundance, distribution and composition along a latitudinal gradient in the Atlantic Ocean. Mar. Pollut. Bull., 2017, 115 (1), 307-314. Cole, M.; Lindeque, P.; Fileman, E.; Halsband, C.; Galloway, T. S., The Impact of Polystyrene Microplastics on Feeding, Function and Fecundity in the Marine Copepod Calanus helgolandicus. Environ. Sci. Technol., 2015, 49 (2), 1130-1137. Cole, M.; Lindeque, P.; Fileman, E.; Halsband, C.; Goodhead, R.; Moger, J.; Galloway, T. S., Microplastic Ingestion by Zooplankton. Environ. Sci. Technol., 2013, 47 (12), 6646-6655. 19 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50

16. Lin, V. S., Research highlights: impacts of microplastics on plankton. Environ. Sci.-Proc. Imp., 2016, 18 (2), 160-163. 17. Bergami, E.; Bocci, E.; Vannuccini, M. L.; Monopoli, M.; Salvati, A.; Dawson, K. A.; Corsi, I., Nanosized polystyrene affects feeding, behavior and physiology of brine shrimp Artemia franciscana larvae. Ecotox. Environ. Safe., 2016, 123 (Supplement C), 18-25. 18. Booth, A.; Hansen, B.; Frenzel, M.; Johnsen, H.; Altin, D., Uptake and toxicity of methylmethacrylate-based nanoplastic particles in aquatic organisms: Ecotoxicity and uptake of nanoplastic particles. Environ. Toxicol. Chem., 2016; Vol. 35, p 1641-1649. 19. Chae, Y.; An, Y.-J., Effects of micro- and nanoplastics on aquatic ecosystems: Current research trends and perspectives. Mar. Pollut. Bull., 2017. 20. Miranda, D. d. A.; de Carvalho-Souza, G. F., Are we eating plastic-ingesting fish? Mar. Pollut. Bull., 2016, 103 (1), 109-114. 21. Desforges, J.-P. W.; Galbraith, M.; Dangerfield, N.; Ross, P. S., Widespread distribution of microplastics in subsurface seawater in the NE Pacific Ocean. Mar. Pollut. Bull., 2014, 79 (1), 9499. 22. AshaRani, P. V.; Low Kah Mun, G.; Hande, M. P.; Valiyaveettil, S., Cytotoxicity and Genotoxicity of Silver Nanoparticles in Human Cells. ACS Nano, 2009, 3 (2), 279-290. 23. Beer, C.; Foldbjerg, R.; Hayashi, Y.; Sutherland, D. S.; Autrup, H., Toxicity of silver nanoparticles—Nanoparticle or silver ion? Toxicol. Lett., 2012, 208 (3), 286-292. 24. Bundela, H.; Bharadwaj, V., Synthesis and characterization of hydroxyapatite-poly-(vinyl alcohol) based nanocomposites for their perspective use as bone substitutes. Polym. Sci. Ser. A., 2012, 54 (4), 299-309. 25. Chan, C. Y. S.; Chiu, J. M. Y., Chronic Effects of Coated Silver Nanoparticles on Marine Invertebrate Larvae: A Proof of Concept Study. PLOS ONE, 2015, 10 (7), e0132457. 26. Stensberg, M. C.; Wei, Q.; McLamore, E. S.; Porterfield, D. M.; Wei, A.; Sepúlveda, M. S., Toxicological studies on silver nanoparticles: challenges and opportunities in assessment, monitoring and imaging. Nanomedicine-UK, 2011, 6 (5), 879-898. 27. Burden, D. K.; Spillmann, C. M.; Everett, R. K.; Barlow, D. E.; Orihuela, B.; Deschamps, J. R.; Fears, K. P.; Rittschof, D.; Wahl, K. J., Growth and development of the barnacle Amphibalanus amphitrite: time and spatially resolved structure and chemistry of the base plate. Biofouling, 2014, 30 (7), 799-812. 28. van Dam, J. W.; Trenfield, M. A.; Harries, S. J.; Streten, C.; Harford, A. J.; Parry, D.; van Dam, R. A., A novel bioassay using the barnacle Amphibalanus amphitrite to evaluate chronic effects of aluminium, gallium and molybdenum in tropical marine receiving environments. Mar. Pollut. Bull., 2016, 112 (1), 427-435. 29. Gambardella, C.; Morgana, S.; Ferrando, S.; Bramini, M.; Piazza, V.; Costa, E.; Garaventa, F.; Faimali, M., Effects of polystyrene microbeads in marine planktonic crustaceans. Ecotox. Environ. Safe., 2017, 145 (Supplement C), 250-257. 30. Ali, U.; Karim, K. J. B. A.; Buang, N. A., A Review of the Properties and Applications of Poly (Methyl Methacrylate) (PMMA). Polym. Rev., 2015, 55 (4), 678-705. 31. Frazer, R. Q.; Byron, R. T.; Osborne, P. B.; West, K. P., PMMA: An Essential Material in Medicine and Dentistry. J. Long Term Eff. Med. Implants, 2005, 15 (6), 629-639. 32. Vollrath, A.; Schallon, A.; Pietsch, C.; Schubert, S.; Nomoto, T.; Matsumoto, Y.; Kataoka, K.; Schubert, U. S., A toolbox of differently sized and labeled PMMA nanoparticles for cellular uptake investigations. Soft Matter, 2013, 9 (1), 99-108. 33. Reisch, A.; Klymchenko, A. S., Fluorescent Polymer Nanoparticles Based on Dyes: Seeking Brighter Tools for Bioimaging. Small, 2016, 12 (15), 1968-1992. 34. Rosenbauer, E.-M.; Wagner, M.; Musyanovych, A.; Landfester, K., Controlled Release from Polyurethane Nanocapsules via pH-, UV-Light- or Temperature-Induced Stimuli. Macromolecules, 2010, 43 (11), 5083-5093.


Page 20 of 21

Page 21 of 21 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43

35. Akbulut, M.; Ginart, P.; Gindy, M. E.; Theriault, C.; Chin, K. H.; Soboyejo, W.; Prud'homme, R. K., Generic Method of Preparing Multifunctional Fluorescent Nanoparticles Using Flash NanoPrecipitation. Adv. Funct. Mater., 2009, 19 (5), 718-725. 36. Sengupta, S., Synthesis of Regioisomerically Pure 1,7-Dibromoperylene-3,4,9,10-tetracarboxylic Acid Derivatives. J. Org. Chem., 2014, 79 (14), 6655-6662. 37. Chai, C. L. L.; Teo, S. L. M.; Jameson, F. K. M.; Lee, S. S. C.; Likhitsup, A.; Chen, C.-L.; Rittschof, D., Loperamide-based compounds as additives for biofouling management. Int. Biodeter. Biodegr., 2014, 89 (Supplement C), 82-87. 38. López, D. A.; López, B. A.; Pham, C. K.; Isidro, E. J.; De Girolamo, M., Barnacle culture: background, potential and challenges. Aquac. Res., 2010, 41 (10), e367-e375. 39. Piazza, V.; Ferioli, A.; Giacco, E.; Melchiorre, N.; Valenti, A.; Del Prete, F.; Biandolino, F.; Dentone, L.; Frisenda, P.; Faimali, M., A standardization of Amphibalanus (Balanus) amphitrite (Crustacea, Cirripedia) larval bioassay for ecotoxicological studies. Ecotox. Environ. Safe., 2012, 79 (Supplement C), 134-138. 40. Neo, M. L.; Todd, P.; Lay-Ming Teo, S.; Chou, L., The effects of diet, temperature and salinity on survival of larvae of the fluted Giant clam, Tridacna squamosa. J. Conchol., 2013; Vol. 41, p 369376. 41. Canesi, L.; Ciacci, C.; Fabbri, R.; Marcomini, A.; Pojana, G.; Gallo, G., Bivalve molluscs as a unique target group for nanoparticle toxicity. Mar. Environ. Res., 2012, 76 (Supplement C), 16-21. 42. Rittschof, D.; Lai, C.-H.; Kok, L.-M.; Teo, S. L.-M., Pharmaceuticals as antifoulants: Concept and principles. Biofouling, 2003, 19 (sup1), 207-212. 43. Siegel, S. M., Chapter 1 - Constitution and Architecture in the Cell Wall. In The Plant Cell Wall, Pergamon: 1962; pp 1-40b. 44. Sun, D.; Hussain, H. I.; Yi, Z.; Siegele, R.; Cresswell, T.; Kong, L.; Cahill, D. M., Uptake and cellular distribution, in four plant species, of fluorescently labeled mesoporous silica nanoparticles. Plant Cell Rep., 2014, 33 (8), 1389-1402. 45. Li, X.; Schirmer, K.; Bernard, L.; Sigg, L.; Pillai, S.; Behra, R., Silver nanoparticle toxicity and association with the alga Euglena gracilis. Environ. Sci.: Nano, 2015, 2 (6), 594-602. 46. Cole, M.; Galloway, T. S., Ingestion of Nanoplastics and Microplastics by Pacific Oyster Larvae. Environ. Sci. Technol., 2015, 49 (24), 14625-14632. 47. Al-Aidaroos, A. M.; Satheesh, S., Larval development and settlement of the barnacle Amphibalanus amphitrite from the Red Sea: Influence of the nauplii hatching season. Oceanol. Hydrobiol. St., 2014, 43 (2), 170-177. 48. Jones, L. W. G.; Crisp, D. J., The larval stages of the Barnacle Balanus improvisus Darwin. Proc. Zool. Soc. Lond., 1954, 123 (4), 765-780. 49. Jamieson, C. D., The effects of temperature and food on naupliar development, growth and metamorphosis in three species of Boeckella (Copepoda:Calanoida). Hydrobiologia, 1986, 139 (3), 277-286. 50. Clancy, B.; Cauller, L. J., Reduction of background autofluorescence in brain sections following immersion in sodium borohydride. J. Neurosci. Meth., 1998, 83 (2), 97-102. 51. Schelkle, K. M.; Schmid, C.; Yserentant, K.; Bender, M.; Wacker, I.; Petzoldt, M.; Hamburger, M.; Herten, D.-P.; Wombacher, R.; Schröder, R. R.; Bunz, U. H. F., Cell Fixation by Light-Triggered Release of Glutaraldehyde. Angew. Chem. Int. Edit., 2017, 56 (17), 4724-4728.

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