Literature study
Determination of microplastics by
mass spectrometry
Niels Janssen, 11934042
Amsterdam
October 2019
MSc Chemistry
Analytical Sciences
Literature Study
Determination of
microplastics by mass
spectrometry
by
Niels
Janssen
11934042
October 2019
Examiner:
Second examiner:
ABSTRACT
Microplastics are emerging global contaminants found in the aquatic, terrestrial and atmospheric environment and carry potential risks to the environment, animals and humans. These risks are a result of leaching and accumulation of potential toxic chemicals from and to microplastics, and ingestion of microplastics by a wide range of organisms. Microplastics are most commonly determined by spectroscopic techniques such as Fourier-transform infrared spectroscopy, Raman spectroscopy, and scanning electron microscopy, where samples are vigorously pretreated, and quantification is generally performed by manual counting. However, these methods are time-consuming and propose a high risk of error. To overcome these shortcomings, mass spectrometric techniques such as pyrolysis- and thermal extraction and desorption-gas chromatography-mass spectrometry and liquid chromatography-tandem mass spectrometry are utilized. In this study, all mass spectrometric techniques for the determination of microplastics in varying environmental matrices are extensively reviewed and additionally general concerns regarding microplastic determination are elaborated upon. It was found that pyrolysis-gas chromatography-mass spectrometry and liquid chromatography-tandem mass spectrometry required most sample preparation but showed to be most sensitive. Lower sensitivity was found using thermal extraction and desorption-gas chromatography-mass spectrometry, but this technique showed to be capable of measuring relatively untreated samples. Lastly, high risks of contamination, a lack of reference materials, and the near absence of interlaboratory comparison studies affect the trustworthiness of the acquired data in microplastic research. As for future research, studies on human samples could aid in the elucidation of the risk of microplastics to humans.
ABBREVIATIONS
(5%-Phenyl)-methylpolysiloxane (HP-5ms) accelerated solvent extraction (ASE) advanced polymer chromatograph (APC) atmospheric pressure chemical ionization (APCI) atmospheric pressure photoionisation source (APPI) attenuated total reflectance (ATR) attenuated total reflectance-Fourier transform-infrared spectroscopy (ATR-FTIR)
chlorinated PE (cPE)
chlorosulfonated polyethylene (csPE) cooled injection system (CIS) Correct Nominal Kendrick Mass (CNKM) desorption electrospray ionisation (DESI) differential scanning calorimetry (DSC) direct analysis real-time (DART) ethylene-vinyl acetate (EVA)
expanded polystyrene (EPS)
extracted ion chromatogram (EIC)
focal plane array (FPA)
Fourier transform-infrared spectroscopy (FT-IR) high-density polyethylene (HDPE)
internal standard (IS)
Kendrick Mass Defect (KMD)
limit of detection (LOD)
limit of quantification (LOQ) liquid chromatography-high-resolution mass spectrometry (LC-HRMS) low-density polyethylene (LDPE) low-density polypropylene (LDPP)
mass spectrometry (MS)
matrix-assisted laser desorption ionisation (MALDI) medium-density polyethylene (MDPE) methyl dimethyl diisocyanate polyurethane (MDI-PUR)
micro furnace (MF)
organic plastic additives (OPAs) phenyl arylene polymer (DB-5ms) poly (methyl methacrylate) (PMMA) poly(acrylonitrile-co-styrene-co-butadiene) (ABS)
poly(vinyl acetate) (PVA)
polyamide (PA)
polybutadiene (PB) polycarbonate (PC) polydimethylsiloxane (PDMS) polyester (PET) polyformaldehyde (POM) polypropylene (PP) polystyrene (PS) polytetrafluoroethylene (PTFE) polyurethane (PUR)
polyvinyl acetate (PVAc)
polyvinylchloride (PVC)
pyrolysis-gas chromatography-mass spectrometry (Pyr-GC-MS) relative standard deviation (RSD) relative standard error (RSE) scanning electron microscopy (SEM) scanning electron microscopy energy-dispersive X-ray spectroscopy (SEM-EDS) secondary ion mass spectrometry (SIMS) selected ion monitoring (SIM) size-exclusion chromatography (SEC) solid analysis probe (ASAP) solid-phase adsorber (SPA) styrene–butadiene block copolymer (SBS) styrene–butadiene rubber (SBR) suspended particulate matter (SPM) tetramethylammonium hydroxide (TMAH) thermal desorption-gas chromatography-mass spectrometry (TDS-GC-MS) thermal extraction desorption gas chromatography (TED-GC) thermogravimetry analysis (TGA)
time of flight (TOF)
CONTENTS
1. Introduction 5
2. Microplastics 6
2.1 Primary and secondary microplastics 6 2.2 Sources of microplastics 7 2.3 Distribution of microplastics 8 2.4 Risks of microplastics 10
3. Overview of the general workflow of the determination of microplastics 12
3.1 Sampling 12
3.2 Sample preparation 13
3.3 Particle analysis 14
3.4 QA/QC and blank issues 16
4. Overview of methods for the determination of microplastics using MS 17
4.1 Thermogravimetry analysis-mass spectrometry 17 4.2 Pyrolysis-gas chromatography-mass spectrometry 18 4.3 Thermal extraction and desorption - gas chromatography mass spectrometry 28 4.4 Time-of-flight-secondary ion mass spectrometry 34 4.5 Liquid chromatography-mass spectrometry 35
5. Discussion and Conclusion 38
1.
Introduction
Microplastics are major global contaminants: plastic microlitter is found mainly throughout marine systems and freshwater1–5 but is also found in the terrestrial environment6,7 and even air8–11. Due to their
small and variable size (1 – 5000 µm) predicting particle transport following release into the environment is difficult. Also, the environmental impact of microplastics is not yet fully understood but additional to the potential leaching of harmful additives from microplastics12, persistent organic pollutants and certain
metals are able to adsorb onto the microlitter.13,14 Also, ingestion of microplastics has been identified
within species spanning from invertebrates to large marine animals.15,16 In order to understand the
sources, fate, and pathways of microplastics in the environment, suitable techniques should be applied to measure individual microplastics and microplastics in environmental matrices. Common and techniques involve microscopic detection of microplastics and measurements by FTIR/Raman spectroscopy; however poor structural information is obtained.17 Therefore, in this article mass
spectroscopic methods for the determination of microplastics will be reviewed. Also, information about physical properties, sources, distribution, and risks of microplastics will be provided along with the general workflow of microplastic determination in environmental samples.
2.
Microplastics
2.1 Primary and secondary microplastics
Microplastics are tiny plastic fragments, granules and fibers that are present in several size-ranges. In general all plastic fragments, granules and fibers smaller than 5 mm are considered microplastics.18,8
However, microplastic particles of a few micrometers in size are also reported, which makes it challenging to compare data from different studies.18,19 Just as the size, the shape of microplastics can
vary a lot as well: common shapes (in the marine environment) include: pellets, fibers, fragments, spheres, sheets and foams.20 Furthermore, microplastics are categorized into two main categories:
primary- and secondary microplastics.18 Primary microplastics are produced on purpose to be used
typically in facial cleansers and cosmetics, air-blasting media and medicine. Also, virgin plastic production pellets (2 – 5 mm) can be considered microplastics although these pellets are usually used for the production of macroplastics (larger plastics). Furthermore, primary microplastics in exfoliating scrubbers and cleansers are replacing natural (degradable) microscopic particles such as oatmeal and ground almonds and since the patenting of microplastic scrubbers in cosmetics the amount of cosmetics containing plastics has increased substantially.21 These ‘’micro-beads’’ can vary in shape and composition
depending on the product they are used for; however, different sizes, polymers, and shapes have been found in a single cosmetic product.22 Another source of primary microplastics is air blasting technology;
this technology includes the cleaning of e.g. machinery and engines of rust and paint by blasting microplastic scrubbers onto it. Also, these microscopic scrubbers become often contaminated with heavy metals which increases the potential risk of these microplastics.18
On the other hand, microplastics that are not produced on purpose but are a result of breakdown of lar-ger plastic (macroplastic) debris are called secondary microplastics.18 These tiny plastic fragments are a
result of chemical, biological and physical processes on both sea and land that weaken the structural integrity of the plastic debris which ultimately leads to fragmentation. Furthermore, on land macroplastics are exposed to a relatively high amount of sunlight for prolonged periods which causes the macroplastic to undergo photo-degradation. This event is caused by cleavage of bonds in the polymer matrix due to oxidization by ultraviolet radiation present in sunlight. Aside the risks of (secondary) microplastics themselves, degradation of macroplastics may result in additives being released from the polymer matrix. Theses additives were added to virgin plastics in order to enhance certain desired properties of the plastics; common additives are fillers, colorants, lubricants, plasticizers, antioxidants and flame retardants.23 Furthermore, different kinds and amounts of additives are being used in different
kinds of applications of plastic, so a complex manufactured plastic contains a complex mixture of additives. More on the general risks of microplastics and the risks of the additives will be provided in paragraph (2.4).
In addition, this degradation of plastics is an ongoing process which occurs most effectively on land (including beaches) since beside the lack of direct sunlight, the cold and haline conditions of the marine environment most likely prevent this photo-oxidation to happen.18 This ongoing degradation results in
have different properties and bioavailability than microplastics and may therefore cause environmental problems at another level.24 Lastly, the replacement of traditional plastics by biodegradable plastics may
also contribute to the increase of microplastics in the (marine) environment. 18 These biodegradable
plastics are usually composed of starch, vegetable oils or chemicals that ought to decrease degradation times, combined with a certain degree of synthetic polymers. Subsequently, after decomposition, in industrial composting plants under hot, humid and well-aerated conditions, the synthetic polymer will be left behind.24 However, if these bio-plastics end up in sea (80% of marine debris are plastics)25 no
terrestrial microbes are present and even the degradable components will take longer to degrade. Finally, when decomposition occurs, microplastics are released into the marine environment.26
2.2 Sources of microplastics
Plastics are being used extensively in industry and by society which results in a variety of sources of general plastic waste and, therefore, primary and secondary microplastics.27 For instance, in Europe
plastic packaging (39.5%), building and construction (20.1%), and automotive industry (8.6%) are the main sectors demanding the most plastic material. In addition, electrical and electronic industry (5.7%), and agriculture (3.4%) also contribute to the European plastic usage. Lastly, other uses such as consumer and household appliances, sport, health and safety contribute for 22.7% to the European plastic consumption. However, these sectors may not be contributing similarly to the production of microplastics or leakage of microplastics into the environment.
For instance, sewage treatment plants play a big role in removing microplastics from wastewater but also in spreading microplastics back into the environment.28,29 For example, sewage treatment plants filter
and clean primarily household sewage wastewater using several chemical, biological and physical processes. In developed countries sewage treatment plants have (at least) two treatment stages: a primary stage where oils, sand and other large solids are removed by physical processes and the secondary stage where organic matter is broken down by bacteria. In both treatment stages microplastics have been observed and it was estimated that one microplastic particle per liter of treated water was released back into the environment.28 One particle per liter is not disproportionately large;
however, microplastics are mainly removed in the primary stage resulting in relatively high concentrations microplastics in sewage sludge. Sewage sludge is a semi-solid material that is produced during the primary stage of wastewater treatment and in some countries this sludge is used as soil fertilizer. Subsequently, microplastics end up into bodies of water and in addition macroplastics and relative large microplastics are exposed to sunlight and other biological factors which in turn causes the plastic to (further) fragment, releasing even more microplastics into environmental waters.29
Another significant contributor to the release of microplastics into the environment is the wear and tear of car and truck tires.30 The global average flow of microplastics into the environment per capita caused
by the wear and tear of car and truck tires was estimated at 0.81kg/year. Other transport related factors that contribute include airplane tires, artificial turf, brake wear and road markings. Moreover, depending on local factors like sewage systems or road type, wear and tear of tires alone contributes to 5 – 10% of the global amount of microplastics eventually reaching the oceans.31 In air, particles from tires smaller
Also, as already touched upon in paragraph 2.1, certain cosmetics (face wash, hand soap, other personal care products) contain high levels of micro-beads. These microplastics are usually composed of polyethylene but other polymers are also imbedded in the beads.21 After use of these cosmetics the
beads are transferred to sewage treatment plants where on average 95 – 99.99% of the micro-beads are filtered from the sewage water. This results in an average of 0 – 7 micro-micro-beads per liter of treated water, in combination with an average discharge of 160 trillion liters of water per day, approx. 8 trillion micro-beads enter the environment every day per treatment plant.32 Furthermore, many
companies have removed micro-beads from their personal care products but there are still around 80 different scrubs that do have micro-beads as their main component. This contributes to the discharge of 80 metric tons of micro-beads per year by the United Kingdom alone, which not only negatively impacts the food chain and wildlife but also toxicity levels since micro-beads can absorb pesticides and polycyclic aromatic hydrocarbons.32 The risks associated with exposure to microplastics are described in more
detail in paragraph 2.4.
Other sources of microplastics reaching the environment are: washing of clothes33, manufacturing of
small pellets for plastic production18, recreational and commercial fishing34, packaging, and shipping.35
For instance, it was found that every piece of clothing could shed up to 1900 microplastic fibers during laundry with fleeces reaching the highest percentage of fibers.
2.3 Distribution of microplastics
Microplastics end up from different sources and locations and through different mechanisms in the environment (see paragraph 2.2); however, there are only four major environmental matrices where microplastics distribute to. These four matrices are the marine environment, freshwater environment, terrestrial environment and atmospheric environment.8
The occurrence of microplastics in the marine environment has been documented for most habitats of the open and enclosed seas.8 Data from remote shorelines to populated coastlines, from deep-sea
sediments to surface water, and from Polar Regions to the equator is available on the distribution of microplastics. However, exact distributions of microplastics in the marine environment are not fully understood. In addition, mainly common microplastics such as high-density polyethylene (HDPE) (0.94 g/cm3), low-density polyethylene (LDPE) (0.91–0.93 g/cm3), polystyrene (PS) (1.05 g/cm3), polypropylene
(PP) (0.85–0.83 g/cm3), and polyamide (PA or nylon) (1.02–1.05 g/cm3) have been the focus of concern.
Other plastics such as polyformaldehyde (POM) (1.42g/cm3), polycarbonate (PC) (1.2 g/cm3), and
polyester (PET) (1.37g/cm3) occur less in literature. Reasons for this are that common plastics are
produced on the greatest scale worldwide and that research mainly has focused on microplastics with a density lower than approx. 1.025 g/cm3 due to floatation. Furthermore, many factors play a role in the
distribution of microplastics in the marine environment. For instance, plastics that are denser than seawater sink to variable depths and may be transported by underlying currents. Another reason for microplastics to sink is biofouling, this is the accumulation of plants, algae, micro-organisms or animals on wetted surfaces.36 Also, stranded microplastics may relocate to seawater again after changes in solar
that most accumulation of (micro)plastics occur in the converge zones of the five subtropical gyres. However, high concentrations of floating plastics debris are present in the North Atlantic, probably as a result of the dense coastal population. For example, up to approx. 5% of all generated plastic waste in 192 coastal countries entered the ocean in 2010; this might not seem a lot but once plastic enters the ocean there is no human control over the plastic, compared to plastic landfills where contact with the environment and migration of the waste is limited.27
Aside the well documented marine environments, microplastics also distribute to freshwater environments.8 However, microplastics in freshwater have not been researched quite as extensively as
seawater until 2010.8,37,38 Most of the plastic debris in sea originates from land-based sources and
transport through rivers may have a big influence in the amount of microplastics reaching the marine environment.8 Furthermore, microplastic concentrations found in freshwater lakes were just as severe as
in seawater and microplastics originating from household laundry and wastewater treatment plants end up in freshwater prior to potential migration to the sea. Compared to seawater, the microplastic content of rivers and lakes is reported to be highly heterogeneous, even within-site.37 The microplastic content of
some European rivers such as the river Rhine, the river Main and the river Danube was measured and in the river Danube the concentration microplastics exceeded the concentration fish larvae.39 Furthermore,
studies on the river Rhine indicated that, especially in the German section of the river, high abundances of microplastics were observed. Average concentrations reached approx. 900,000 particles km-2 for
surface water and 4000 particles kg-1 were found for shore sediment.40 Lastly, microplastics are also
present in sediments; lakeshore sediments from Lake Garda in Italy contained relatively high concentrations (108–1108 particles m-2) of PE and PS microplastics.41 Other microplastics with higher
densities such as polyvinylchloride (PVC) and PET were also found, this underlined the variety of plastics found in shore sediment. Unfortunately all over the world microplastics are being measured in freshwater sediments, up to 466,305 particles km-2 in North America.8,37
Studies on microplastics in the terrestrial environments have been rarely carried out.6 Reasons for this
might be that the ecological research on marine and terrestrial environments is relatively separated so that ideas and/or information are not shared effectively. Secondly, microplastics are relatively easy to separate and analyze from an aqueous sample and not so easy from a complex organo-mineral soil sample. Lastly, marine environments tend to provide habitat for filter feeders, organisms that feed by straining suspended matter and food particles from water, which may increase the risk of microplastics due to enhanced accumulation. Nevertheless, studies on the ingestion of microplastics by terrestrial organisms have been performed and are described in more detail in paragraph 2.4.8,7 Furthermore,
microplastics can enter the terrestrial environments through deposition from the air and irrigation water. In addition, agricultural sites and horticultural fields are a great source of plastic debris and due to the oxygen availability and abundance of sunlight, fragmentation of macroplastics into microplastics is significantly higher than in the aqueous environment.8 As a result, a study in 2018 found significantly
greater abundance of all kinds of microscopic (0.05 – 1 mm) plastics particles (mainly fibers) in cropped areas compared to a forest buffer zone. The study concluded that the ‘’application of soil amendments and irrigation with wastewater must be controlled to reduce accumulation of microplastics in agricultural soils’’.42
Lastly, microplastics can even distribute to the atmosphere.8,9 A study from 2016 that examined outdoor
air in Paris found mostly microscopic plastic fibers in the atmospheric fallout.10 An average of 118
particles m-2 day-1 was found with sizes ranging from 1000+ µm (50%) and 100 – 1000 µm (50%). A year
later the same leading author found microplastic fibers ranging from 50 – 3250 µm in indoor and outdoor air.11 An air pump with 1.6 µm filters was used to sample air and collect the fibers. Overall,
indoor concentrations ranged from 1.0 to 60 fibers m-3, however, outdoor concentrations were
significantly lower, ranging between 0.3 and 1.5 fibers m-3. Nevertheless, more research must be carried
out on microplastics in air due to possible inhalation by humans and especially young children.8
2.4 Risks of microplastics to humans and the environment
Microplastics present in the environment bring certain risks to the health of humans and to the wellbeing of the environment itself.8 For instance, an abundance of microplastics could change the
physical properties of beach sediment; this could affect life in these sediments.43 Also, floating
microplastics on the water could change the spread and therefore bioavailability of sunlight to underlying organisms, affecting their normal life activities. Unfortunately, just mentioned effects of microplastics on the environment are relatively small and less worrisome than the risks that occur when microplastics are ingested since microplastics could be a source and sink of toxic chemicals.8
Furthermore, since 1960 more than 180 kinds of organisms are reported to ingest (micro)plastics, including turtles, fish, mammals and birds.44 Microplastics were found in different tissues such as
digestive glands, gills, stomach and hepatopancreas and translocation of the ingested microplastics from gut the circulatory system and cell tissue was reported.8,45,46 More importantly, microplastics can be
transferred throughout the food chain from lower trophic levels to their predators47.
Next to that microplastics are ingested and transferred easily, they are also potentially a source of toxic chemicals.8 One of the reasons for this is that plastics in general are extensively contaminated with
additives; 50% of the plastics contain hazardous monomers, additives and byproducts.48 Also, several
plastics such as PS, PC, and PVC have been shown to leak toxic monomers that are linked to reproductive abnormalities and cancer.8 Furthermore, a wide variety of additives, such as colorants, plasticizers and
flame retardants are added to plastics during manufacturing in order to improve the performance of plastics.48 However, numerous of these additives are toxic; for instance, brominated flame retardants and
nonylphenol are carcinogens and endocrine disruptors. Lastly, it was reported that 38% of plastics exposed to artificial sunlight produces leachates that caused acute toxicity to Nitocra spinipes49 and that
a fluorescent additive, leached from microplastics, was toxic to Chlorella vulgaris.12
Microplastics could not only be a source but also a sink for toxic chemicals.8 Accumulation of persistent
organic pollutants such as polychlorinated biphenyls, organochlorine pesticides, polyaromatic hydrocarbons and metals from the surrounding environment onto microplastics has been reported from ng g-1 to mg g-1.13,14 Organic contaminant concentrations up to 6 orders of magnitude greater than those
in ambient seawater have been found accumulated by microplastics.50 Moreover, plastics like
polyethylene (PE), PP and PVC have a greater affinity for these compounds than natural sediments.8 In
hydrophobicity and the modification of the surface area of microplastics. On the other hand, aqueous metals are considered to be relatively inert toward (micro)plastics; however, loss of aqueous metals to plastic containers has been reported.14 Furthermore, next to lab experiments, long-term field
experiments have shown that microplastics can accumulate metals.8 Modification of the microplastic’s
surface by adhering organic matter (biofouling, paragraph 1.3) is the main contributor to metals accumulating to microplastics. This organic matter forms complexes with metal ions and interacts with silt and clay that in turn is able to accumulate metals.14 Lastly, it has been reported that presorbed
pollutants on microplastics and additives can transfer into the tissue of animals51 and pollutants have
3.
Overview of the general workflow of the determination of microplastics
Research on microplastics in the marine- and freshwater environment has been performed extensively compared to terrestrial- and atmospheric environments. Sampling and sample preparation of terrestrial samples is similar to those of sediment samples.53 For this reason, the sampling, sample treatment and
identification of microplastics in aqueous environment (including sediment) is described in this chapter. Figure 1 shows possible strategies for the analysis of microplastics in water and sediment.
Figure 1: Possible strategies described in literature for the analysis of microplastics in water and sediment samples.
Starting with sampling to the report of the results; sample preparation is split into pretreatment, density separation and posttreatment. Pyrolysis- or thermal desorption-gas chromatography/mass spectrometry (Pyr-GC-MS, TDS-GC-MS), Fourier transform infrared spectroscopy (FT-IR) and scanning electron microscopy energy-dispersive X-ray spectroscopy (SEM-EDS) are deployed for the analysis. Reproduced from Klein et al., (2018).37
3.1 Sampling
In the aquatic environment an aqueous phase (surface water and a water column) and a sediment phase (lakebed sediments, riverbed or shoreline sediments) are differentiated.37 The abundance of
microplastics in the aqueous phases is significantly lower so therefore larger sample volumes (up to 100 liters) should be taken. Liquid samples from the water surface are filtered mainly by neuston or plankton nets with mesh sizes ranging from 50 – 3000 µm; filtering is supported by a flow meter in order to
measure the sample volume. Mesh sizes <300 µm typically result in clogging, that’s why 300 µm is a standard mesh size used to sample microplastics. Subsequently, microplastics smaller than 300 µm are sampled nonquantitatively. However, methods utilizing filter cascades are developed that result in a size fractionation during the sampling and the reduction of the matrix burden of the small mesh sizes.54 Also,
water from beneath the surface is analyzed for microplastics; the water column is sampled by the acquisition of batch samples or is directly filtered with the use of submersible pumps.37 Recently, a
continuous flow centrifugation device is being developed to counter drawbacks such as filter clogging, contamination and particle size-discrimination and is successfully tested on lake water to sample microplastics.55
Sediment samples are typically divided into samples from the shoreline and from the river- or lakebed; sediment from the marine environment is typically sampled from beaches.37 Standardized sampling
instruments are able to collect bed sediment in a relatively comparable manner so that results are comparable and variation is minimized.56 Also, drills are used for the sampling of deeper layers of
sediment but this results in relatively small sample volumes.37 Throughout literature, shore sediments
are collected perpendicular, parallel or randomly at different distances to the shoreline. Typically grid samples at depths of 2 – 5 cm from the sediment surface are taken. Moreover, stainless steel spoons, trowels or shovels are used for sample collection since it is obviously important to avoid contact with plastic equipment. However, if plastic containers are used for transportation of the sample, blank samples ought to be taken in order to quantify their contribution to the concentration microplastics in the sample. Furthermore, biota samples such as mussel and fish are also analyzed for microplastics. Samples include e.g. digestive residues57, stomachs58, gastrointestinal tract58 or the alimentary tract.59
3.2 Sample preparation
Microplastics in water are easily filtered from the sample during the sampling process, microplastics in sediment samples however must first be separated from the sediment.37 Density separation is a
commonly used technique to separate microplastics from sediment particles. A high density solution (usually salt solutions) is added to the sediment containing the microplastics, this results in the relatively light plastic particles to float; the dense sediment particles do not.60 Different density separation setups
were developed in order to improve the repeatability, effectivity and the ease of handling of this technique.37 Initially Erlenmeyer flasks or beakers were used but those were substituted by separation
funnels, stainless steel separators with high volume capacity and vacuum-enhancement. Another technique to separate microplastics from soil particles is accelerated solvent extraction (ASE); also referred to as pressurized liquid extraction.61 In ASE microplastics are extracted under higher pressure
which increases the extraction speed, this is typically done is small metal cells. Furthermore, ASE does not require an additional sample purification step and is easily automated; controversially, the volume of the extracted sediment is limited due to the relatively small metal cells.37
After the volume reduction, samples are treated in order to remove natural debris.37 Natural debris often
accompanies microplastics during density separation or sampling of water samples and could prevent the identification of microplastics. Therefore, the amount of biological material in samples is minimized in order to prevent underestimation and/or misidentification of microplastics and to minimize
background during measurements. Natural debris is removed from samples by either a chemical- or enzymatic approach, both having their pros and cons. Natural debris is chemically destructed by treatment of the sample with Fenton-like reactions (Fe3+/H
2O2)62, hydrogen peroxide, and mixtures of
sulfuric acid and hydrogen peroxide.37 However, these harsh treatments may result in the destruction of
plastic particles that are unstable in relatively strong acidic conditions or that are relatively easily oxidized. Examples of such plastics are PC or poly (methyl methacrylate) (PMMA). In order to avoid this loss of plastic particles, treatment with sodium hydroxide was proposed; however, it was reported that such treatment might also damage the microplastics.63 Furthermore, research showed that treatment of
samples with potassium hydroxide caused least damage to the plastic content compared to the approaches mentioned above.59 Next to chemical destruction of biological material in soil and water
samples, enzymatically catalyzed reactions are also utilized.37 Cole et al., (2014) developed a treatment
for marine surface water samples using single-enzyme approaches (proteinase K) or mixtures of enzymes to remove natural debris.63 Enzymatic digestion was performed under relatively normal temperature and
pH in order to detect pH-sensitive polymers. However, in comparison to chemical treatments, enzymatically catalyzed reactions might not remove the entire organic content and are expensive and time-consuming.37
3.3 Particle analysis
There are several ways to identify microplastics; in this paragraph three techniques will be briefly explained: scanning electron microscopy (SEM), Fourier-transform infrared spectroscopy (FT-IR) and Raman spectroscopy. Approaches utilizing mass spectrometry (MS) will be elaborated upon in further detail in chapter 4.
SEM
The characterization or analysis of microplastics can be divided into two types: physical or morphological classification and chemical classification.64 Physical classification focuses mainly on the shape, size,
distribution and color whereas chemical classification focuses more on the type of polymer and potential additives. Typically, microscopic, spectroscopic and mass spectrometric methods are utilized to study both the morphological and chemical composition of microplastics. Furthermore, morphology studies can be performed by SEM but this technique does not differentiate plastic particles from natural particles with the use of spectral data. To counter this issue, SEM has been coupled with energy-dispersive X-ray spectroscopy (SEM-EDS); this resulted in high resolution images and elemental analysis of the measured plastic particles.1 Moreover, in SEM-EDS, the surface of the particle is scanned by an
electron beam; as a result secondary electrons and element-specific X-ray radiation are emitted from the surface. These secondary electrons and X-ray radiation can be processed to an image of the microplastic particle and the elemental composition can be identified. Subsequently, SEM-EDS could distinguish microplastics from inorganic particles such as aluminum silicates.
Raman spectroscopy
SEM-EDS is not the only approach that combines microscopy and spectroscopy.64 Raman spectroscopy is
for instance coupled to a microscope, enabling the collection of spectra from distinct points of a microscopic image. Morphological and chemical information is hereby acquired from single microplastic
particles. Furthermore, lasers in the UV-vis range are being used most frequently in the characterization of microplastics and since the spatial resolution is dependent on the excitation wavelength, spatial resolutions in the micrometer range is obtained.65 This, however, limits the particle size that can be
analyzed by this technique. Shorter wavelength could be used but at the same time result in relatively higher fluorescent interferences and higher intensities of backscattered light.64 After measuring, the
recorded spectra are compared to a database in order to identify the plastic particles. Also, since water molecules are weakly active in Raman spectroscopy and it is a non-destructive method, Raman can be coupled with confocal laser scanning. This enables scanning organic tissues for microplastics and is therefore used in toxicology studies for the understanding of the uptake and effects of microplastics on organisms.66 Also, the absence of extensive sample preparation is one of the features that contributes to
the strength of this scanning technique.64 One of the limitations of Raman spectroscopy however is the
presence of fluorescence. Moreover, sorption of organic substances to the surface of plastic particles is a major source of fluorescence and the artificial dyes that are added to the plastics may contribute more to the recorded spectrum than the actual polymer itself. To combat this, more intensive sample preparation should be performed.
FT-IR
Similar to Raman, FT-IR is also used for the analysis of microplastics.64 Either the reflectance or the
transmittance mode of FT-IR can be used but for particles larger than 500 µm the special mode attenuated total reflectance (ATR) is preferred. In this mode the surface of a microplastic particle is put on a crystal; consequently, contamination of the plastic sample and damage to the crystal might be a problem. Moreover, limits of this technique include manual placement of the sample on the crystal which is time consuming and the minimum particle size of 500 µm. Nonetheless, for larger microplastics (in the mm range) this technique is suitable. Furthermore, in water samples microplastics often end up in filters. Even though it is very time consuming and only 10% of the total area is analyzed, with the use of FT-IR or Raman coupled with microscopy filters can be investigated.64 However, focal plane array (FPA)
detectors are often used to measure samples in high throughput or scan entire filters.67 This is possible
due to FPA detectors being composed of multiple detectors, for instance in a 64x64 grid which allows the FPA detector to record thousands of spectra simultaneously. By scanning a filter only for the carbon-hydrogen stretch using such FPA detectors, images of the analyzed filter locating the plastic particles can be produced.
Lastly, for physical and chemical studies on microplastics in environmental samples, FT-IR and Raman are suitable (non-destructive) methods.64 Nonetheless, interferences originating from organic matter on the
plastic surface and typically extensive sample preparations are major things to consider. The difference between Raman and FT-IR lies in the size of the plastic particles that can be analyzed: Raman can measure particles as small as 1 µm whereas FT-IR in the transmittance mode has a lower limit of 10 µm. Using both ATR and transmittance mode when measuring microplastics using FT-IR may be beneficial since larger particles can be put manually on the ATR crystal and smaller particles can be analyzed with transmittance measurements.
3.4 QA/QC and blank issues
Since microplastics are present in air and clothes55, and laboratory equipment includes a substantial
amount of plastics, the risk of contamination is relatively high in microplastic research. This problem is addressed by several (recent) articles and appropriate measures have been taken in order to reduce contamination55,68–70; however, lack of interlaboratory data and reference materials is still affecting data
quality in microplastic research.71,20 Risk of contamination starts at sampling; especially nylon nets and
pumping systems are potential sources of microplastic contamination. Non-plastic alternatives are usually not able to process similar, large volumes of water; which leaves the choice between representative samples and eliminating potential contamination.70 The use of metal spoons, glass
bottles, and thoroughly rinsing of nets are examples of measures taken to reduce risk of contamination during sampling.68 Furthermore, covering of samples after sampling and during processing, and working
in rooms where air-circulation is controlled help reducing contamination.70 Keeping samples covered
during sample treatment and visual identification was shown to reduce contamination by >90 %72
whereas working in a fume hood decreases microplastic contamination by 50 %.73 As for the lack of
interlaboratory studies, the Vrije Universiteit has collaborated with several other organizations in order to fill this data gap by establishing an interlaboratory comparison study that includes microplastic determination by several techniques and provides additional workshops.
4.
Overview of methods for the determination of microplastics using MS
For the determination of microplastics, mass spectrometry is utilized extensively in the last decade and much progress has been made.64 Especially thermo-analytical methods such as thermogravimetry
analysis (TGA), pyrolysis gas chromatography (Pyr-GC) and thermal extraction desorption gas chromatography (TED-GC) are being coupled to MS for the determination of microplastics. However, liquid chromatography (LC) is also coupled to MS and the use of time of flight (TOF) instruments has also been reported. This chapter will discuss abovementioned techniques and review literature on the topic.
4.1 Thermogravimetry analysis-mass spectrometry (TGA-MS)
In TGA, samples are heated in a controlled fashion and the weight of the sample is measured during the heating process.74 Heating can be performed by either combustion or pyrolysis of the sample, both being
thermochemical reactions.75 Moreover, combustion of a fuel can emit light and heat since combustion is
an exothermic chemical reaction (oxidation). Pyrolysis of a sample results in decomposition of the organic material by the addition of heath. The biggest difference between both reactions is oxygen; for combustion oxygen is necessary but pyrolysis is performed in the (near) absence of oxygen. Furthermore, in TGA the sample is placed in a pan on a precision scale; this complex is in turn placed in a furnace.74 The sample can therefore be heated and cooled while the weight of the sample can be
monitored precisely. To control the atmosphere around the sample, a sample purge gas flows through the sample pan; this gas may be oxygen (reactive) or inert (e.g. nitrogen). Upon heating certain compounds in the sample undergo degradation and gaseous degradation products (also named thermal degradation product or pyrolysis products throughout this article) emerge. These gasses can either be captured by differential scanning calorimetry (DSC), MS or GC-MS for the purpose of microplastic analysis.76 In order to transfer a defined amount of degradation gas to the MS, heated capillaries or
capillary-less orifices are used. Then, based on the relation between mass loss at a specific temperature and the recorded m/z by the MS, analytes with similar pyrolysis products as the sample matrix can be measured separately from each other. For instance, soil pyrolysis products and polymer pyrolysis products may have the same m/z but decompose at different temperatures.
TGA-MS has successfully been carried out on soils, sediments and polymers, however, in 2018 David et al., (2018) performed TGA-MS on a complex mixture of microplastic polymers in soil.76 Their goal was to
evaluate a capillary TGA-MS system for the qualitative and quantitative determination of PET in soil samples. Characteristic pyrolysis products of PET such as benzoic acid and biphenyl were measured from standard loamy sand samples spiked with PET. Moreover, two calibration series were prepared: one series without internal standard (IS) by spiking approx. 50 mg of soil with 0.56 – 4.18 wt % PET and one series with IS by spiking approx. 45 mg soil with 0.23 – 4.59 wt % PET. PET dust was obtained by shredding PET bottles and cysteine was used as IS since it provided an easily detectable pyrolysis product (SH-) that was absent in both PET and soil organic matter (SOM) pyrolysis products. Furthermore,
samples were exposed to a 5 K min-1 pyrolysis ramp from 40 to 1000 °C under a 20 mL min-1 argon flow.
acid (m/z 150) of 0.07 and 1.72 wt % PET, respectively. Figure 2a shows the overall, relative mass loss of the sample and the relative abundance of benzoic aicd at different wt % PET during the pyrolysis ramp; figure 2b shows the calibration curve of benzoic acid at increasing wt % PET. At 205 – 250 °C a sharp weight loss is observed, this corresponds to the degradation of the IS. The calibration curve showed an R2 of 0.987 and relative standard error (RSE) of 3.21%. As a result, the authors concluded that even
though lower or equal LOQs and LODs were found with other techniques such as TED-GC-MS, Pyr-GC-MS and LC-MS, TGA-MS is still a very suitable technique for the determination of microplastics (PET in this case) in soil. TGA-MS requires (almost) no sample preparation, is relatively cheap and can handle samples up to 1 g; this is important due to heterogeneity of soil samples. Lastly, TGA-MS analysis for microplastics in soil could serve as a first assessment to investigate if soils are contaminated with microplastics in order to evaluate whether more sensitive methods should be applied.
Figure 2: TGA-MS curves with the relative mass loss of the sample and the relative abundance of benzoic acid at
different wt % PET spiked soil samples (A) and the corresponding calibration curve; the dashed lines represent the 95% confidence interval (B). Reproduced from David et al., (2018).76
4.2 Pyrolysis – gas chromatography – mass spectrometry
Just as the destructive thermoanalytical method TGA-MS, Pyr-GC-MS also utilizes pyrolysis of the sample in order to decompose relatively large analyte molecules into smaller volatile compounds.77 These
thermal degradation products are then separated by GC, measured by MS and structural information about the e.g. polymer is obtained. Moreover, identical samples can be sequentially pyrolyzed at different pyrolysis conditions (e.g. pyrolysis temperature) in order to e.g. extract organic plastic additives (OPAs) prior to measuring the pyrolysis products of the polymers. In general, the increase in temperature is relatively fast, temperatures of 700°C are reached in approx. 10 seconds.78 Lastly, only a small fraction
of the thermal degradation products are lead to the GC-MS and pyrolysis of the sample can be done within the same instrument or is separately performed. The main advantages of Pyr-GC-MS compared to spectroscopic techniques for the determination of (microscopic) polymers is the potential absence of intensive sample pretreatment (measuring untreated environmental samples), the ability to measure OPAs and microplastics in the same run and gain more structural information of the polymer which could aid to the identification of the source and distribution of microplastics.79 The main disadvantages of
Pyr-GC-MS are the high instrumental costs and the relatively large minimum amount of sample (individual microplastic) needed for analysis. Compared to a minimum sample size of 1 µm and 10 µm for Raman and FT-IR spectroscopy, respectively, Pyr-GC-MS needs 50 µm or approx. 0.1 mg of sample for one analysis due to difficulties in manual transfer of the single particle to the pyrolysis cup. However, the difference is that spectroscopic techniques measure generally single microplastics whereas Pyr-GC-MS is able to measure environmental samples containing microplastics. Nevertheless, such small samples are not sufficiently representative of a heterogeneous environmental sample containing multiple kinds of microplastics. Lastly, no physical information is obtained such as the morphology when microplastics are analyzed by Pyr-GC-MS. Columns such as HP-5ms ((5%-Phenyl)-methylpolysiloxane) and DB-5ms (Phenyl Arylene polymer virtually equivalent to a (5%-Phenyl)-methylpolysiloxane) are typically used; ‘’ms’’ stands for a low-bleed version of the HP-5 and DB-5 columns.
The beginning
Pyr-GC-MS has been used for the determination of microplastics for at least almost two decades.80 In
2000 Fabbri et al., (2000) proposed a Pyr-GC-MS method to determine PVC particles (1 – 10 µm) in sediments and suspended particulate matter (SPM). Also, an alternative method using Soxhlet-extraction and precipitation to isolate the polymer from the sediment was described. 10 g of wet sediment was mixed with 10g of anhydrous sodium sulphate and DCM and extracted for 40 h (extraction yields remained constant after 16 h), afterwards DCM was evaporated and the residue was re-dissolved in DCM and added drop-wise to n-hexane, precipitates were dried and analyzed. Furthermore, Pyr-GC-MS analysis was performed on SPM (dried matter retained on filters; 5 mg), isolated polymers (0.2 mg) and sediments (13 mg) by pyrolizing the samples at 700 °C for 10s and the highest heating rate. Ions were measured by an ion trap instrument using scanning modus; total ion chromatogram (TIC) and extracted ion chromatogram (EIC) were used for identification. As a result, recoveries of 75 – 95% were found for the extraction, the relative standard deviation (RSD) for benzene (a pyrolysis product of PVC) was reported at 14% and the LOD for PVC in isolated polymer samples was 0.1 mg/g whereas the LOD for PVC in sediment samples was 0.01 mg/g. Pyrograms from sediment analyses contained numerous peaks originating from thermal decomposition products of organic matter but the peaks arising from synthetic polymers were easily identified. Predominantly aromatic compounds such as toluene, benzene, ethylbenzene and styrene were observed; these are known thermal degradation products from PVC and other polymers. Moreover, the presence of chlorobenzene in the pyrolysate confirms the presence of PVC in the sample; however, the concentration of chlorobenzene was too low to be used in quantitative calculations. Instead, Fabbri et al., (2000) used benzene for quantification. Also, matrix effects such as organic matter and co-occurrence of styrene plastics (since PVC also contributes to styrene in the pyrolysate) were considered during quantification by also measuring calibration plots in unpolluted sediment. Lastly, concentrations PVC found by direct pyrolysis of sediment samples compared to isolated PVC from sediment samples compared reasonably well with each other based on orders of magnitude and distribution. However, differences up to four times were found. In 2001 Fabbri et al., (2001) determined several other kinds of microplastics such as polybutadiene (PB), poly(vinyl acetate) (PVA), poly(acrylonitrile-co-styrene-co-butadiene) (ABS), styrene–butadiene block copolymer (SBS) and styrene–butadiene rubber (SBR) in sediments using a similar method.81 Lastly, these methods are used
for untargeted microplastic determination instead of specific polymer analysis; if more literature or larger databases were available that time, more sufficient identification could potentially be performed.
Polymers and additives
In 2009 Trimptin et al., (2009) identified polymer type and additives of small pieces of plastic such as pieces of PET thread or Nylon.82 The aim of the study was to circumvent drawbacks like labor intensity
and time cost of methods such as TGA-, (Pyr)GC-, or LC-MS by developing a relatively rapid analysis. Furthermore, samples (1 mm of plastic thread) were either put into an atmospheric solid analysis probe (ASAP) and heated to maximally 500 °C or put into a pyroprobe coil to be heated to either 450 °C or 700 °C. The ASAP method mainly identified additives present in the plastic whereas the higher temperature of the pyroprobe was necessary in order to identify the polymer; volatile thermal degradation products were ionized by atmospheric pressure chemical ionization (APCI) in both methods. Moreover, the ASAP method was able to analyze several additives in less than a minute by raising the temperature from 100 to 400 °C. However, when the temperature was increased to 500 °C ions from low mass oligomers are produced. A cyclic trimer of PET was observed at 500 °C but it was not certain whether this trimer was an actual degradation product of PET or originated from residual trimer that had vaporized, since residual trimer leached from PET was measured in bottled water by applying 5 µl of bottled water on the ASAP. In addition, a piece of a nylon fiber was analyzed at 500 °C by the ASAP method and the pyroprobe method at 700 °C. At 500 °C mainly cyclic oligomers from the mono- to the heptamer were measured whereas at pyrolysis at 700 °C produced a higher abundance of lower mass fragments (monomer) compared to the ASAP method. Nevertheless, both methods produced ions that clearly identified the polymer as nylon-6. Lastly, the most time-consuming step was placing the sample on the melting point of the ASAP or on the pyroprobe platinum wire coil. Furthermore, due to small sample size and possible heating to 1000 °C contamination was limited. Even though these methods provide fast measurements of small polymer particles and its additives, the analyzed individual microplastics did not originate from an environmental matrix so the method should be tested on microplastics contaminated with organic matter. In that case, sample preparation would probably be necessary.
Streams and digestion
In 2016 the microplastic content of up- and down streams of several wastewater treatment plants was investigated by McCormick et al., (2016); additionally the bacterial assemblages on the microplastics were determined.83 One liter surface water was sampled using nets with a 333 µm mesh size and
microplastics were picked out using forceps and transported in 20 ml site water; pellets, fibers, and fragments were the dominant microplastic types. These 20 ml samples were filtered (0.33 – 4.75 mm) and dried in an oven at 75 °C, additionally the organic material adsorbed on the microplastics was degraded by wet peroxide oxidation; wet peroxide oxidation resists plastic but degrades organic matter. Next, microplastics were separated by density separation and filtered again before being loaded into the pyroprobe and pyrolyzed at 750 °C for 90 s and analyzed by GC-MS. Furthermore, an ion trap MS was used to detect all m/z between 34 and 551 in scan mode and pyrograms were obtained for every sample by averaging mass spectra over the chromatogram. Obtained pyrograms for each microplastic were compared to pyrograms from the CDS Analytical 2013 pyrolysis library and the best match was used. As a result, PE, low density PP (LDPP), PS and ethylene were identified among the microplastics. This method
shows adequate qualitative measurements of different types of microplastics; however, sampling and sample preparation are quite labor intensive and further methods specifications such as LOD/LOQ are not given. Also, only using a database for identification may not be sufficient as literature and standards could be used as well.
Dehaut et al., (2016) similarly performed qualitative analysis on microplastics utilizing Pyr-GC-MS in 2016.59 They tested several protocols to digest organic matter on the deleterious effects on fifteen plastic
types. Aside deleterious effects, the effect of digestion on the ability to identify the individual microplastics by Pyr-GC-MS was also determined. Furthermore, spiked samples of mussel, crab, and seabream were digested, filtered, and density separated; single microplastics (<0.5 mm3) were put in a
pyrolysis cup and heated to 600 °C before being directly injected in the GC-MS that scanned from m/z 33 to 500. Average mass spectra were compared to an external and internal database; no use of EICs of indicator ions was reported. Nevertheless, this resulted in a majority of the plastics being correctly identified by Pyr-GC-MS (>80 % match with database); however, identifying cellulose acetate caused some problems after most of the digestion protocols since the corresponding pyrogram was identified as ‘’wood powder’’. To conclude, Dehaut et al., (2016) evaluated the identification of fifteen types of microplastics by Pyr-GC-MS after several different digestions. This shows Pyr-GC-MS being used as a suitable technique to determine a wide range polymer types of single microplastics after sample preparation.
Fish
In 2017 Fischer and Scholz-Böttcher, (2017) performed a rather extensive study on the qualification and potential quantification of microplastics in fish samples; they additionally compared a direct pyrolysis method with a thermo-chemolysis (methylation under pyrolytic conditions)84 method.58 PE, PP, PS, PET,
PVC, PMMA, PC, and polyamide 6 (PA6) were simultaneously detected within a single GC-MS run. As for the thermochemolysis, tetramethylammonium hydroxide (TMAH) was added to the samples for the thermochemolysis directly prior to pyrolysis. For identification polymer standards were measured either with or without TMAH and sample pyrograms were compared to an in-house database and to literature. For calibration, 0.4 – 1070 µg of standard polymer was weighed into Al2O3 and measured individually and
in mixtures and identified utilizing indicator compounds (characteristic pyrolysis products). For recovery experiments, fish samples (stomachs and gastrointestinal tracts) of 20 g were spiked with several kinds of plastics and enzymatically and chemically digested in order to remove organic content; then, samples were dried and degreased. If the sand content was too high additional density separation was performed. Dried samples were then milled in a small mortar and placed into the pyrolysis target cup and pyrolyzed at 590 °C directly or after adding TMAH. As a result of the TMAH, pyrolytic mechanisms changed from e.g. random- or end chain scission to thermochemolytic transmethylation and subsequently the main signals changed to signals from methylated compounds. As a result, the addition of TMAH resulted in more specific reaction products which enhanced the detection sensitivity for PET and PC. As for the sensitivity of the method, LODs in the calibration were determined at 0.4 µg. No significant losses during sample pretreatment or altered indicator signal intensities were observed in spiked fish and PVC, PMMA, and PS were qualitatively determined in non-spiked stomach samples. Additionally, several sources of naturally occurring polymers that could generate indictor compounds
that could interfere with indicator compounds of synthetic polymers were pyrolyzed in order to determine their interference with microplastic determination. Chitin, cellulose, pine wood, wool and cotton were pyrolyzed under addition of TMAH and Fischer and Scholz-Böttcher, (2017) found that the indicator compounds of PP, PET, PA6, and PMMA were not affected by these biopolymers. To conclude, this method showed to be very sensitive and no significant losses were observed during sample preparation of the fish samples. In addition, for the first time three types of microplastics were determined in solid environmental stomach samples; further research could potentially include calibration in stomach matrix in order to quantitatively measure microplastics in fish samples. Also, for the first time the use of standard polymers for identification has been reported which improved identification.
Lakes, ingestion, bivalves, sediments, and solvent-solubility
In order to quantify the distribution of microplastics and identify the microplastic content of a lake in the USA, Hendrickson et al., (2018) applied Pyr-GC-MS (and ATR-FTIR) on microplastics extracted from aqueous samples.68 Surface microplastics were sampled from surface water using nets with mesh size of
333 µm; extra caution was given to prevent contamination during sampling and transport by additional rinsing of petri dishes and wearing wool or cotton based clothes instead of synthetic fibers. As for pretreatment, samples were dried, oxidized to remove organic matter and density separation was performed to separate plastic particles from inorganic matter such as sand. Lastly, pretreated samples were filtered and examined by microscope; particles identified as microplastics were analyzed by Pyr-GC-MS. Moreover, particles <20 µg were analyzed by splitless introduction and particles >20 µg were introduced in the GC using a 1:100 split; the MS scanned m/z 10 – 550. Pyrograms were firstly compared to a mass spectral library, secondly compared to plastic standards and thirdly compared to literature. If less than 3 or 4 pyrolytic products were measured or in too low abundance, peaks of pyrolytic products were integrated in the pyrogram and their presence was established, similarly as Fischer and Scholz-Böttcher, (2017)58. As a result, 14 out of 68 analyzed particles showed no signal in the 10 – 550 m/z range
and 3 particles could not confidently be assigned to a single polymer; the rest of the particles were identified. The authors concluded that for reliable microplastics identification more than one specific pyrolytic product should be present in sufficient abundance, therefore, if this is not the case FTIR should be used complementary to Pyr-GC-MS. Furthermore, unassigned particles could also be misidentified as plastic particles by microscopy, hence no polymer type could be assigned. The successfully assigned particles belonged to PVC, PP, PE, PET, chlorinated PE (cPE), PS, and polydimethylsiloxane (PDMS). Aside some unassigned particles, Hendrickson et al., (2018)68 applied a suitable Pyr-GC-MS method for the
identification of seven different microplastic types in aqueous environmental samples.
In the same year Peters et al., (2018) used the abovementioned Pyr-GC-MS method to identify microplastics ingested by 1381 fish from six different species.85 Stomach contents were washed with
distilled water and filtered through filters of 1000 µm, 243 µm, 118 µm and 53 µm. After filtering, microplastics were identified utilizing microscopy and then analyzed by Pyr-GC-MS; however, particles <10 µg were introduced in the absence of a split flow whereas Hendrickson et al., (2018)68 used a split
flow above 20 µg. The following standards were used for identification: medium density PE (MDPE), PVC, PS, and PET. Furthermore, 30 particles including fibers, fragments and spheres were analyzed by
Pyr-GC-MS resulting in the identification of PVC, PET, silicone, epoxy resin and nylon. Nevertheless, 42% of the analyzed particles could not be assigned to a polymer class but 50% of these unidentified particles shared a very similar pyrogram consisting of common pyrolysis products, indicating a similar composition and origin. In figure 3 the pyrogram and pyrolytic products of PVC and an unidentified sample are shown. It was hypnotized by the authors that the unidentified samples could originate from the petroleum industry, as pyrolytic product diethyl phthalate (a plasticizer) was measured and not all particles shared the same morphology. To conclude, this study showed that Pyr-GC-MS is a suitable technique for the identification of microplastics in a fish ingestion study, and in comparison to Fischer and Scholz-Böttcher, (2017)58, significantly less sample preparation is required. However, constraints such as time cost for
sample preparation and analysis, significant decrease analysis success for particles <10 µg, not representable samples when multiple types of microplastics are present, and lack of comparative standard data are sources of shortcomings of this method. Lastly, Fisher et al., (2017) pyrolyzed dried fish samples whereas this method analyzed single microplastics; this leaves a lot of smaller particles undetected.
Figure 3: Pyrogram and pyrolytic products of PVC (A) and Unknown Subsample A (B). Reproduced from Peters et al.,
(2018).85
Also in 2018, Ceccarini et al., (2018) performed a rather large study on the determination of organic solvent soluble, free and adsorbed microplastics (<2 mm) and their degradation products.19 Microplastics
were not only determined by Pyr-GC-MS but FT-IR, 1HNMR spectroscopy, and gel permeation
chromatography were also used. Furthermore, from each sample spot 1.5 kg sand was sampled in glass vessels, samples were homogenized, sieved at 5 mm, dried at 60 °C, sealed and stored, then further sieved at 2 mm before performing solvent extraction. Microplastic particles within the 2 – 5 mm range were picked out and stored separately. The aim of the solvent extraction of the treated sand samples was to extract polyolefins (e.g. PE and PP) and most vinyl polymers (e.g. PE, PP, PS, PVC, Polyvinyl acetate
(PVAc), and polyacrylonitrile in two separate extractions: refluxing DCM and xylenes at boiling point. Moreover, 160 g of dried sand was punt in cylinder extracted with 90 mL refluxing DCM for three hours at 37 °C. Then, extracted sand samples were again extracted with refluxing xylenes at 135 – 140 °C in order to collect the DCM insoluble, less degraded polyolefin fraction. These two extractions were carried out in triplicate for every sand sample. After extraction the resulting solutions were put in a rotary evaporator and then evaporated to dryness under a nitrogen stream. Pyr-GC-MS was performed at a 650 °C pyrolysis temperature, 1:10 split injection, and using a single quadrupole. Pyrograms were compared to literature and internal and external mass-spectral databases in order to identify pyrolysis products and the interpretation of pyrograms. As for the xylene extracts, gravimetric analysis was carried out and only a nearly negligible amount of non-fragmented, non-oxidized, relatively high molecular weight, intact polyolefins were found. This was explained by the authors by the fragmentation of the most common polymers into micrometric and submicrometric particles with increasing polarity, promoting their solubility in DCM. Furthermore, analysis of the DCM extract by Pyr-GC-MS found markers that related to PS and PE based on literature and internal databases. PE was identified based on its typical pyrolytic profile consisting of a series of clusters corresponding to linear hydrocarbon fragments whereas PS was identified by the given that all markers of PS feature a tolyl ion fragment. The lowest concentration polymer containing solids in the sand samples extracted by DCM and gravimetrically measured was 5.9 ± 0.4 mg/kg sand; for the xylene extracts polymers as low as 0.9 ± 0.3 mg/kg sand were quantified. To conclude, total amounts of polymer containing solids (<2 mm) was determined by TGA and not by Pyr-GC-MS but further identification of the present polymers was successfully carried out by Pyr-Pyr-GC-MS. Also, discrimination between different microplastics in terms of solubility in the extraction solvent may occur so recovery experiments utilizing suitable internal standards would be advisable. Nevertheless, PE and PS were identified in extracts from sand samples; this was the first time terrestrial samples were analyzed
Table I: LOD for eight common polymer types and their corresponding theoretical size of different morphologies.
Reproduced from Hermabessiere et al., (2018).86
Another promising study was done by Hermabessiere et al., (2018) they optimized the method of Dehaut et al., (2016)59 and applied the method to unknown environmental samples, including bivalves.86
