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Analytical Methodswww.rsc.org/methods

ISSN 1759-9660

PAPERTetsuo Okada et al.Chiral resolution with frozen aqueous amino acids

Volume 8 Number 1 7 January 2016 Pages 1–224

Analytical Methods

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Wang, S. Ghosal, C. M. Rochman, M. Gassel and S. Wall, Anal. Methods, 2016, DOI:

10.1039/C6AY02396G.

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1

Novel Method for the Extraction and Identification of Microplastics

in Ocean Trawl and Fish Gut Matrices

Jeff Wagnera, Zhong-Min Wang

a, Sutapa Ghosal

a, Chelsea Rochman

bc, Margy Gassel

d, and

Stephen Walla

aCalifornia Dept. of Public Health, Environmental Health Laboratory Branch, Richmond, CA bUniversity of California, Aquatic Health Program, School of Veterinary Medicine, Davis, CA cUniversity of Toronto, Department of Ecology and Evolutionary Biology, St. George, Ontario, Canada dCalEPA Office of Environmental Health Hazard Assessment, Oakland, CA

Abstract

This work presents alternative extraction and analysis techniques to identify microplastics in the

environment. This study aims to address previously noted issues with methods that use

aggressive extraction treatments or optical microscopy identification techniques alone. Pulsed

ultrasonic extraction with ultrapure water was used to remove microplastics from fish stomachs

without dissolving the stomach tissues or microplastics. The technique is relatively simple and

minimizes issues with hazardous disposal and laboratory safety. Microplastics were

characterized using optical microscopy, scanning electron microscopy plus energy-dispersive x-

ray spectroscopy (SEM/EDS), Fourier transform infrared (FTIR) micro-spectroscopy, and

Raman micro-spectroscopy (RMS). These methods were demonstrated successfully on

laboratory fish exposed to reference microplastics and on ocean surface trawl and fish samples

taken from subtropical gyres. Polyethylene (PE), polypropylene (PP), and blended PE + PP

microplastics were detected in the stomachs of ocean-caught lanternfish, with the majority

consisting of PE. One nearly empty lanternfish stomach contained a long PE fiber that appeared

to block the digestive tract. Minor amounts of fat, proteins, and carbohydrates were detected by

FTIR on many microplastic surfaces. The Pacific Ocean trawl samples yielded similar plastic

compositions as the fish stomachs, plus one polystyrene particle. Of the 115 ocean particles

analyzed by FTIR (15 µm – 5 mm), 25 particles were microplastics (600 µm – 5 mm). The

microplastic PE + PP copolymer blends were the most visibly degraded of the four observed

types. FTIR and SEM/EDS identified micro-shell pieces in the ocean fish stomachs that

resembled microplastics by optical microscopy alone.

Revisions submitted to Analytical Methods, October 2016

Page 1 of 24 Analytical Methods

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View Article OnlineDOI: 10.1039/C6AY02396G


2

Introduction

Microplastics are anthropogenic pollutants that accumulate in marine and freshwater ecosystems

globally 1, both in the form of engineered particles in consumer products (e.g., microbeads in

personal care products) and degradation products from larger plastic products 2. Microplastics

are defined here as plastic particles <5 mm 3.

This work is part of a larger collaboration with United States Environmental Protection Agency

Region 9 to investigate microplastics ingested by ocean fish, characterize microplastic surfaces,

and determine any association between plastic debris and levels of persistent organic pollutants

in fish. This paper focuses primarily on method development.

The presence of microplastic is increasingly reported in more complex matrices, e.g., sediments

rich with organic matter, manta trawls filled with plant matter and woody debris, and biota of all

shapes and sizes 4. A variety of methods have been developed to measure microplastics in these

matrices. A key aspect of these analytical methods is the extraction of microplastics from

interfering biomass. Many previous studies have employed one or more fairly aggressive

chemicals (e.g., KOH, H2O2, HClO4, HNO3) to dissolve the biomass, which can be destructive to

the plastic particles and their surfaces. 5-7 Of the above chemicals, KOH was reported to cause

the least plastic damage 5. Enzymatic digestion methods have also been used to minimize

damage to plastics 8, 9. Still, even “plastic-safe” treatments have the potential to redeposit

dissolved tissue residues on plastic surfaces. Preliminary extractions for this work using KOH

(described in more detail below) created interferences that were problematic for micro-

spectroscopy-based analyses. These findings motivated the development of alternative

preparation and extraction methods.

Another analytical consideration involves the methods by which plastic types are identified.

Plastic identification methods include combustion techniques (CHN analyzers, pyrolysis-gas

chromatography/mass spectrometry) and various types of microscopy and micro-spectroscopy 8.

Studies that rely on visual identification alone, either unaided or using optical microscopy, are

vulnerable to misidentification of microplastics 8. Fourier transform infrared (FTIR) micro-

spectroscopy and Raman micro-spectroscopy (RMS) are two distinct but complementary

vibrational techniques which can be used for molecular identification of specific microplastic

types and non-plastic interferences 5, 8, 10

. Scanning electron microscopy plus energy-dispersive

x-ray spectroscopy (SEM/EDS) offers high resolution imaging with identification of chemical

elements 11, and has also shown promise for distinguishing between plastics and non-plastic

interferences 12.

In this work, California Department of Public Health’s Environmental Health Laboratory Branch

(EHLB) optimized methods to extract microplastics using non-chemical techniques, then identify

and characterize particles in terms of type, size, and morphology using a screening protocol

employing complementary optical microscopy, SEM/EDS, FTIR, and RMS. These methods

were developed on laboratory fish exposed to reference microplastics, and were subsequently

tested on ocean surface trawl and fish samples taken from subtropical gyres. For clarity, the

results are described primarily using FTIR data, with select, complimentary data from SEM/EDS

and RMS. The SEM/EDS and RMS results will be presented in more detail in separate papers.

Page 2 of 24Analytical Methods

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3

Experimental

Laboratory Fish Samples

For preliminary method development and evaluation of extraction methods, two sets of

freshwater Japanese medaka (Oryzias latipes) were selected from a laboratory population

maintained by the Aquatic Health Program at the University of California Davis (UCD). Care,

maintenance, handling, and sampling followed protocols were in accordance with UCD Animal

Care and Use Committee and approved by the UCD Animal Care and Use Committee. In the

laboratory, these fish were exposed to a mixture of several common microplastic types mixed in

with a purified casein fish diet (polyvinyl chloride (PVC), polyethylene (PE), polypropylene

(PP), polystyrene (PS), polyethylene terephthalate (PET)), and a mixture of anthropogenic fibers

from laundry lint. The plastics were donated as pellets by SABIC Innovative Plastics (Mt

Vernon, IN, USA) and micronized by Custom Processing Services (Reading, PA, USA) into

particles 100 – 2,000 µm in size. The lint was sampled from a clothes dryer run with a mix of

typical household laundry. Fish were placed into 2L beakers. Each type of microplastic was

added in small scoops with a laboratory spatula. Because a mass balance was not desired, the

concentrations were not quantified, but relatively high amounts were used to assure the fish ate

the plastic.

Clean microplastic particles were analyzed by SEM, FTIR, and RMS as reference samples.

These results were compared to the results obtained from particles extracted from the medaka

gastrointestinal (GI) tracts.

One set of medaka GI tracts was dissolved in 10% KOH solution, made with 10 g KOH in 100

mL of ultrapure H2O (purified, deionized, and filtered through a 0.1 µm membrane with a

Millipore Milli-Q system.) For initial analyses, particles were transferred directly to microscopy

substrates using a pipet. For subsequent analyses, KOH aliquots were filtered onto 0.8 µm pore

size PC (Millipore polycarbonate membrane) filters. To prevent damage to the PC filters, 10%

HNO3 was added to the KOH solution to create a neutralized (pH=7) solution prior to filtration.

The second set of medaka GI tracts was received frozen in aluminum foil. For this set, instead of

using chemicals to dissolve biota, an alternative extraction method was developed to remove

ingested particles from fish stomachs while minimizing tissue dissolution and re-deposition on

microplastic surfaces. First, the medaka GI tracts were removed from freezer storage, thawed,

and immediately dissected with a clean stainless steel scalpel blade in a clean glass petri dish

under a stereozoom microscope (see below for microscope details). Any observed particles that

visually resembled microplastics were set aside in another clean glass petri dish for SEM and

vibrational micro-spectroscopy analysis (Fig. 1a). Each dissected GI tract was then rinsed with

room temperature, ultrapure H2O through a glass funnel into a 40 mL glass vial. The GI tract was

subsequently processed with pulsed ultrasonic extraction (PUE) using a Ney Prosonik controller

and water tank (40-PRO-0506N). The PUE process utilized a series of square envelope bursts

modulated by a 39 – 41 KHz sweep wave form, with quiet time between bursts and de-gas time

between burst trains (Fig. 1b). The sealed vial was mounted in the PUE water tank with the vial


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cap above the waterline. PUE was conducted for six minutes at a burst power of 360 watts. The

vial contents were then poured immediately through a 1mm stainless-steel sieve mounted in a

glass funnel onto a 47 mm, 10 µm pore-size PC filter mounted inside a polysulfone (Millipore

Nalgene Sterafil) filtration unit. A backing filter (2 µm PTFE Zefluor) was used to assure

uniform filter deposition onto the PC filter. Stomach tissue and any other particles >1mm were

rinsed off of the stainless steel sieve into a clean glass Petri dish for analysis. The <2 µm

ultrapure H2O suspension was collected downstream of the filters in a glass jar and archived

along with the 2 µm backing filter for potential, future microplastics analysis.

Ocean Surface Trawl and Fish Samples

Extraction and analytical methods were further developed and verified using samples from ocean

surface manta trawls and the stomach contents of lantern fish from the family Myctophidae.

Three surface manta trawl samples and six myctophid fish were selected from a manta trawl

collected in August 2009 during a Project Kaisei research cruise in the North Pacific Ocean 13.

Seven additional myctophid fishes were selected from fish collected via manta trawl on a sailing

trip led by 5Gyres across the South Atlantic Ocean, November-December 2010 14. The average

length and mass of the analyzed Pacific fish were 9.2 +/- 0.5 cm and 7.5 +/- 0.9 g, respectively

(mean and standard deviation). The average length and mass of the analyzed Atlantic fish were

9.3 +/- 1.3 cm and 12 +/- 3.9 g, respectively.

Pacific Ocean surface trawl samples were received frozen in large glass jars sealed with

polytetrafluoroethylene (PTFE) lined screw top lids. For each, jar contents were rinsed with

approximately 100 mL ultrapure H2O through a 1mm stainless steel sieve (Gilson) mounted in a

glass funnel (Fig. 2). Particles >1 mm were rinsed from the sieve with ultrapure H2O and

collected for analysis in a pre-cleaned glass Petri dish. The sieved <1mm fraction was collected

in a pre-cleaned glass jar, from which three 15 mL subsamples were drawn. The three parallel

subsamples were collected on 25 mm, 10 µm pore-size PC filters mounted in glass vacuum

filtration units, one each for SEM, FTIR, and RMS analysis. The jar was swirled to homogenize

the sample before each filtration to obtain equivalent subsamples. The <10 µm fractions

collected downstream of the 3 filters were subsequently filtered through a 25 mm, 0.1 µm pore-

size PC filter to identify any smaller microplastics or colloidal materials that may have been

adhered to the larger trawl particles.

Myctophid GI tract samples from the Pacific Ocean were received frozen in aluminum foil.

Atlantic Ocean myctophid GI tract samples were received frozen and embedded in optimum

cutting temperature (OCT) media on cork substrates, along with a set of OCT media blanks.

OCT media had been used because the samples were originally prepared for a different analysis

that required cryosectioning. For all myctophids, only the stomach was processed to minimize

the introduction of decomposed foil or OCT artifacts, observed in some cases to be intermixed

with the other organs. The myctophid stomachs were dissected and processed using the same

ultrapure H2O-PUE protocol developed for the laboratory fish (see previous section).

Cleaning and Quality Control


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Glass Petri dishes, funnels, sieves, and Nalgene filtration units were cleaned between filtrations

using four sequential treatments: 1) scrub with detergent and DI H2O, 2) immerse in DI H2O in

an ultrasonic bath (Branson 3510) for 15 minutes, 3) repeat step 2 with fresh DI H2O, and 4)

rinse with ultrapure H2O. The 50 mL glass vials (ESS) and glass jars (I-Chem) with PTFE screw

cap liners were single-use, and certified contamination free by the vendors. Scalpel blades and

tweezers were rinsed with isopropyl alcohol (Baker Analyzed ACS Reagent Grade 2-Propanol)

and inspected under the stereozoom microscope for cleanliness prior to each use. The use of

laboratory wipes was prohibited during this step to minimize fiber contamination. Ultrapure

water and isopropyl alcohol were administered using dedicated PE wash bottles, as no suitable

non-plastic alternative was identified for this purpose. All filtrations, cleaning, and sample

preparations were performed using powder free, disposable nitrile gloves.

Typically, a laboratory blank was prepared simultaneously along with 1-2 sample filtrations.

Blanks were analyzed for the presence of plastics due to improper cleaning, as well as any

degradation of the polysulfone funnels, PE squeeze bottles, PTFE backing filters or cap liners, or

nitrile gloves. OCT media blanks were analyzed to account for any contamination of particles

from the Atlantic fish.

All filtrations and drying of cleaned filtration equipment were conducted inside particle-free

ventilation hoods with HEPA filtered air curtains, located within a cleanroom rated at class 2k

(less than 2,000 particles > 0.5 µm per cubic foot). Glassware and filtration unit cleaning was

also performed in the clean room. Monitoring of the cleanroom and hoods with an optical

particle counter (LasAir II, Particle Measuring Systems) typically yielded no detectable airborne

particles > 0.5 µm per cubic foot.

A representative filter subsampling approach 15, 16

was used to measure presence/absence of

microplastic types per trawl or per fish stomach. Because the accuracy of this approach depends

upon homogeneous filter loadings, filter deposit uniformity was verified under the stereozoom

microscope in all cases. Separate subsamples were obtained for each analytical technique.

Randomly selected, 1 cm2 squares were cut from each PC filter with a clean scalpel blade.

Particles were selected randomly from each filter subsample for analysis.

Optical Microscopy

Two reflected-light stereozoom microscopes (Leica S8APO and S6D) with digital CCD cameras

(Leica DFC 420 and 320) were used for initial documentation, gut dissection, and pre-screening

of any observed particles that visually resembled microplastics. Optical microscopy was

conducted at 6.3x – 80x with a minimum resolution of approximately 10 µm.

SEM/EDS

SEM/EDS was conducted using an FEI XL30 Environmental SEM and a Thermo Fisher

Scientific Noran System 7 EDS System to screen for potential microplastics using particle

morphology and elemental chemistry profiles.


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Pipetted particles from the first medaka gut set were deposited on copper tape mounted on

aluminum stubs. All subsequent individual particle and PC filter subsamples were prepared for

SEM/EDS by mounting them on double-sided adhesive carbon tabs on aluminum stubs.

To minimize sample charging under the electron beam, 0.6 mBar of water vapor was injected

into the SEM chamber for wet-mode imaging. When desired, this technique enabled artifact-free

analysis of the same subsample by FTIR or RMS, unlike other alternatives such as metal or

carbon-coating. Samples were imaged at 50x – 10,000x using an atomic number sensitive, back-

scattered electron (BSE) detector with a resolution of approximately 0.1 µm.

FTIR Micro-spectroscopy

FTIR microscopy was performed on individual, 15 µm – 5 mm particles using a Thermo

Scientific Nicolet Continuum FTIR Microscope with 10x optical objective, 15x IR objective, and

a Nexus 470 spectrometer. Depending on the sample, particles were analyzed via bulk attenuated

total reflectance (ATR), micro-ATR, transmission, or reflection techniques. The majority of the

particles were analyzed with a diamond micro-ATR microscope objective, either on glass slides

or directly on PC filter subsamples prepared on SEM stubs as described in the preceding section.

Selected larger particles were analyzed with a Smart Miracle bulk ATR with ZnSe crystal.

Transmission-FTIR microscopy was performed on a subset of the smaller particles, which were

positioned alongside KBr reference crystals inside a diamond compression cell. Reflection-FTIR

microscopy was conducted on selected smaller particles using mirrored, aluminum-coated slides.

All acquired FTIR spectra were compared to quantitative matches from EHLB’s in-house and

commercial (Thermo Scientific) FTIR libraries of over 100,000 spectra. Additional spectral

interpretation was often necessary when additional surface species were present.

Particle sizes were measured with the 10x optical objective, calibrated with a National Physical

Laboratory (UK)-certified, reflected-light stage micrometer (SPI Supplies). For non-fiber

spheroids, representative particle sizes were calculated using the average of the length and width.

The effective size, deff, for fibers in an oriented fluid flow was calculated using the following

expression 17, 18

:

deff = W × (9/4 × ρf/ρ0 × [ln(2L/W) - .807])0.5

(1)

where W = fiber diameter, ρf = fiber density, ρ0 = unit density, and L = fiber length.

Raman Micro-spectroscopy

RMS analyses were performed using both Renishaw inVia and Bruker Senterra dispersive

Raman microscopes with 785 nm and 532 nm lasers. Several different objectives (5x, 20x, 50x

and 100x) were used to optimize the analytical laser spot size for spectral analyses. To minimize

laser-induced damage of the sample, laser power and acquisition times were varied depending on

sample sensitivity to thermal damage. Data analyses utilized spectral matching performed using

GRAMS software, which is equipped with commercially available Raman spectral libraries 19.


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Automated spectral matching using this software package was accomplished using a correlation

based search algorithm.

Results

Laboratory Fish Samples

Analyses of microplastics from the first set of medaka GI tracts dissolved in 10% KOH often

revealed re-deposited gut tissue residues and KOH reaction products. The filter preparations

(Fig. 3a) generally showed less residues than the pipetted particle preparations (Fig. 3bc), but for

both types of KOH preparations, FTIR of the microplastic surfaces detected strong peaks from

proteins, fats, and potassium salts (Fig. 3ab) not present in the original reference plastics.

SEM/EDS of microplastics from the KOH preparations revealed fine particulate coatings and

strong potassium peaks (Fig. 3c).

In contrast, the ultrapure H2O - PUE-extracted medaka GI tracts exhibited much cleaner surfaces

(Fig. 3d). All of the 28 particles extracted from these GI tracts were identified as microplastics

by FTIR, including a PET fiber from the laundry lint. In all cases, the plastic peaks and

morphologies of these particles matched those of the reference microplastic particles closely,

suggesting that the plastics were not damaged by the PUE extraction. Figure 3d shows data for

four ingested microplastic types: PE, PS, PVC, and PET. The FTIR-analyzed microplastics

ranged from 100-300 µm, with a mean and standard deviation of 165 +/- 64 µm.

Particles > 500 µm in size (corresponding to some of the PE and all of the PP particles) were not

observed in these Medaka GI tracts. The observed absence of larger particles may be related to

the mouth sizes or feeding behavior of these laboratory fish, though this result cannot be easily

extrapolated to fish in other environments.

RMS analyses of parallel subsamples confirmed these results. SEM/EDS of the PVC particles

confirmed a strong Cl peak, in addition to the C peaks present in all types.

Manta Trawl Samples

Of the 46 trawl particles analyzed by FTIR, 20 were identified as microplastics (Fig. 4).

Analyzed particle sizes ranged from 0.25 mm – 5.0 mm (mean and standard deviation = 1.5 +/-

1.5 mm). The identified microplastic sizes ranged from 0.7 mm – 5 mm (mean and standard

deviation = 2.5 +/- 1.3 mm). The identified microplastic types were PE, PP, PE + PP copolymer

blends, and PS. A separate set of trawl particles examined by RMS also identified a similar set of

polymers. The majority of the plastic particles were PE (70%) and > 1mm. Phthalocyanine dye

was also detected in one bright blue PE particle, exhibiting peaks in the 1090-1200 cm-1 region

that were distinct from the PE peaks. The three PE+PP particles were the most visibly degraded

of the 4 observed types; two of the PE particles were so brittle that they fractured after contact

with the FTIR ATR objective (Fig. 4c). The remaining 26 trawl particles were identified as

various other marine constituents: cellulose (plant matter), chitin, sodium phosphate, silicates


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(sand and clays), proteins, and carbohydrates. Laboratory blanks exhibited no plastics in all

cases.

SEM/EDS screening for microplastic particles enabled quick differentiation from similarly

shaped, non-plastic, mineral-based marine particles. The mineral particles exhibited a much

brighter BSE signal due to the dominant inorganic elements, primarily Si and Ca..

Analyses of microplastic surfaces revealed adhered mineral crusts, fish scales, radiolarians,

crustaceans, and other microorganisms (Fig. 4b). In addition, a thin biofilm coating was

identified by FTIR on the surface of many of the microplastics, composed primarily of proteins,

carbohydrates, and clay. A similar biofilm with closely matching FTIR spectra was the main

constituent of the <10 µm filtrate from one of the trawl samples. SEM/EDS of this <10 µm

fraction confirmed an agglomeration of fine colloidal particles exhibiting C, O, Al, and Si.

Ocean Fish Samples

The PUE extraction method yielded gut particles that were adequately free of tissue residues for

microscopy and micro-spectroscopy analyses. All dissected stomachs retrieved from the >1 mm

sieve fraction were observed to be intact, with their previous contents visibly removed by the

PUE. Figure 5 shows a typical pre-dissection stomach, the stomach following dissection, and the

relatively clean stomach tissues after PUE treatment. Laboratory blanks exhibited no plastics in

all cases.

Of the 69 particles analyzed by FTIR across all 13 myctophids from both oceans, 5 particles

were identified as microplastics. These results are summarized in Table 1. For all particle types

in the Atlantic fish, the mean and standard deviations of the analyzed particle sizes in the > 1 mm

and 10 µm – 1mm fractions were 1.6 +/- 1.3 mm and 99 +/- 100 µm, respectively, with an

overall mean and standard deviation of 330 +/- 730 µm. The mean analyzed particle sizes from

the Pacific fish in the > 1 mm and 10 µm – 1mm fractions were 1.9 +/- 1.9 mm and 210 +/- 190

µm, respectively, with an overall mean and standard deviation of 500 +/- 940 µm.

For the Atlantic Ocean myctophids, one particle in one stomach was identified as plastic by

FTIR. This yields a prevalence of 1/7 Atlantic fish containing plastics (14%). The plastic was a

13 mm long fiber with a 0.2 mm diameter, composed of PE (Fig. 6). This fiber is too large to be

classified as a microplastic based on its length (> 5mm). However, the smaller fiber diameter (<<

5 mm) may be almost as important, as it likely enabled the fiber to enter the relatively small

stomach opening (Fig. 6a). The high aspect ratio of this fiber likely caused it to become lodged

in the stomach, somewhat analogous to asbestos fiber deposition in airways 18. The effective

size of the fiber calculated with Equation 1, 0.6 mm, reflects this preferential orientation

assumption. The fiber appeared to block most of the stomach length, and virtually no other

additional stomach contents were present. RMS analysis of a different subsample from this fiber

also identified it as PE (Fig. 6c). The PE library spectra in Figures 6 and 7 include both HDPE

and LDPE matches, but rigorous differentiation between the two compounds was not performed,

and all were interpreted as simply PE.


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For the Pacific Ocean myctophids, 4 particles in two fish were identified as microplastics by

FTIR (Fig. 7). This yields a prevalence of 2/6 Pacific fish containing microplastics (33%). The

most prevalent type was again PE (2 particles), followed by one each consisting of PP and PE +

PP. The identified microplastics ranged from 0.8 mm – 4 mm (mean and standard deviation = 1.6

+/- 1.6 mm).

Adsorbed species were detected by FTIR on some of the analyzed particle surfaces. Fats and

“ocean particulate matter” colloids (proteins, carbohydrates, and clays, as described above in the

Pacific Ocean trawl <10 µm fraction) were detected on many of the microplastic surfaces (Fig.

7). In rare cases, OCT media peaks were detected (Fig. 6b).

One of the most prevalent types of non-plastic particles in the myctophids were whole shells and

shell fragments, with the largest shell pieces observed in the Atlantic fish (Fig. 8). The analyzed

shell particles ranged from 25 µm to 3 mm in size, and were clear or white in color. SEM/EDS

exhibited strong calcium peaks, bright BSE signal, and grooved surfaces, while FTIR yielded

good matches to aragonite and calcite, all of which clearly identified these particles as mineral

shells. The stomachs of the Pacific Ocean fish also contained many fish scales (collagen plus

hydroxylapatite). Other constituents of both Atlantic and Pacific myctophids were undigested

crustaceans (proteins plus chitin), and various mixtures of triglycerides, protein + carbohydrates,

fatty or amino acid salts, cellulose, and silicates (sand, clay, and talc).

Discussion

This work focused primarily on method development, employing low numbers of fish compared

to sampling guidelines for contaminant monitoring programs 20-21

. In addition, no attempt was

made to measure the fraction of trawl mass analyzed, microplastics per m2 ocean surface,

fractions of filter areas analyzed, or total numbers of microplastics per fish. Nevertheless, the

observed prevalence of PE and PP particles in these Pacific fish, Atlantic fish, and Pacific trawl

samples were consistent with other studies. 22-23

In addition, the trend in plastic type prevalence

identified by FTIR in the Pacific fish guts (50% PE, 25% PE+PP, and 25% PP) was roughly

consistent with that in the Pacific trawls (70% PE, 15% PE+PP, 10% PP, and 5% PS). The

observed coherence of microplastics results observed by SEM, FTIR, and RMS techniques was

encouraging. However, these techniques should be regarded as complementary rather than

directly comparable due to their differing measurement principles 8, as well as the different

subsamples analyzed by each.

Although KOH extraction methods can be optimized to yield satisfactory results 5, 7, the

alternative methods pursued here offer some potential benefits. The ultrapure H2O-PUE

extraction technique minimizes hazardous disposal and lab safety issues compared to methods

that employ chemicals. The time required for the relatively simple H2O-PUE extraction is short

(< 1 hr) compared to multi-step chemical and enzymatic treatments (typically on the order of

several hours to several weeks 5, 7, 9

).

In this study, the use of multiple identification techniques and instruments required additional

laboratory time and resources, but their consistent findings together provided added confidence


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10

in the extraction methods developed in this study. The time/labor costs of each of the individual

analysis techniques alone were roughly proportional to their power to identify microplastics

(stereozoom requiring the least time, followed by SEM, followed by Raman and FTIR). Their

individual capital costs scaled with spatial resolution (stereozoom lowest, SEM highest.)

The remnants of OCT media detected by FTIR in the Atlantic Ocean fish samples suggest that

the substance was not completely removed through water extraction. The presence of OCT was

thus somewhat undesirable for these analyses, though its successful identification (via reference

to OCT blanks) minimized the impact of this intermittent interference. The other gut preservation

technique used in this study, wrapping in aluminum foil, also generated artifacts. Reflective

aluminum fragments were observed under the stereozoom microscope in some of the Pacific fish

samples, though these fragments were easily distinguishable from plastics using SEM, FTIR, and

RMS. Foil may degrade less in other samples that are frozen for shorter durations than these fish

(6 years).

FTIR was used to identify non-plastic particles as small as 15 µm and microplastics as small as

100 µm in laboratory fish, though the smallest microplastic detected in ocean fish was

considerably larger, 600 µm. Hypothetically, if smaller microplastics were less prevalent in

these ocean fish than larger ones, these FTIR analyses may not have had enough statistical power

to detect them. In addition, if the biological films observed in ocean fish guts coated smaller

particles more heavily than larger ones, plastics < 600 µm may have been more difficult to detect

by FTIR.

Many of the shell fragments observed in the ocean fish stomachs (25 µm - 3 mm) visually

resembled microplastics under the stereozoom microscope. Conversely, the brittle, angular PE

particles observed in the ocean trawl sample (Fig. 4c) could conceivably be mistaken for shell

particles using optical techniques. Together, these results suggest that studies that employ optical

detection alone may be prone to either false positives or false negatives.

Conclusions

Alternative microplastic extraction methods employing ultrapure H2O, PUE, sieving, and

filtration successfully isolated stomach contents while leaving the stomach tissues largely intact.

As a result, the majority of microplastics were found to be adequately free of tissue remnants for

microscopy and micro-spectroscopy analyses. Equally important, comparisons between extracted

microplastics from laboratory fish and clean reference particles suggest this method did not

dissolve or alter the properties of the microplastics themselves.

A combination of optical microscopy, SEM, FTIR, and RMS was used to detect microplastics in

ocean trawls and fish prepared using these methods. SEM/EDS was a useful screening tool for

identifying potential microplastics and ruling out mineral species confounders. FTIR identified

PE, PP, blended PE + PP, and PS microplastics in these ocean trawl and fish samples. FTIR and

SEM/EDS identified micro-shell pieces in the ocean fish stomachs, confirming that caution is

required in analyses that rely on visual methods alone.


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11

Acknowledgments

Funding for laboratory work was provided by United States Environmental Protection Agency

Region 9. Rochman was funded by a David H. Smith Postdoctoral Fellowship. We thank 5Gyres

and Project Kaisei for making it possible to collect myctophids from the S. Atlantic and N.

Pacific, respectively, and the Aquatic Health Program at UC Davis for providing Japanese

medaka. We also thank Anna-Marie Cook and Harry Allen of United States Environmental

Protection Agency Region 9 and Swee J. Teh at UC Davis for their advice on this work.

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Tables

Table 1. Summary of Ocean myctophid stomach particle FTIR analyses.

Ocean

# of fish

analyzed

by FTIR

Number of fish

containing

microplastics (percent)

# of

analyzed

particles

size range of

analyzed

particles (um)*

size range of

plastic particles

(um)*

identified

plastic types

Atlantic 7 1 (14%) 52 15 – 3,000 570 PE

Pacific 6 2 (33%) 17 35 – 4,000 750 – 4,000 PP, PE, PP+PE

*for spheroids: deff = (L+W)/2; for fibers: deff = W × (9/4 × rf/r0 × [ln(2L/W) - .807])0.5


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14

Figure Captions

Figure 1. Fish gut microplastic extraction method. (a) Manual large particle removal from

dissected gut, Pulsed Ultrasonic Extraction (PUE) of stomach contents, 1 mm sieving, and 10 µm

filtration to generate indicated fractions. (b) PUE modulated wave form with 2.8 ms bursts, 1 s

sweep from 39 – 41 KHz, 1 s burst train duration, 2.8 ms of quiet time between bursts, and 0.66 s

degas time between burst trains. Adapted from Puskas and Ferrell 24.

Figure 2. Extraction method for ocean surface trawl microplastics. Initial1 mm sieve collection is

followed by filtration to yield 1 mm – 10 µm fractions for different analysis methods, and an

archived 10 µm – 0.1 µm fraction to investigate the potential for smaller microplastics.

Figure 3. Microplastic particles extracted from laboratory medaka GI tracts. For all FTIR spectra,

extracted microplastics (red) are compared to the original reference microplastics (blue.) a) 150

µm PVC and b) 300 µm PET particles prepared in 10% KOH, showing strong FTIR peaks for

proteins and fats (red arrows) and potassium salts (green arrows). c) SEM/EDS of KOH-treated

microplastic showing redeposited particulate material and strong potassium peak. d) 150 µm PS,

150 µm PE, 250 µm PET, and 150 µm PVC extracted with ultrapure H2O and PUE, exhibiting

reduced FTIR protein and fat peaks and no salt peaks.

Figure 4. Stereozoom images of Pacific Ocean trawl microplastic particles, some with adhered

crustaceans, mineral crusts, or radiolarians. Visibly degraded PE+PP particles marked with

arrows in (b). (c) shows brittle PE microplastics; the left particle fragmented upon FTIR ATR

contact.

Figure 5. 10x stereozoom images of an Atlantic myctophid stomach preparation: a) pre-

dissection, b) post-dissection showing stomach contents, and c) post-PUE showing stomach

tissues devoid of stomach contents.Figure 6. a) Stereozoom (STZ) images, b) FTIR and c) RMS

spectra showing consistent results for 13 x 0.2 mm PE fiber extracted from Atlantic Ocean

myctophid stomach. Arrows in (b) denote minor peaks corresponding to OCT.

Figure 6. a) Stereozoom (STZ) images, b) FTIR and c) RMS spectra showing consistent results

for 13 x 0.2 mm PE fiber extracted from Atlantic Ocean myctophid stomach. Arrows in (b)

denote minor peaks corresponding to OCT.

Figure 7. FTIR spectra and stereozoom images from Pacific myctophid stomach microplastics: a)

800 µm PE, b) 4 mm PP, c) 750 µm PE d) 750 µm PE+PP. Red arrows denote minor peaks

corresponding to proteins, fats, and carbohydrates. Talc (blue peak) is a common plastic additive.

Figure 8. Broken shell pieces from Atlantic Ocean myctophid guts. a) stereozoom (STZ) image

of 2 mm shell in gut, b) SEM image of 600 µm fragment, c) FTIR microscope image of 100 µm

fragment, and d) FTIR spectrum from particle in (c) matching aragonite plus calcite.


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Figure 1

194x236mm (300 x 300 DPI)


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Figure 2

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Figure 3abcd

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Figure 4

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Figure 5

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Figure 6

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Figure 7

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Figure 8

115x80mm (300 x 300 DPI)


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Graphical Abstract for Table of Contents

This alternative microplastic extraction method employs ultrapure water, ultrasonication, and

identification using complementary optical microscopy, SEM/EDS, FTIR, and RMS techniques.


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39x19mm (300 x 300 DPI)


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