Sampling, isolating and identifying microplastics
ingested by fish and invertebrates†
A. L. Lusher,
*
a
N. A. Welden,
b
P. Sobral
c
and M. Cole
d
Microplastic debris (<5 mm) is a prolific environmental pollutant, found worldwide in marine, freshwater and
terrestrial ecosystems. Interactions between biota and microplastics are prevalent, and there is growing
evidence that microplastics can incite significant health effects in exposed organisms. To date, the
methods used to quantify such interactions have varied greatly between studies. Here, we critically
review methods for sampling, isolating and identifying microplastics ingested by environmentally and
laboratory exposed fish and invertebrates. We aim to draw attention to the strengths and weaknesses of
the suite of published microplastic extraction and enumeration techniques. Firstly, we highlight the risk
of microplastic losses and accumulation during biotic sampling and storage, and suggest protocols for
mitigating contamination in the field and laboratory. We evaluate a suite of methods for extracting
microplastics ingested by biota, including dissection, depuration, digestion and density separation. Lastly,
we consider the applicability of visual identification and chemical analyses in categorising microplastics.
We discuss the urgent need for the standardisation of protocols to promote consistency in data
collection and analysis. Harmonized methods will allow for more accurate assessment of the impacts
and risks microplastics pose to biota and increase comparability between studies.
1 Introduction
Over the past century there has been an exponential increase in
plastic demand and production.
1
Concurrently, improper
disposal, accidental loss, and fragmentation of plastic mate-
rials, have led to an increase in tiny plastic particles and bres
(microplastic, <5 mm) polluting the environment.
2,3
Micro-
plastics have been observed in marine,
4
freshwater
5,6
and
terrestrial
7
ecosystems across the globe, and biotic interactions
are widely evidenced (Fig. 1). Microplastics can be consumed by
a diverse array of marine organisms, across trophic levels,
including protists,
8
zooplankton,
9–17
annelids,
18–26
echino-
derms,
27–31
cnidaria,
32
amphipods,
19,26,33
decapods,
34–41
isopods,
42
bivalves,
43–60
cephalopods,
61
barnacles,
62
sh,
58,66–94
turtles,
95
birds
96
and cetaceans.
97,98
Over 220 different species
have been found to consume microplastic debris in natura. Of
these, ingestion is reported in over 80% of the sampled
populations of some invertebrate species.
34,38,41
Interactions
between microplastics and freshwater invertebrates, sh and
birds are increasingly reported
99–107
although some researchers
are focussing on model species such as Daphnia magna.
108–111
The consumption of microplastics by terrestrial organisms is
poorly documented, however, laboratory studies indicate
earthworms ( Lumbricus terrestris) can consume plastic particles
present in soil.
112
There are a number of exposure pathways by which organ-
isms may interact with microplastic debris. Direct consumption
of microplastic is prevalent in suspension feeders, including
Fig. 1 Publication trend of studies investigating biota interactions with
microplastics until 30th June 2016.
a
Department of Animal Ecology I, University of Bayreuth , Universitaetsstr. 30, 95440
Bayreuth, Germany. E-mail: amy.lusher7@gmail.com
b
Faculty of Science, Technology, Engineering and Maths, Open University, Milton
Keynes, MK7 6AA, UK
c
MARE – Marine and Environmental Sciences Centre, Faculdade de Ci
ˆ
encias e
Tecnologia, Universidade NOVA de Lisboa, Campus da Caparica. 2829-516
Caparica, Portugal
d
College of Life and Environmental Sciences: Biosciences, University of Exeter, Geoffrey
Pope Bui lding, Stocker Road, Exeter, EX4 4QD, UK
† Electronic supplementary information (ESI) available. See DOI:
10.1039/c6ay02415g
Cite this: Anal. Methods,2017,9,1346
Received 27th August 2016
Accepted 22nd October 2016
DOI: 10.1039/c6ay02415g
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zooplankton,
10
oysters
59
and mussels,
43–58,60–63
and deposit
feeders, such as sea cucumbers,
28
crabs
35–37,39,40
and Neph-
rops,
34,41
owing to their inability to differentiate between
microplastics and prey. Predators and detritivores may indi-
rectly ingest plastic while consuming prey (i.e. trophic transfer)
or scavenging detrital matter (e.g. marine snows, faecal pellets,
carcasses) containing microplastic.
13,34,35,41,113–115
Micro- and
nanoplastics can adhere to external appendages, including the
gills of the shore crab (Carcinus maenas)
37
and mussels (Mytilus
edulis),
62
and setae of copepod swimming legs and antennules.
10
Other studies have identied that microplastics can bind to
microalgae
116–118
or macroalgae.
119
Microplastic exposure has
been associated with a suite of negative health effects, including
increased immune response,
49
decreased food consump-
tion,
20,22
weight loss,
20
decreased growth rate,
112
decreased
fecundity,
59
energy depletion
22
and negative impacts on subse-
quent generations
59,104
. Microplastics have also been shown to
readily accumulate waterborne persistent organic pollutants
including pesticides, solvents and pharmaceuticals, which may
pose further health effects such as endocrine disruption and
morbidity.
106,120,121
The United Nations Environment Programme (UNEP) has
identied plastic pollution as a critical problem, the scale and
degree of this environmental issue is comparable to that of
climate change.
3
There is currently much public and political
debate surrounding the issue of microplastics as additives to
household and industrial products, and the methods by which
impacts of said microplastics on the environment are to be
measured. Determining the degree to which biota consume
microplastics is essential to determine and monitor ‘good
environmental status’ for plastic pollution (e.g., EU Marine
Strategy Framework Directive, 2008/56/EC; UNEA, US EPA).
Equally, the development of robust environmental legislation is
reliant on toxicological studies with ecological relevance,
requiring an accurate measure of microplastic loads in
natura.
122
As such, it is imperative that researchers are able to
accurately isolate, identify and enumerate microplastic debris
consumed by or entangled with biota. Here we systematically
and critically review methods employed in the extraction,
identication and quantication of microplastic particles
ingested by biota. We consider the effectiveness and limitations
of a range of eld sampling, laboratory exposure, extraction,
and analytical techniques, and consider steps for mitigating
contamination. Our review primarily focuses on peer-reviewed
publications that have investigated the interactions between
invertebrates and sh from the wild, and following controlled
laboratory exposure. A review on extraction of microplastics
from larger marine organisms has been conducted by Pro-
vencher et al. (this issue).
2 Methodological review
For this literature review, we examined original peer-reviewed
research articles, grey literature and conference proceedings
from the 1970s to July 2016. We identied literature referring to
the extraction of microplastics from marine, freshwater and
terrestrial biota using Web of Knowledge, Science Direct,
Scopus and Google Scholar. We also mined the journals Marine
Pollution Bulletin, Environmental Pollution and Environmental
Science and Technology owing to the regularity with which they
publish relevant material. Analysis of microplastic specic
studies was expanded to include historical literature that did
not necessarily have microplastics as the central theme of the
research, such as studies which used uorescent latex beads as
a tracer for feeding and retention experiments. Of the 120
papers included in our meta analysis, 58.3% of studies were
conducted in the laboratory, 38.3% focused on organisms
collected from the wild and 3.4% involved both laboratory
exposure and eld collection (Fig. 2). There were 96 studies
wholly focused on marine organisms, 21 on freshwater, two
studies on both marine and freshwater organisms and one
published study on a terrestrial species.
2.1. Sampling
2.1.1. Field collected organisms. Observations of micro-
plastic uptake by environmentally exposed organisms have now
been reported in a range of habitats, including the sea-surface,
water column, benthos, estuaries, beaches and aquaculture.
4
The diversity of the organisms studied and the habitats from
which they are sampled require a range of collection tech-
niques:
123
the sampling method employed is determined by the
research question, available resources, habitat and target
organism. Benthic invertebrate species such as Nephrops nor-
vegicus may be collected in grabs, traps, and creels, or by bottom
trawling,
34,41
and planktonic and nektonic invertebrates by way
of manta and bongo nets.
10,12,14,16
Fish species are generally
recovered in surface, midwater and benthic trawls, depending
on their habitats.
69–92
Gill nets have been used in riverine
systems.
102
Some species are collected from the eld by hand;
this is common practice for bivalves, crustaceans and anne-
lids.
21,35,37,42,56
Another method is direct collection from shellsh
or sh farms
15,55,56
or from commercial sh markets, where the
capture method is oen unknown.
58,103
Avoiding contamination
and biases during sampling and sample analysis is paramount,
and mitigation protocols are described below.
Fig. 2 Studies of biota interactions with microplastic in the laboratory
and field.
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2.1.1.1. Microplastic losses during eld sampling. Handling
stress, physical movement, and the physiological and behav-
ioural specicities of the sampled organism, may result in the
loss of microplastics prior to animal preservation. Gut evacua-
tion times for animals are varied, ranging from as little as
30 minutes for decapod crustaceans (N. Welden, personal
observations), <2 hours for calanoid copepods,
10
10 to 52 hours
for sh
68,124
to over 150 hours in larger lobsters.
125
Therefore,
some animals might egest microplastic debris prior to anal-
ysis.
41
In such cases, the time between sample collection and the
preservation of the animal must be as short as possible.
Care must also be taken to minimise handling stress or
physical damage. This will reduce the potential for microplastic
regurgitation; the frequency with which animals expel
consumed plastics during sampling is unknown. The copepod
Eurytemora affinis
126
and some sh species have been observed
regurgitating their stomach contents.
127
The main cause of
regurgitation in sh is thought to be related to the expansion of
gas in the swim bladder: this causes the compression of the
stomach and may, in extreme cases, result in total stomach
inversion.
128
Compression of a catch in the cod end might
induce regurgitation in sh.
129
The likelihood of regurgitation
increases with depth of capture, and gadoids are more prone to
regurgitation than atsh. Piscivorous predators are prone to
regurgitation due to their large distensive oesophagus and
stomach.
128,130
As such, regurgitation may bias the stomach
content estimation, affecting consumption estimates and the
presence of plastic debris.
2.1.1.2. Microplastic accumulation during eld sampling.
Laboratory studies have identi ed that nano- and micro-
plastics can adhere to external appendages of marine cope-
pods.
10
Cataloguing such inter action s in natura is complicated
as determining whether the resulting accumulation has
occurred nat urally, or as a by-product of the sampling
regimen, is prohibitive. While most studies focus on the
consumption of plastic, any research considering external
adherence of microplastics should be aware that observed
entanglement may have occurred durin g samp ling and may be
unrepresentative of microplastic–biota interactions a t large. A
similar interaction may occ ur with organisms feeding on
micropla stics during capture in n ets, this is par ticularly
a concern when the mesh size of the net is capable of col-
lecting microplastics, for example, in manta nets (common
mesh size 0.33 mm.
69
Control of microplastic contamination is
discussed in Section 2.4.
2.1.1.3. Sample storage. Consideration should be given to
the storage of biotic samples. Choice of pre servation tech-
nique will largely depend on the research question being
considered; for example, will the xative affect the structure,
microbial surface co mmunities, c hemic al c omposition, colour
or analytical properties of any microplastics within the
sample? 4% formaldehyde and 70% ethanol are commonly
used xatives, however, consultation of resistance tables
suggests these preservatives, albeit at higher concentra tions,
can damage some polymers; for example, polyamide is only
partially resistant to 10% formaldehyde solution, while
polystyrene can be damaged by 100% alcohol (ESI, Table S 1†).
Alternative methods for storage of organisms include desic-
cation
12
and freezing.
41,77,83,89
2.1.2. Laboratory exposed organisms. Laboratory studies
have been implemented to better understand the interactions
between microplastics and biota. Controlled laboratory expo-
sures facilitate monitoring of the uptake, movement and
distribution of synthetic particles in whole organisms and
excised tissues (e.g. gills, intestinal tract, liver). Fluorescently
labelled plastics, either purchased or dyed in the lab
17
allow
visualization of microplastics in organisms with transparent
carapaces,
10,15,30
circulatory uids,
47,49
or histological sections.
105
Where dissection is prohibitive (e.g. mussels) uorescent
microplastics can be quantied by physically homogenising
tissues followed by microscopic analysis of sub-sampled
homogenate.
35
Coherent anti-Stokes Raman scattering (CARS)
has also been used to visualise non-uorescent nano- and
microplastics in intestinal tracts and those adhered to external
appendages of copepods and gill lamellae of crabs.
10,35
Bio-
imaging techniques, however, are not feasible with eld-
sampled biota as environmental plastics do not uoresce, and
may be obscured by tissues or algal uorescence.
2.2. Isolating microplastics
In recent years an increasing number of techniques have been
developed to detect microplastics consumed by biota. Methods
for extracting microplastics from biotic material include
dissection, depuration, homogenisation and digestion of
tissues with chemicals or enzymes. Here we consolidate a range
of optimised methods, and evaluate their benets, biases and
areas of concern:
2.2.1. Dissection. In a large proportion of studies
researchers target specic tissues, primarily the digestive tract
(including the stomach and intestine). In larger animals,
including squid,
64
whales,
97,98
turtles
95
and seabirds,
96
dissec-
tion of the gastrointestinal tract and subsequent quantication
of synthetic particles from the gut is the predominant method
for assessing plastic consumption. In laboratory studies, it is
more common for the whole organism (42% of studies) or the
digestive tract (26% of studies) to be digested or analysed
(Fig. 3A). In comparison, 69% of eld studies targeted the
digestive tract, and 27% looked at the whole organism (Fig. 3B).
Excision of the intestinal tract can also be used to ascertain
consumption of microplastics by invertebrates and vertebrates
including pelagic and demersal sh.
19,34,41,65–67,69–80,83–93
Investi-
gation of stomachs and intestines is relevant for microplastic
>0.5 mm in size. Microplastics larger than this do not readily
pass through the gut wall without pre-existing damage, and
the likelihood of translocation into tissues is too low to
warrant regular investigation.
131,132
Localisation of microplastics
<0.5 mm can be determined by excising organs, such as the liver
or gills,
62,81,105
or, where the research question relates to risks of
human consumption: edible tissues, for example, tail muscles
of shrimp.
38
Microplastics present in dissected tissues can be
isolated using saline washes, density otation, visual inspec-
tion, or digestion (see below).
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2.2.2. Depuration. Should microplastic ingestion be the
primary focus of the study, it is important that any externally
adhered plastics are removed prior to treatment; typically this is
achieved by washing the study organism with water, saline
water or using forceps.
16,61
A depuration step can be used to
eliminate transient microplastics present in the intestinal tract.
Depuration is facilitated by housing animals in microplastic-
absent media (e.g. freshwater, seawater, sediment), with or
without food, and leaving sufficient time for complete gut
evacuation;
54
media should be refreshed regularly to prevent
consumption of egested microplastics.
23
Depuration ensures
only microplastics retained within tissues or entrapped in the
intestinal tract are considered.
23,37,51
Depuration also provides
opportunities for the collection of faecal matter, typically
sampled via siphon, sieve or pipette; faeces can subsequently be
digested, homogenised or directly visualised to assess and
quantify egested microplastics. Faecal analysis has been used to
determine microplastic consumption in a range of taxa,
including sea cucumbers,
28
copepods,
13,17
isopods,
42
amphi-
pods,
33
polychaetes
22
and molluscs.
43–55
2.2.3. Digestion. Enumerating microplastics present in
biota, excised tissues or environmental samples can be chal-
lenging because the plastic may be masked by biological
material, microbial biolms, algae and detritus.
12
To isolate
microplastics, organic matter can be digested, leaving only
recalcitrant materials (Table 1). Traditionally, digestion is con-
ducted using strong oxidizing agents, however, synthetic poly-
mers can be degraded or damaged by these chemical
treatments, particularly at higher temperatures. In Table S1
(ESI†), we have amalgamated chemical resistance data to
highlight the sensitivity of polymers to a range of digestion
agents and storage media. Environmentally exposed plastics,
which may have been subject to weathering, abrasion and
photodegradation, may have reduced structural integrity and
resistance to chemicals compared to that of virgin plastics used
in these stress tests.
133
As such, data ascertained using caustic
digestive agents should be interpreted with caution, and the
likely loss of plastics from the digestive treatment carefully
considered.
2.2.3.1. Nitric acid. Nitric acid (HNO
3
) is a strong oxidizing
mineral acid, capable of molecular cleavage and rapid dissolu-
tion of biogenic material.
134
When tested against hydrochloric
acid (HCl), hydrogen peroxide (H
2
O
2
) and sodium hydroxide
(NaOH), HNO
3
resulted in the highest digestion efficacies, with
>98% weight loss of biological tissue.
51
The optimised protocol
involved digesting excised mussel tissue in 69% HNO
3
at room
temperature overnight, followed by 2 h at 100
C. Desforges
et al.
16
also tested HNO
3
, HCl and H
2
O
2
in digesting
zooplankton, and similarly identied nitric acid as the most
effective digestion agent based on visual observations; here the
optimised digestion protocol consisted of exposing individual
euphausids to 100% HNO
3
at 80
C for 30 minutes. Adaptations
of nitric acid protocols have been successfully used to isolate
bres, lms and fragments from a range of organ-
isms.
23,38,54,61,64,105
While largely efficacious in digesting organic
material, a number of studies observed that oily residue and/or
tissue remnants remained post-digestion,
15,51,63
which have the
potential to obscure microplastics. In response, De Witte et al.
52
proposed using a mixture of 65% HNO
3
and 68% perchloric
acid (HClO
4
) in a 4 : 1 v/v ratio (500 ml acid to 100 g tissue) to
digest mussel tissue overnight at room temperature followed by
10 minutes boiling; this resulted in the removal of the oily
residue. Recovery rates for 10 and 30 mm PS microspheres
spiked into mussel tissue and subsequently digested with nitric
acid range between 93.6–97.9%.
51
However, the high concen-
trations of acid and temperatures applied resulted in the
Fig. 3 Target tissues of animals exposed to microplastics (A) under
laboratory conditions; and (B) in the environment. Total number of
studies ¼ 120.
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destruction of 30 200 mm Nylon bres and melding of 10 mm
polystyrene microbeads following direct exposure. Researchers
have found that polymeric particles, including polyethylene
(PE) and polystyrene (PS), dissolved following overnight expo-
sure and 30 minutes boiling with 22.5 M HNO
3
.
81,135
Polyamide
(PA, Nylon), polyester (PET) and polycarbonate have low resis-
tance to acids, even at low concentrations; furthermore, high
concentrations of nitric, hydrouoric, perchloric and sulphuric
acid are likely to destroy or severely damage the majority of
polymers tested, particularly at higher temperatures (ESI, Table
S1†). The absence of synthetic bres in biota digested using
HNO
3
is likely a reection of the destructive power of the acid.
57
2.2.3.2. Other acids. Formic and hydrochloric acid (HCl)
have also been suggested as digestive agents. With scleractinian
corals (Dipsastrea pallida), formic acid (3%, 72 h) has been used
to decalcify polyps to assist in the visualisation of ingested blue
polypropylene shavings.
32
HCl has also be trialled as a digestant
of microplastics from pelagic and sediment samples; however
this non-oxidizing acid proved inconsistent and inefficient in
digesting organic material.
12
2.2.3.3. Alkalis. Strong bases can be used to remove bio-
logical material by hydrolysing chemical bonds and denaturing
proteins.
136
Excised sh tissues, including the oesophagus,
stomach and intestines, have been successfully digested using
potassium hydroxide (KOH, 10%) following a 2–3 week incu-
bation.
75,89
The protocol has been adapted for the dissolution of
gastrointestinal tracts of sh and mussel, crab and oyster
tissues, either directly or following baking (450
C, 6 h), by
incubating tissues in 10% KOH at 60
C overnight.
58,135
This
latter method has proven largely efficacious in removing
biogenic material, being well suited to the dissolution of
invertebrates and sh llets, but proving less applicable for sh
Table 1 Optimised protocols for digesting biota or biogenic material to isolate microplastics. Assumptions: ‘overnight’ given as 12 h; ‘room
temperature’ given as 20
C
Treatment Exposure Organism Author
HNO
3
(22.5 M) 20
C (12 h) + 100
C (2 h) Blue mussels Claessens et al. (2013)
51
HNO
3
(22.5 M) 20
C (12 h) + 100
C (2 h) Blue mussels oysters Van Cauwenberghe & Jansen (2014)
54
HNO
3
(22.5 M) 20
C (12 h) + 100
C (2 h) Blue mussels lugworms Van Cauwenberghe et al. (2013)
23
HNO
3
(100%) 20
C (30 min) Euphausids copepods Desforges et al. (2015)
16
HNO
3
(69–71%) 90
C (4 h) Manilla clams Davidson & Dudas (2016)
61
HNO
3
(70%) 2 h Zebrash Lu et al. (2016)
105
HNO
3
(22.5 M) 20
C (12 h) + 100
C (15 min) Brown mussels Santana et al. (2016)
63
HNO
3
(65%) 20
C (12 h) + 100
C (10 min) Blue mussels De Witte et al. (2014)
52
HClO
4
(68%) (4 : 1)
HNO
3
(65%) 20
C (12 h) + 100
C (10 min) Brown shrimp Devriese et al. (2015)
38
HClO
4
(68%) (4 : 1)
CH
2
O
2
(3%) 72 h Corals Hall et al. (2015)
32
KOH (10%) 2–3 weeks Fish Foekema et al. (2013)
75
KOH (10%) 60
C (12 h) Fish Rochman et al. (2015)
58
KOH (10%) 2–3 weeks Fish Lusher et al. (2016)
89
H
2
O
2
(30%) 60
C Blue mussels Mathalon & Hill (2014)
53
H
2
O
2
(30%) 20
C (7 d) Biogenic matter Nuelle et al. (2015)
137
H
2
O
2
(15%) 55
C (3 d) Fish Avio et al. (2015)
81
H
2
O
2
(30%) 65
C (24 h) + 20
C (<48 h) Bivalves Li et al. (2015)
57
NaClO (3%) 20
C (12 h) Fish Collard et al. (2015)
82
NaClO
3
(10 : 1) 20
C (5 min)
Proteinase K 50
C (2 h) Zooplankton copepods Cole et al. (2014)
12
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