Review of micro- and nanoplastic contamination in the food chain

Figures & data

Table 1. Number of published studies on microplastic contamination in different animal species, identified by authors since 2010.

Animal species	Number of published studies since 2010
Bear	1
Birds	12
Bivalves (e.g. mussels, clams, scallops, etc.)	22
Chicken	1
Crustaceans (e.g. lobsters, shrimps, langoustines, crabs, etc.)	20
Fish and sea mammals (e.g. seals, dolphins, whales)	208
Insects	2
Seabirds	152
Turtles	5

Table 2. Levels of microplastic contamination in mussels. When reviews cite other papers, the original paper is cited between brackets. ‘Item’ means a microplastic piece. SD indicates Standard Deviation.

Common name/Taxonomy	Level of microplastic contamination (± SD)	Samples	Location and sampling rationale	Method	Control/blank	Cross-contamination prevention	References
Brown mussel/ Perna perna	0.35 items/g (commercial brands) 0.26 items/g (wild mussels from groyne) 0.51 items/g (wild mussels from quayside)	5 mussels per location (6 for wild, plus 3 brands at market for commercial) from Belgian department stores and Belgian groynes and quaysides	-	Acid digestion with HNO3:HClO4 (4:1 v:v). + filtration + stereomicroscope observation	3 blanks (=without mussel tissue) analyses per location	Samples stored in aluminium foil in the freezer prior analysis. Solutions filtered (10–20 µl). Laboratory glassware cleaned with acetone and filtered type 1 water. Mussel body rinsed with filtered type 1 water to remove the intervalve water. Digestion solution covered with a clock glass to avoid contamination by air. No synthetic clothing allowed in the lab.	De Witte et al. 2014
Mediterranean mussel/ Mytilus Galloprovincialis	~2.5 ± 1.5 items/g (all individuals contaminated)	18	Most commercially popular. From fishery market in China	Hydrogen peroxide treatment + Floatation and filtration with saline (NaCl) solution + microscope observation and µ-FT-IR verification	One blank extraction group without tissue	Liquid (freshwater, salt water and hydrogen peroxide) filtered with 1mm filter paper prior to use. Containers and beakers rinsed three times with filtered water.Samples covered if not in use. Experimental procedures finished as soon as possible	Wesch et al. 2016 (Li et al. 2015)
	0.35 items/g (consumption mussels) 0.26 items/g (wild mussels from groyne) 0.51 items/g wild mussels from quayside)	5 mussels per location (6 for wild, plus 3 brands at market for commercial) from Belgian department stores and Belgian groynes and quaysides	-	Acid digestion + filtration + stereomicroscope observation	3 blank (= without mussel tissue) analyses per location	Samples stored in aluminium foil in the freezer prior analysis. Solutions filtered (10–20µl). Laboratory glassware cleaned with acetone and filtered type 1 water. Mussel body rinsed with filtered type 1 water to remove the intervalve water. Digestion solution covered with a clock glass to avoid contamination by air. No synthetic clothing allowed in the lab.	De Witte et al. 2014
	3.6–12.4 items/individual (4.4–11.4 items/g)	10	3 commercial stocks of mussels from local markets (provenience: central Adriatic Coast; Ligurian Sea Coast, North East Sardinia, Tyrrhenian Sea), + 1 stock from natural rocky bottom in Tuscany (central Tyrrhenian Sea)	Digestion with H2O2 30% + filtration (0.45 μm) + stereomicroscopy	Filters (n  =  3) on glass Petri dish exposed to laboratory air. Blank tests	Glass materials and cotton dresses	Renzi et al. 2018
Blue Mussels/ Mytilus edulis	0.36 ± 0.07 items/g	36	Samples from a mussel farm in Germany and originated from the North Sea	Acid digestion + filtration (0.8 and 5 μm membrane filter) + microscope observation + micro-Raman spectrometer analysis	Procedural blanks (i.e. samples containing no tissue) included in every acid destruction	100% cotton lab coat, equipment rinsed 3 times with filtered deionised water (0.8 μm membrane filter) before use and all sample processing performed in a clean laminar flow cabinet.	Luscher., 2015-Li et al. 2016-Wesch et al. 2016 (Van Cauwenberghe and Janssen 2014)
	0.35 items/g (consumption mussels) 0.26 items/g (wild mussels from groyne) 0.51 items/g (wild mussels from quayside)	5 mussels per location (6 for wild, plus 3 brands at market for commercial) from Belgian department stores and Belgian groynes and quaysides	-	Acid digestion with nitric acid + filtration (5 μm) + stereomicroscope observation	3 blank (= without mussel tissue) analyses per location	Samples stored in aluminium foil in the freezer prior analysis. Solutions filtered (10–20 µl). Laboratory glassware cleaned with acetone and filtered type 1 water. Mussel body rinsed with filtered type 1 water to remove the intervalve water. Digestion solution covered with a clock glass to avoid contamination by air. No synthetic clothing allowed in the lab.	Luscher. 2015; Li et al. 2016; Wesch et al. 2016 (De Witte et al. 2014)
	First analysis: 170 (wild) −375 (culture) fibres/5 mussels Second analysis: average number per observed to be ∼126 items/wild mussel from McCormack’s Beach, ∼106 items/wild mussel from Rainbow Haven Beach, and ∼178 items/farmed mussel	10 wild mussels + 10 cultured mussels for first analysis, 15 wild mussels + 15 cultured mussels for second analysis	Comparison of aquaculture and wild habitat. Mussels from an aquaculture site (off the west coast of Newfoundland and Labrador) and two protected beaches along Nova Scotia’s Eastern Shore, on the Atlantic coast of Canada	Dissection + digestion with H2O2 + separation via flotation with concentrated saline solution of NaCl in H2O + filtration (0.8 μm) + microscopic observation	Control samples to test extraction efficiency	New and unopened bags and Petri dishes. All containers and beakers rinsed with distilled water prior use. Samples kept covered when possible.	Wesch et al. 2016 (Mathalon and Hill 2014)
	0–10 items/individual (average 0.2 ± 0.3 items/g) (all contaminated)	Sample size not described	Comparison of locations along French–Belgian–Dutch coastline	clearance + acidi digestion with 69% nitric acid + filtration (5 μm) + microscopic observation + Raman spectroscopy	Blank extractions	Filtered solutions	Wesch et al. 2016 (Van Cauwenberghe et al. 2015)
	5–19 items/individual	23	North Sea	Digestion with HNO3 and H2O2 + extraction based on density separation + filtration (0.7 μm) microscope observation	One blank per 3 sample analyses to correct for background contamination	All equipment rinsed with Milli-Q water before use. Samples, subsamples and filters covered. Glassware was used where possible	Karlsson et al. 2017
	3.4 ± 0.48 items/individual	62	From Scottish commercial suppliers and deployed in cages in the estuary of the Forth River, Edinburgh, UK, in Port Edgar	enzymatic digestion with Corolase® 7089 + filtration (0.8 μm) + dissection microscope + FT-IR	Airborne fibre contamination: 1 Milli-Q water control sample (100 ml) . Procedural blanks (taken into account in calculations)	Samples covered, vials capped with aluminium, cotton lab coats, equipment rinsed with Milli-Q water, and glassware acid-washed	Catarino et al. 2018
	coastal mussel tissue: 1.1 to 6.4 items/individual (0.7 to 2.9 items/g); supermarket products: live mussels: 0.9 items/g, processed mussels: 1.4 items/g; seawater: 3.5 ± 2.0 items/L; size range: 8 µm to 4.7 mm (most particles between 5 µm and 250 µm); 50% of randomly selected particles confirmed as microplastic, 37% rayon or cotton, ~10% cellulose	all mussel and seawater samples contaminated	mussels from 8 sites along UK coasts and 8 UK supermarkets (farmed, live and processed mussels), 6 replicates from each; seawater sampled from the same sites of the UK coast except 2 sites	tissue extraction using 30% H2O2, incubation at 65°C in oscillator at 80 rpm for 24h, then room temperature for 24–48 h; density separation of particles and debris in concentrated saline solution, overlying water filtered on 5 µm cellulose nitrate membrane filter, filter observed under stero microscope; particle identity confirmed by µFT-IR	airborne contamination: blank extraction without tissue, 6 replicates: 0.67 ± 0.75 (mean ± SD) item per filter compared to 5.70 and 8.63 items/filter for supermarket and coastal mussel tissues, respectively	water and H2O2 filtered with 1 µm filter paper, apparatus rinsed 3 times with filtered water; no description of laminar flow nor special clothing precautions	Li et al. 2018
Blue mussel, Mediterranean mussel, bay mussel/ Mytilus spp (M. edulis, M. galloprovincialis, M. trossulus)	3.2 ± 0.52 tems/individual	36	From various locations along Scottish coast	enzymatic digestion with Corolase® 7089 + filtration (0.8 μm) + dissection microscope + FT-IR	Airborne fibre contamination: 1 Milli-Q water control sample (100 ml) . Procedural blanks (taken into account in calculations)	Samples covered, vials capped with aluminium, cotton lab coats, equipment rinsed with Milli-Q water, and glassware acid-washed	Catarino et al. 2018
Northern horsemussel/ Modiolus modiolus	3.5 ± 1.29 tems/individual	6	From various locations along Scottish coast	enzymatic digestion with Corolase® 7089 + filtration (0.8 μm) + dissection microscope + FT-IR	Airborne fibre contamination: 1 Milli-Q water control sample (100 ml) . Procedural blanks (taken into account in calculations)	Samples covered, vials capped with aluminium, cotton lab coats, equipment rinsed with Milli-Q water, and glassware acid-washed	Catarino et al. 2018

Table 3. Levels of contamination reported in honey, beer, salt and water. ‘item’ means microplastic piece.

Product	Level of microplastic contamination (sometimes average ± SD)	Samples	Location and sampling rationale	Method	Blanks	Contamination prevention	References
Honey	40 to 660 coloured fibres/kg of honey, with a mean value of 166 ± 147/kg of honey; 0–38 fragments/kg of honey; mean 9 ± 9/kg of honey; fibres and fragments supposed to be synthetic	19	From Germany, France, Italy, Spain and Mexico, from supermarkets (8) or producers (11)	Filtration (40 μm) + 30% H2O2 + filtration (0.8-µm) + dissection microscope + staining with fuchsin and rose Bengal	-	Limited exposure of samples to the open air	Liebezeit and Liebezeit 2013
	10 to 336 fibres/kg and 2 to 82 fragments/kg supposed to be synthetic	47	From German local supermarkets or bee-keepers	Filtration (40 μm) + 30% H2O2 + filtration (0.8-µm) + dissection microscope + staining with rose Bengal	Blank filters through complete scheme (maximum of 2 fibres/filter)	Filtration unit and all glassware covered during procedure. Laboratory air sucked through a previously rinsed filter for 1 h at 50 L min–1 (2 fibres, 0 fragments and 3 granular particles)	Liebezeit and Liebezeit 2015
	32–108 coloured fibres/kg (mostly cellulose but a minor part being PET fibres); 8–28 blue particles/kg (unknown origin)	5	From 5 locations in Switzerland	Filtration (30 μm) + sonication in ethanol + 30% H2O2 + filtration (1-µm) + microscope + FTIR–ATR and Raman spectroscopy	Particle background: filtration and preparation procedures performed on ultrapure water.	Material cleaned in a dishwasher and rinsed with ultrapure water. Ultrapure water from laboratory water purification system used for dilution of honey. Honey handling, dilution, and filtration performed in a laminar flow box (SKANAIR VSE 2000). All inner surfaces of the laminar flow box cleaned with particle-free detergent and cleanroom wipes. Cleanroom-grade gloves and sleeves.	Mühlschlegel et al. 2017
Salt	16–84 item/kg (sea salt), 8–102 item/kg (lake salt) and 9–16 item/kg (rock salt)	16	From Turkish market	Digestion with 30% H2O2 + filtration (0.2 μm) + 4M NaI + Microscopic and Raman examination	Empty petri dish uncovered near working area during process.	All equipment and work surfaces cleaned with pure ethanol 70%, and all operations performed under a closed fume hood. Cleaning with cellulose towel (any Raman spectrum presenting cellulose removed from results). Bottles kept capped and all other equipment kept in lidded glass containers.Microscope covered with Nylon bag	Gündoğdu, S., 2018
	46.7 ± 0.58–806 ± 15.3 particles/kg (average: 212 particles/kg, all positive)	12	From US grocery stores (from North, Celtic, Sicilian, Mediterranean, Utah, Hawaiian, Atlantic, Pacific, Baja Seas and Hymalaya)	Filtration (11 μm) + biological stain Rose Bengal + dissection microscope	Bottle blanks run by filling 2 HDPE bottles with deionized water. Lab blankswith only deionized water (1 blank was processed for each brand (n = 12 for each).	All glassware covered with a watch glass and washed between trials. Laminar airflow cabinet. Workspace was wiped down every week. Filters inspected under a microscope. Filtration times recorded so that the window of time for potential contamination was known. Cotton lab coat and sterling nitrile powder free exam gloves.	Kosuth et al. 2018
	1 to 10 items/kg (16 positive)	17	From a Malaysian market originating from 8 countries (Australia, France, Iran, Japan, Malaysia, New Zealand, Portugal, South Africa)	Filtration (150 μm) + NaI (4.4M) + filtration + stereomicroscope + Raman spectroscopy	1 procedural blank with only filtered deionized water tested simultaneously during extraction procedure and 1 procedural blank with only NaI solution tested simultaneously during density separation process	Cotton lab coat and nitrile gloves. All liquids filtered through 2.7 μm glass fibre filter membranes. Glassware washed with dishwashing liquid, with deionized water, with ethanol. The work surface cleaned with 70% ethanol. Horizontal laminar flow cabinet	Karami et al. 2017
	50± 7–280± 3 items/kg salt	21	From sea: Atlantic Ocean (Huelva, Cádiz, Lanzarote, La Palma, Galicia), Mediterranean sea (Barcelona, Gerona, Valencia, Alicante, Murcia, Menorca) From well: Alicante, Cuenca, Añana	Filtration (5 μm) + microscope + FT-IR	-	-	Iñiguez et al. 2017
	7–680 items/kg (550−681 particles/kg in sea salts, 43–364 particles/kg in lake salts, and 7−204 particles/kg in rock/well salts)	15	From supermarkets throughout China	Digestion with 30% H2O2 + Floatation and Filtration (5μm) + Stereomicroscope + μ-FT-IR	One blank extraction group without salt tested simultaneously for procedural contamination	All liquids filtered (1μm). All containers and beakers rinsed with filtered water. Samples immediately covered.	Yang et al. 2015
Sugar	217 ± 123 transparent and coloured fibres/kg of sugar; 32 ± 7 fragments/kg of sugar	5 commercial sugars		Filtration (0.8-µm) + 30% H2O2 + dissection microscope + staining with fuchsin and rose Bengal	-	Samples not exposed longer than necessary	Liebezeit and Liebezeit 2013
Canned sardines and sprats	1 −3 items/contaminated brand (4 contaminated brands – cans containing 2–30 fish each)	20 brands (cans containing 2–30 fish each)	From Australian and Malaysian markets (originating from Canada, Germany, Iran, Japan, Latvia, Malaysia, Morocco, Poland, Portugal, Russia, Scotland, Thailand,Vietnam)	Digestion with KOH + filtration (149 μm) + NaI + filtration (8 μm) + Stereomicroscope + Micro-Raman spectroscopy + field emission scanning electron microscope	Procedural blanks run in parallel with the samples.	Extraction process inside a pre-cleaned (with ethanol using SCOTT® paper towels made of cellulose) horizontal laminar flow cabinet. Cotton lab coat and nitrile gloves. All glassware and forceps washed with a detergent, rinsed with deionized water, with ethanol, and dried in an oven at 50 °C for 5 h. Caps of laboratory bottles loose, and the forceps placed in beakers covered with an aluminum foil. Solutions filtered through 8 μm filter membranes (one filter membrane per ≈ 2 L of solution/solvent). The gloves were removed from the packaging and worn inside the laminar flow cabinet.	Karami et al. 2018
Beer	2–79 fibres/L (RSDs = 130%), 12–109 fragments/L (RSDs = 205%), 2–66 granules/L (RSDs = 103%)	24	From German local supermarkets	Filtration (0.8 μm, 40 μm for wheat beers) + staining of natural organic particles with Rose Bengal + dissecting microscope	Blank filters through complete scheme (maximum of 2 fibres/filter)	Filtration unit and all glassware covered during procedure. Filters rinsed with filtered water and inspected microscopically. Laboratory air sucked through a previously rinsed filter for 1 h at 50 L min–1 (2 fibres, 0 fragments and 3 granular particles)	Liebezeit and Liebezeit 2014
	16 ± 15 fibres/L (Blank sample: 15 ± 9 fibres/L); 21 ± 16 fragments/L (Blank sample: 20 ± 13 fragments/L); 27 ± 10 granules/L (Blank sample: 15 ± 12 granules/L)			Same method as in Liebezeit and Liebezeit 2014 to show the inconsistency of results when compared to blanks, due to lack of validated methodology for detection of microplastics in food and beverages	Same method as in Liebezeit and Liebezeit 2014	Same method as in Liebezeit and Liebezeit 2014	Lachenmeier et al. 2015
	0–14.3 particles/L (average: 4.05 particles/L, all positive)	12	From breweries using water from the five Laurentian Great Lakes (US)	Filtration (11 μm) + biological stain Rose Bengal + dissection microscope	Bottle blanks run by filling 2 HDPE bottles with deionized water. Lab blankswith only deionized water (1 blank per brand (n = 12 for each).	Rinsed sample collection bottles. All glassware covered with a watch glass and washed between trials. Laminar airflow cabinet. Workspace was wiped down every week. Filters inspected under a microscope. Filtration times recorded so that the window of time for potential contamination was known. Cotton lab coat and sterling nitrile powder free exam gloves.	Kosuth et al. 2018
Bottled water	10.4 particles (>100 µm)/L; 325 particles (6.5–100 µm)/L; 242 bottles contaminated, all single-use plastic bottles except 1 glass bottle	259	From 19 locations in 9 countries (China, USA, Brazil, India, Indonesia, Mexico, Lebanon, Thailand, Germany) – 11 brands	Nile Red solution addition + filtration (1.5 μm) + optical microscope + FTIR	13 blanks with deionized water or acetone	Laminar flow fume hood. Workspace wiped down every week. All glassware covered with a watch glass and washed. Filters inspected under a microscope. Cotton lab coat and sterling nitrile powder free exam	Mason et al. 2017
	118 ± 88 particles/L (returnable bottles), 14 ± 14 particles/L (single-use plastic bottles), 11 ± 8 particles/L (beverage cartons), 50 ± 52 particles/L (glass bottles).	22 bottles, 3 cartons, and 9 glass bottles	From grocery stores in Germany	Filtration + μ-Raman spectroscope	Blank with ultra-pure water: 14 ± 13 particles/L	Laminar flow workbench. 100% cotton laboratory coat, particle free nitrile gloves and arm sleeves. Gloved hands washed with detergent and rinsed with Milli-Q water. Glass funnel and vessels cleaned with deionized water, detergent and cleanroom wipes, sonicated in Milli-Q water, rinsed with Milli-Q water. Imaging microscope on the clean bench for before and after filtration pictures. ‘Flush filter’ wiped off with a cleanroom wipe.Outside of water bottles or beverage cartons cleaned Labels removed and package rinsed with Milli-Q water	Schymanski et al. 2018
Tap water	0–61 particles/L (average: 5.45 particles/L)	159	From Cuba (1), Ecuador (24), England (3), France (1), Germany (2), India (17), Indonesia (21), Ireland (1), Italy (1), Lebanon (16), Slovakia (8), Switzerland (2), Uganda (26),US (36)	Filtration (2.5 μm) + biological stain Rose Bengal + dissection microscope	Bottle blanks run by filling 2 HDPE bottles with deionized water. Lab blanks with only deionized water (once each day that samples were processed (n = 30))	Sample collection bottles rinsed twice with running tap water before filling. All glassware covered with a watch glass and washed between trials. Laminar airflow cabinet. Workspace was wiped down every week. Filters inspected under a microscope. Filtration times recorded so that the window of time for potential contamination was known. Cotton lab coat and sterling nitrile powder free exam gloves.	Kosuth et al. 2018
	Particles >100 µm: 15.6 particles/50 L (Blank sample: 13.2 particles/50 L) Limit of detection = mean blank + (1.645 x SD for blank) = 29 particles/50 L	17 + 3	Danish tap water	17 samples treated with acetic acid 32 % for 1 h to dissolve inorganic particles, sample column flushed with ethanol onto a stainless steel filter (10 µm) + stereomicroscope (44 % of the particles further analysed by µFT-IR) 3 samples filtered on Anodisc 0.2 µm filter + µFT-IR	5 blank samples	Stainless steel or glass equipment for sampling, rubber O-rings and pipes cleaned with a 5% detergent and water, air filtration	Strand et al. 2018

Table 4. Advantages and limitations of the methods for sample preparation and characterisation of microplastics.

Sample preparation methods	Advantages	Limitations
Filtration	Simple Low cost Categorises particles by size Suitable for water & not viscous beverages	Clogging on small pore size filter Possible loss of particles on large pore size filter Possible sample contamination if using filter fibers (glass fibre filter, cellulose filter) Lower contamination requires use of pore etched membrane filters Possible difference between the nominal and the effective cut-off of the filter
Flotation/Sedimentation	Simple Low cost Separates microplastics from inorganic matter	Critical choice of the type of gradient Plastic particles must be detached from matrix materials Recycling or disposal of contaminated salts required
Matrix digestion
Acidic chemical digestion	Suitable for biofilms & animal tissues, organs	Need optimisation to digest all organic matter Can modify, damage or destroy plastic particles
Alkaline chemical digestion	Suitable for biofilms & animal tissues, organs	Need optimisation to digest all organic matter Can modify, damage or destroy plastic particles
Enzymatic digestion	Less damage to polymers Suitable for biological matrices	Need careful optimisation Might require several laboratory steps More expensive Variable enzymatic activity depending on matrix, storage, origin
Oil extraction	Suitable for aqueous media Separates microplastics from inorganic matter Compatible with FT-IR	Possible damage of low melting point polymers Particles must be sufficiently hydrophobic to be extracted by oil
Methods of detection/identification/quantification
Optical vibrational spectroscopy
Micro-FTIR spectroscopy	Identifies polymer Selective & reproducible detection Requires small sample amounts Requires limited sample preparation Non-destructive FTIR-microscopy can identify individual particles with sizes down to around 2 microns and quantify the number of particles of each type Able to localise debris in living organisms	May not be reliable on weathered samples, degraded samples or mixed polymer samples. FTIR-microscope combination are expensive
Micro-Raman spectroscopy	Identifies polymer Selective & reproducible detection Requires small sample amounts Requires limited sample preparation Partially non-destructive Raman-microscopy can identify individual sub-micron particles and quantify the number of particles of each type Able to localise debris in living organisms	Auto-fluorescence of samples may mask the Raman signal May not be reliable on weathered samples, degraded samples or mixed polymer samples Samples may be damaged by extended laser exposure Time consuming for wide area imaging Raman-microscope instruments are expensive
Thermal analysis
Differential scanning calorimetry (DSC)	Fast, simple Identifies polymers using reference materials	Limited application to mixtures of polymers
Pyrolysis gas chromatography-mass spectrometry (Py-GC-MS)	Identifies polymers and organic additives Quantifies small quantities in mass concentration (<0.5 mg) Requires limited sample pre-treatment Suitable for biological matrices and screening of environmental samples	Time consuming No data on particle number, size & shape Need an expert operator Manual manipulation of sample required which limits lower particles size to fractions of mm
Fluorescent tagging with Nile Red	Detects & quantifies particles 20 µm – 1 mm Low cost Semi-automated	Not reliable with weathered samples Residual organic matter in background matrix may become stained producing false positives Not able to identify polymer types, needs additional instrumentation e.g IR spectroscopy Dye fluorescence may prevent identification by Raman spectroscopy
Time-of-Flight Secondary Ion Mass Spectrometry (ToF-SIMS)	High spatial resolution imaging possible Suitable for mixtures of particles Able to detect particles in the low µm range Can potentially provide information on inorganic and organic chemical contaminants (heavy metals) Identifies polymers	Complex to use High cost Relative quantification of different polymers is usually not possible due to matrix effects Identification complicated by weathering or by surface contaminants Sample must be vacuum compatible

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