Introduction to the AnalyticalMethodologies for the Analysisof Microplastics

João Pinto da Costa and Armando C. Duarte

ContentsIntroduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2Sampling and Sample Handling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

Marine Water Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3Freshwater and Estuarine Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7Soil and Sediment Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11Biota Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14Atmospheric Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16Sample Handling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17

Extraction/Separation of Microplastics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19Identification of Microplastics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22Recommendations, Knowledge Gaps, and Future Venues of Research . . . . . . . . . . . . . . . . . . . . . . . . . 25References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

Abstract

Ubiquitous and highly pervasive microplastics have been found in all compart-ments of the environment. However, correctly assessing their prevalence isgreatly hampered by the current methodological and technical limitations, aswell as data reporting and analysis. Herein, we discuss the most pressing issuesassociated with the analysis of microplastics in environmental samples (water,sediments, and biological tissues), from their sampling and sample handling totheir identification and quantification. Furthermore, the need for analytical qualitycontrol and quality assurance associated with the validation of analyticalmethods, including the use of reference materials for the quantification of micro-plastics, is also examined. Lastly, the current challenges within this field ofresearch and foreseeable routes to overcome such limitations are discussed.

J. P. da Costa (*) · A. C. DuarteCESAM and Department of Chemistry, University of Aveiro, Aveiro, Portugale-mail: jpintocosta@ua.pt; joao.pinto.da.costa@gmail.com; aduarte@ua.pt

© Springer Nature Switzerland AG 2020T. Rocha-Santos et al., Handbook of Microplastics in the Environment,https://doi.org/10.1007/978-3-030-10618-8_1-1

1

Keywords

Microplastics · Analytical chemistry · Pollution · Environment

Introduction

In recent years, plastic pollution and its consequences have become major causes ofconcern by scientists, policy-makers, and the public, in general. Owing to theexponentially growing production and use of these materials, plastic has becomepervasive throughout the different spheres of the environment. Consequently, plasticparticles have been in areas as remote as Antarctica (Reed et al. 2018) and theMariana trenches (Peng et al. 2018) or isolated, inhabited areas (Allen et al. 2019).This is not surprising, considering that the global production of plastics surpassed359 million tons in 2018, of which approximately 40% was intended for packaging(PlasticsEurope 2019) and, therefore, for immediate (or nearly immediate), disposal.Plastic pollution is therefore ubiquitous, and although the effects of this pollution aremore evident for larger fragments, such as the ingestion of plastic materials byseabirds or whales or the entrapment of seals or sea turtles, smaller particles,commonly referred to as microplastics, could perhaps be more pervasive andconstitute a more prominent risk towards global environmental health and, ulti-mately, to human health (da Costa et al. 2016). Microplastics are commonly definedas plastic particles <5 mm, and, albeit the generally accepted definition, it should benoted that other size-based definitions have been proposed, varying between<1 mmand <10 mm (da Costa et al. 2017). Microplastics (MPs) accumulate in the envi-ronment due to either their direct release or through the fragmentation of largerparticles and are thus classified as primary or secondary MPs, respectively. Thescientific community has devoted particular attention in identifying, quantifying, anddetermining the potential effects of these materials once they reach the environment,and, although this interest has surged in the last decade, all of the currently describedrisks have been identified since the 1970s, when Carpenter and co-workers firstreported the presence of microsized plastic particles in the environment and imme-diately identified their associated hazards and potential consequences (Carpenteret al. 1972). These included not only the ingestion of these materials by numerousorganisms, similarly to what is observed for larger plastic debris, but also the likelyadsorption of persistent organic pollutants (POPs) found in the ambient water andthe colonization of these particles by microorganisms. The ever-expanding array ofapplications of plastic materials, potentiated by the use of numerous additives aimedat improving their characteristics, such as malleability, light and heat stabilization,antioxidant properties, and mechanical resistance, among others, has led to addi-tional risks, such as the already described leaching of these additives to both theenvironment and biota (Koelmans et al. 2014).

It is hence not surprising that a large amount of information about microplastics,namely, their prevalence, risks, and effects, has been gathered and made available inrecent years. Nonetheless, there are obvious discrepancies and disparities regarding

2 J. P. da Costa and A. C. Duarte

the presence of these materials in the environment and a noticeable lack of consis-tency in the different reported field studies (da Costa et al. 2017). It is clear that, toadequately determine the real hazards of microplastics, there is a pressing need todevelop and implement standardized protocols for sampling, quantification, andcharacterization of microplastics, including data treatment and visualization (Silvaet al. 2018), as well as quality control assurance. Only then it will be possible tocompare data gathered from different studies and, from such harmonized data,adequately gauge the potential effects of MPs.

In the following sections, the analytical techniques for sampling, sample han-dling, identification, and quantification of microplastics in different environmentalmatrices, namely, water, sediments, and biological tissues, are described. The ana-lytical quality control and quality assurance protocols associated with the validationof these analytical methods are also considered, including the use of referencematerials for the quantification of microplastics in defined environmental samples.The challenges in sampling and quantification of microplastics are also highlighted,and potential routes to overcome such hurdles are also discussed.

Sampling and Sample Handling

The foundation for any meaningful and representative study is the choice of sam-pling location, while considering the types of samples to be collected and theirpreservation, the spatial and temporal variation, as well as the available samplingtechniques. This is of special relevance for microplastics, as not only do thesematerials do not distribute uniformly throughout the different environmental matri-ces, they may undergo chemical and physical modifications in the environment, andthere are still no specifically standardized methods for the sampling of these mate-rials (Stock et al. 2019). Furthermore, due to aerial fallout, background contamina-tion should always be considered during the process. The following sections detailthe sampling and sample handling in distinct environmental compartments and someof the closely related issues associated with each of these partitions.

Marine Water Sampling

The vast majority of studies regarding the presence, fate, and effects of microplasticsare focused on marine waters. Hence, it would be reasonable to assume that, atleast for this medium, sampling and sample handling standardized protocols doexist. However, despite the existing large body of research, this is not the case.Subsequently, ensuing studies, particularly those delving into the potential (eco)toxicological effects of these materials, are severely impacted, owing to the littleconsistency between microplastics found in the field and those used in laboratoryexperiments. Multiple efforts have been devised to overcome this limitation, andspecific recommendations for the sampling of the sea surface and the water columnhave been issued by the Marine Strategy Framework Directive (MSFD) Technical

Introduction to the Analytical Methodologies for the Analysis of. . . 3

Subgroup of Marine Litter (EU-JRC 2013), including the need to quantify all“microplastics in the size range 20μm to 5mm.” However, the Technical Reportalso lists the possibility of using either neuston nets or manta trawls, depicted inFig. 1, for sampling surface water (e.g., Gajšt et al. 2016), strategies that have beenreported to yield vastly different results (Barrows et al. 2017). The same TechnicalReport states that, for floating marine debris, the unit reporting should beitems�km�2. However, this recommendation is highly debatable, as sampled oceanareas will display inherently different physical parameters throughout the year andseason and, frequently, throughout the day, including winds and currents, which mayaffect the total filtered volume and, consequently, the number of items per area.Hence, whenever possible, the total area and sampled volume should be reported.

Given the relatively low concentration of microplastics in water (Cole et al. 2014;Löder and Gerdts 2015), sampling of microplastics usually requires large volumes ofwater. Therefore, reduced volume approaches are needed. This strategy allowssampling of large volumes relatively quickly into a small, concentrated final sample.As plastic particles are generally buoyant, most researchers resort to surface sam-pling for investigating the occurrence of these materials, and, although there is noclear definition of what constitutes “surface water layer” when sampling micro-plastics (Crawford and Quinn 2017), this is generally considered the top 15 cm,which is where 95% of small plastic debris is concentrated (Carson et al. 2011).

Net type and design, including aperture, length, and mesh size, are all character-istics that influence the sampling process and, ultimately, the reported concentrationsof microplastics. In most cases, the net opening is not provided in publications, butthis usually ranges from 5 to 200 cm, while net lengths are commonly 350–400 cmlong (Liu et al. 2020). Typically, mesh sizes vary from ~50 μm to 3 mm (Crawfordand Quinn 2017), although the vast majority of studies are conducted using plank-tonic nets, i.e., with a 333 μm mesh size. Flow rate can also influence sampling, andit is a necessary parameter to adequately quantify the filtered/sampled volume, whichcan be determined using Eq. 1:

V ¼ πr2d ð1Þwhere V is the filtered water volume, r is the radius of the net aperture, and d is thetowed distance. A flowmeter may be used to accurately measure the towing distance,by multiplying the shown number of revolutions by the pitch of the impeller. Themost commercially successful flowmeters have an impeller pitch of 0.3 m perrevolution, meaning that the towing distance may be calculated by multiplying thenumber of revolutions by a factor of 0.3.

Of note is the possibility of net clogging, which may introduce errors, and the useof two flowmeters is highly recommended. The first should be placed outside the net,to estimate net speed, and the second is best positioned midway the center and the netrim (Crawford and Quinn 2017). The combination of the two readings thus allowsfor a better indication of filtration efficiency and potential net clogging. Carefullyconsidering all these factors may yield an accurate quantification of microplasticsin a given sample. Nonetheless, results of a recent meta-study suggest that the

4 J. P. da Costa and A. C. Duarte

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Introduction to the Analytical Methodologies for the Analysis of. . . 5

commonly used 300–350 μm mesh size nets are undoubtedly underestimatingmicroplastics’ concentrations anywhere between one and four orders of magnitude.Based on data gathered from 46 studies, the authors concluded that microplasticconcentration approximated a negative log-linear relationship with increasing meshsize (Covernton et al. 2019). This was the case even for net tows, in whichparticularly large volumes of water were filtered, and, whether this is due to theuneven distribution of microplastics in a given volume of water or due to method-ological factors remains unclear. Notwithstanding, a compromise should be reached,and sampling methodology must mirror the research questions. It is clear, however,that for a reliable assessment of the prevalence of microplastics and their potentialecotoxicological effects, it is necessary to at least pair filtration methods (filters<10 μm), able to retain narrow microfibers, with large volume towing methods.Only then will it be possible to accurately execute laboratory ecotoxicologicalstudies that better reflect the physiological response of organisms exposed to envi-ronmentally relevant concentrations of microplastics (da Costa et al. 2017).

Freshwater and Estuarine Sampling

Estuaries, transition areas between freshwater and marine waters, are widely con-sidered as zones of accumulation, or hotspots, of microplastics. This is not surpris-ing, as estuaries are generally in or in the vicinity of urban areas and are subject toboth marine and freshwater influences, including tides, inward waves, and outwardflow of freshwater and sediments in a partially enclosed area (Pritchard 1967).Nonetheless, and despite numerous research on different estuaries across theglobe (e.g., Arias et al. 2019), the fact is that the characterization of microplasticsin estuarine ecosystems remains incomplete, given that the inherent unique charac-teristics of each estuary, including geographical localization; topography; localhydrology; prevailing weather, as wind and rainfall; and surrounding and upstreamanthropogenic activities, greatly influence the distribution, transportation, accumu-lation, and, ultimately, the effects of microplastics in these systems. Lima andcolleagues, for example, found that the density of microplastics was comparable tothe density of fish larvae in the Goiana estuary (Brazil) and that it could, in fact,surpass that of ichthyoplankton (eggs and larvae of fish) (Lima et al. 2014). This wasconfirmed in another location, across the Atlantic, where researchers found anaverage ratio of 1.5 microplastics for each fish larvae (Rodrigues et al. 2019).Such observations highlight the potential for ingestion of microplastics byplanktivorous organisms, as was noted at the Charleston Harbor Estuary, wherenumerous organisms, including the bay anchovy (Anchoa mitchilli) and Atlanticmenhaden (Brevoortia tyrannus), were found to ingest microplastics, which werefound in ~99% of fishes, with an average of 27 microplastic particles per fish and21 microplastic particles per gram of digestive tract (Parker 2020). Interestingly,the authors found that tire wear particles were present in 14.4% of individuals acrossall the surveyed species, the impact of which has been modeled by Unice andcolleagues (Unice et al. 2019a). These researchers estimated that tire and road

6 J. P. da Costa and A. C. Duarte

wear particle mass release in the Seine watershed was 1.8 kg�inhabitant�1�year�1.Furthermore, the model estimated that 18% of this release was transported tofreshwater and 2% was exported to the estuary. Subsequent studies emphasizedthat, however, the accuracy of the model greatly depended on the precise in situcharacterization of the physical characteristics of these particles, namely, diameterand density (Unice et al. 2019b).

It does not appear that these observations may be transposed across all estuaries,although the tidal cycle has been suggested to be of particular relevance and shouldbe considered as a factor when choosing time/season for sampling (Crawfordand Quinn 2017). Sadri and Thompson (Sadri and Thompson 2014) reported asignificant shift towards smaller plastic sized particles during the neap tide, whileWu and colleagues (Wu et al. 2020) observed the exact opposite, larger particlesduring the neap than the springtide period. Though contradictory, this may beattributable to the types of microplastics collected, as, in the former study,researchers observed a higher percentage of polyethylene and polystyrene fragmentswhile the, in the latter, authors found that all samples were comprised of >92% offibers. This may impact how some particles may leave the estuary during the ebb tideand reenter the same zone during flood tide.

When collecting samples in estuaries, depending on the research focus at hand, itmay also be important to consider how samples are preserved and the sterility ofmaterials and utensils used. If evaluating microplastic-associated microorganisms,tools such as sterile forceps and immediate storage at �20 �C are needed. This isbecause, despite the yet limited body of research on this topic, it has been shownthat microbial populations associated to microplastics in estuarine environmentsdiffer substantially from those in the natural environment (Jiang et al. 2018), withthe concomitant potential for the presence of putatively pathogenic species.Polyethylene microplastics have been shown to selectively enrich not only micro-organisms but also antibiotic-resistant genes and antibiotics in rivers, estuaries, andmarine waters (Wang et al. 2020b). Though the authors did observe an increase inthis enrichment with decreased salinity, the associated partition mechanisms remainunclear.

Freshwater systems, including lakes and rivers, are equally influenced by thesame physical parameters, namely, currents and wind, that may influence the distri-bution and accumulation of microplastics (Crawford and Quinn 2017). However,given that these are usually smaller systems, the influence of these parameters isgenerally greater, which may result in more pronounced temporal and spatialdifferences in the patterns of distribution of microplastics. This distribution maybe further impacted by the types of transport occurring in these systems, as advectivetransport (velocity) and dispersive transport (turbulence) may also affect micro-plastic distribution (Crawford and Quinn 2017) and such transports are severelyinfluenced by hydrogeological heterogeneities. Another key difference is that fresh-water systems have a closer proximity to point sources in freshwaters. Takentogether, these stark differences in relation to marine waters may lead to differencesin the types of microplastics present. In fact, riverine systems, for example, mayshow a predictable pattern in the characteristics of microplastics present, such as

Introduction to the Analytical Methodologies for the Analysis of. . . 7

size, shape, and relative abundance, based on waste sources (e.g., household orindustrial) adjacent to the river and downstream distance point source from pointsources (Eerkes-Medrano et al. 2015). These are not the only factors affecting thedistribution of microplastics in rivers, as, for example, tributaries may also play animportant role, given that, when tributaries enter the main flow, a segregated flowmay be present over considerable distances before complete mixing is achieved(González et al. 2016). All these factors actually account for the major difficultiesassociated to river sampling, i.e., the necessary sufficient temporal and spatialcoverage that supports representative estimations of fluxes of litter. Consequently,sampling of riverine waters will inevitably depend on the available informationregarding population density, potential point sources, and sampling location avail-ability. Additional considerations may include sampling of upstream and down-stream points of human pressure (e.g., industry or agriculture), although furtherconsiderations, such as administrative borders, may be included (González et al.2016).

In lakes, the residence time can also affect the prevalence of microplastics, as wellas the seasonal rainfall variations, as noted by Moore and colleagues, whom foundthat rainfall resulted in a substantial increase in the total amount of microplastics incollected samples (Moore et al. 2011). This may be of special relevance in tropicalareas, in which the heavy rains may affect the overall microplastic concentration.Lastly, because freshwater environments typically contain large amounts of organicdebris, particularly from vegetation, sampling may be affected, and therefore adiscerning and careful consideration of the sampling tools used should be conducted,as clogging may occur. Table 1 summarizes typical monitoring methods for thecollection of litter in river waters and the applied methodologies.

Most of the described methods are operationally defined, meaning that the chosenmethodology directly influences the result and its reporting (González et al. 2016).Therefore, disparate unit reporting results are used, including, but not limited to, dataexpressed as concentrations, such as mass per volume; items per volume; fluxes,namely, estimations including mass per day, hours, or year; or area density (mass/items per area). Consequently, these methodologies do not deliver the InternationalSystem of Units (SI) traceable results, and a need for harmonization exists tocompare data. For this, agreed-upon methods are required which should includethe agreement at international levels, including River Commissions, Regional SeaConventions, and the European Union and United Nations (da Costa 2018) availableto all. Hence, detailed documentation of sampling and, when applicable, of thesubsequent analysis procedure(s) is needed and should cover the following infor-mation (adapted from González et al. 2016):

(a) Sampling method, compartment, and size classes sampled(b) Sample size/volume(c) Sampling frequency, timing, and replicates. Particularly for estuarine environ-

ments, should include tide information(d) Sampling equipment (all characteristics should be listed)(e) Sample location and river morphology (should cover different sampling sites)

8 J. P. da Costa and A. C. Duarte

Table 1 Sampling methods in freshwater systems. Used devices and monitoring methods aredescribed, as well as monitoring depth (when available)

LocationDevice/toolcharacteristics Method

Monitoringdepth Observations Ref.

Rhine-Meuse-Scheldt delta(Netherlands)

Manta trawl Trawled for amaximum of30 mins

Top 10 cm Opening:n.a.Mesh size:3.2 mm

(Van derWal et al.2015)

Stationarypump-mantanet

Activepumping of5000 L

Located at30 cmdepth

Mesh size:330 μm

Dalålven(Sweden),Rhine(Netherlands),Po (Italy), andDanube(Romania)rivers

Cage-likesetup

Constructionmounted on apontoon withtwo nets whichsamplefloating andsuspendedlitter

Top 10 cm;10–60 cm

Opening:60 � 10 cmMesh size:330 μm

(Van derWal et al.2013)

Dynamicmanta trawl

Lakesampling:distance 3–4 km to filter320–430 m3 ofsurface water(speed of1.5 m/s, 3 km).Riversampling:Trawl attachedon a ridge for15–30 mins

Top 18 cm Opening:60 � 18 cmMesh size:300 μm

(Faureet al.2015)

LaurentianGreat Lakes(Canada)

Dynamicmanta trawl

Trawl distancecalculated withonboard speedmeter; 60 minsurveys

Top 16 cm Opening:61 � 16 cmMesh size:333 μm

(Eriksenet al.2013)

Lake Hovsgol(Mongolia)

Manta trawl 60 min longwith a targettow speed of3.5 knots

n.a. Opening:16 � 61 cmMesh size:333 lm m

(Freeet al.2014)

Rhine river(Switzerland–Netherlands)

Manta trawl Tows of15 minyieldingfiltered watervolumes of60–250 m3

(mean 150 m3)

n.a. Opening:60 � 18 cmMesh size:300 μm

(Maniet al.2015)

Seine river(France)

Plankton net(and ii)

Upstream15 minsampling

Top10–35 cm

Plankton netmesh size:80 μ

(Gasperiet al.2015)

(continued)

Introduction to the Analytical Methodologies for the Analysis of. . . 9

(f) Detailed information on relevant hydrological (including tributaries) and mete-orological conditions

When feasible, additional metadata should be included, as, as previously noted,multiple factors influence the distribution of microplastics in both riverine andestuarine environments. Such metadata comprise wind direction (before and duringsampling), historical precipitation data, as well as discharge data, depth and flowprofiles, possible point sources (such as wastewater treatment plants, industrial oragricultural activities, and highly densely populated areas, among others), anduncertainty of quantitative results.

Soil and Sediment Sampling

As a man-made product, a large proportion of plastic contamination sources areinland, and a large fraction of the generated plastic waste is sent to landfills. Forexample, in Europe (EU28 + Norway + Switzerland), 24.9% of the 29.1 milliontonnes collected were landfilled (PlasticsEurope 2019), and, in the USA, thispercentage surpasses the 50% mark (Narancic and O’Connor 2017). The vastmajority of studies focusing on the prevalence and effects of microplastics havebeen carried out in aquatic and, more specifically, in marine settings. Nonetheless,there has been an increased awareness by the scientific community of this limitation,and multiple efforts have been devised towards an assessment of these issues interrestrial environments, including both soils and sediments. This limited amount ofresearch is likely due to the inherent technical difficulties in isolating and

Table 1 (continued)

LocationDevice/toolcharacteristics Method

Monitoringdepth Observations Ref.

sampling witha manta trawl

surveys(volumes 182–200 m3)

Manta trawl,opening: n.a., meshsize: 300 μm

Stockholmarchipelago(Sweden)

Trawl Total of21 surfacesamples

n.a. Opening:61 � 16 cmMesh size:335 μm

(Gewertet al.2017)

Dongting andHong Lakes(China)

Pump Total of 20 L(2 � 10L)collected

Top 20 cm Steel 50 μmmesh sieve

(Wanget al.2018)

10 J. P. da Costa and A. C. Duarte

characterizing microplastics from such complex solid or organo-mineral matrices, assoil, when compared to the relative ease with which these particles may be extracted– and analyzed – from aquatic samples. There is no question that soils are contam-inated with (micro)plastics, as not only are some types of soils deliberatelymanufactured containing plastic materials, such as technosols, soils “whose proper-ties and function are dominated by technical human activity as evidenced by either asubstantial presence of artifacts, an impermeable constructed geomembrane, ortechnic hard rock” (Rossiter 2007) but plastic has also been historically used toincrease yields, fruit quality, and water efficiency, as is the case of plastic mulching(Steinmetz et al. 2016). Having been demonstrated that microplastics may exertdeleterious effects on numerous aquatic organisms (reviewed, e.g., in Guzzetti et al.2018), it stands to reason that these materials may equally affect terrestrial biota,particularly considering that smaller organisms, namely, rotifers and ciliates,thrive on thin films of water that cover the soil surface, basically living in water.Mesofauna, namely, collembola (arthropods) or enchytraeidae (annelids), can cer-tainly ingest microplastics, and specific histopathological and molecular effects onthese organisms have been reported (Rodriguez-Seijo et al. 2017). Microplasticsmay reach soils through a variety of routes, such as through wastewater treatment(WWTP) sludge, as sewage sludge, considered a biological residue, is frequentlyused in agriculture as an alternative disposal technique of waste. In fact, by com-bining sewage sludge fate, available national data on farm areas and microplasticcontent guesstimates, Nizzetto and colleagues estimated between 125 and 850 tonsof microplastics/million inhabitants to be annually added to agricultural soils inEurope alone (Nizzetto et al. 2016). This equates to a yearly total ranging from63,000 and 430,000 tons of microplastics deposited in European farmlands. In theUSA, this estimation varies between 44,000 and 300,000 tonnes (Ng et al. 2018).Urbanized areas, including roads and transport infrastructures, as well as storm andrainwater, also contribute to the accumulation of these materials in WWTPs andresulting sludge. Organic fertilizers (compost) obtained from the fermentation andcomposting of biowaste, industrial effluents, landfill leachates, and general litteringand dumping of plastic waste are also important pathways of entry of microplasticsinto soils. A recent additional source is comprised of the famed biodegradable andoxo-biodegradable plastics. Not only are these terms often mistakenly used inter-changeably (Box 1), it has been demonstrated that these products generate micro-sized particles similar to traditional plastics (Shruti and Kutralam-Muniasamy 2019).Owing to the “green” label, such materials could end up being discarded byconsumers with little concern regarding their pervasiveness, thus resulting in theirincreased accumulation in the environment.

Introduction to the Analytical Methodologies for the Analysis of. . . 11

Box 1 What Are Bioplastics?Bio-based does not equal biodegradable. In spite of their interchangeable use,some bioplastics are bio-based, some are not. Moreover, some fossil-basedplastics are biodegradable, while others – the most frequently used types – arenot.

The distinction of bio-based and fossil-based plastics, according to their biodegradability.(Adapted from (EuropeanBioplastics 2018). PP Polypropylene; PE Polyethylene; PA Poly-amide; PHA Polyhydroxyalkanoates; PBS Polybutylene succinate; PET Polyethylene tere-phthalate; PBAT Polybutylene adipate terephthalate; PCL Polycaprolactone)

For soils sampling, as is often the case for water sampling, different researchers/groups/institutions tend to use different strategies. Samples may be collected usingsteel tools (Scheurer and Bigalke 2018), Van Veen grabs (Claessens et al. 2011),coring (Ballent et al. 2016), or a combination of these, depending on the researchquestion.

Figure 2 depicts some of these most commonly used tools. Although suchdifferent sampling strategies may yield comparable results, there is still no exhaus-tive study demonstrating the equivalence – or lack thereof – of these methodologies.It is nonetheless quite likely that different sampling methodologies will yielddifferent results, as sampling tools reach different depths and these may accountfor distinct zones of accumulation of microplastics. Deeper sampling methodologiesmay result in an under- or overestimation of the number of particles per dry weight,owing to their vertical transport in soils, arising from human and/or animal activity.

12 J. P. da Costa and A. C. Duarte

For example, Liu and colleagues sampled 50 � 50 cm (2500 cm2) areas at twodistinct depths: 0–3 cm and 3–6 cm. These authors reported statistically differentconcentrations of microplastics at these depths, with 78.0 and 62.5 items.kg�1 of dryweight at the upper and lower depths, respectively (Liu et al. 2018) (Table 2).

Biota Sampling

Plastics, in general, and microplastics, in particular, are ingested by a wide range ofaquatic organisms, from the largest, as whales (de Stephanis et al. 2013), to those atthe basis of the food chain, such as zooplankton (Desforges et al. 2015). It is,therefore, necessary to accurately ascertain the prevalence of these materials inbiota, so that not only subsequent analyses and assessments of their potential

Fig. 2 Most frequently used tools for sampling soils and sediments. (a) A rotating drum sampler;(b) core soil sample(r); (c) van Veen grab; and (d) stainless steel scoop/spoon. (Adapted from Pintoda Costa et al. 2019)

Introduction to the Analytical Methodologies for the Analysis of. . . 13

ecotoxicity are carried out but meaningful strategies for the minimization of suchimpacts may be devised. Given that many studies are also conducted from a humanhealth perspective, i.e., attempting to determine the exposure of humans to contam-inated seafood, for example, it should be first evaluated whether wild or farmedanimals are to be examined. In an attempt to determine if there are differences infarmed and wild clams, Davidson and Dudas found that these were not significant(Davidson and Dudas 2016). However, it should be noted that, as previouslyunderscored, such conclusions should be not extrapolated and considered as thenorm, as nor only is there a very limited number of studies focusing on suchdifferences, but different areas and farming techniques could result in different levelsof exposure to the cultured organisms. Sampling of biota must also, of course, abideby the associated ethical issues. Although these may be of lower importance fororganisms such as zooplankton, in natura studies face the ethical constraints ofsubjecting large organisms, such as marine mammals, to laboratory analyses inaddition to the inherent difficulties in distinguishing, directly and indirectly, ingestedmicroplastics (Nelms et al. 2018). This limitation may be overcome, for example, bystudying scat samples, whenever possible, which may also help in identifyingpotential areas of more significant exposure in animals that, throughout their lifecycle, cover extensive areas. Nonetheless, the number of analyzed samples remainssomewhat limited, and, consequently, accurate measurements of microplastics inorganisms may be hindered (Mai et al. 2018). For example, when studying the diet of71 individuals belonging to 4 species that were opportunistically collected asdeceased bycatch, Wedemeyer-Strombel and colleagues found no anthropogenic

Table 2 Examples of different sampling techniques for microplastics (MP) in soils and sediments.The major findings are listed. Note that the unit reporting is not the same for all the studies included

Country LocationType ofsoil

Samplingmethodology Findings Ref.

China Suburbs Farmlandsoil

Steel tools Up to 78 MPkg�1

(Liu et al.2018)

Countryside Not reported Average 18760MP kg�1

(Zhang andLiu 2018)

Australia City Municipalsoil

Not reported 500–6900 mg kg�1

(Fuller andGautam2016)

UK Riverbasins

Sediments Steel tools Up to 660 MPkg�1

(Horton et al.2017)

Switzerland River shore Floodplainsoils

Steel tools Up to 593 MPkg�1

(Scheurerand Bigalke2018)

Mexico Suburbs Homegardensoils

Not reported Up to 2770 MPkg�1

(HuertaLwanga et al.2017)

India Lakesediments

Wetland van Veengrab

Mean252.80 � 25.76particles m�2

(Sruthy andRamasamy2017)

14 J. P. da Costa and A. C. Duarte

debris in leatherback sea turtles (Wedemeyer-Strombel et al. 2015). However,the sampling pool was comprised of 45 olive ridleys (Lepidochelys olivacea),22 greens (Chelonia mydas), 2 loggerheads (Caretta caretta), and 2 leatherbacks(Dermochelys coriacea), meaning that leatherbacks were comparatively under-sampled. It certainly does not demonstrate that leatherbacks do not ingest anthropo-genic litter, as has been subsequently demonstrated, for example, by Duncan andco-workers, whom found that microplastic contamination was ubiquitous in allsampled marine turtles (Duncan et al. 2019).

Laboratory-based studies on the ingestion and effects of microplastics by biotafrequently resort to manufactured plastic beads or particles that are obtained from themechanical shredding of the said commercially available plastic beads of knownorigin (e.g., Rodriguez-Seijo et al. 2017). These methods have the advantage ofbeing easily recognized following ingestion and to be relatively easy to compare tothe virgin materials, therefore allowing an immediate assessment of any modifica-tions that these materials may undergo following ingestion. For smaller organisms,like plankton, fluorescent particles may also be used, as this facilitates the recoveryand counting of the ingested particles (e.g., Cole et al. 2013). It should benoted, however, that such studies have been often been questioned regarding theirenvironmental validity, as the concentrations of microplastics used frequently farexceed those reported in the scientific literature, and, therefore, any observedecotoxicological effects may be greatly exacerbated by such high concentrations,as noted by some researchers (da Costa et al. 2016).

Generally, biota samples are dissected to collect microplastics from the varioustissues and organs. Following dissection, when analyses are not carried out imme-diately, these different samples – tissues and organs – should be conserved or frozen,particularly the gut content or the entire digestive system. This may be done by usingplastic-friendly fixatives, such as formalin, or fully dried and kept in the dark.Furthermore, data reporting should include an assessment of microplastics in termsof (a) incidence, i.e., percentage of samples containing microplastics; (b) individualabundance, meaning the average number of particles per analyzed individual; and(c) abundance by mass, i.e., weight in grams, ideally, wet and dry weight. As perrecommendation of an EU Technical Report, this figure should be accurate to thefourth decimal (EU-JRC 2013).

Atmospheric Sampling

Among the different environmental compartments, air is undoubtedly the leaststudied regarding the prevalence of microplastics. This is not surprising, consideringthat atmospheric contamination is likely comprised of rather small microparticles,which subsequently increase the inherent technical limitations associated withtheir collection, isolation, and analyses. In the limited scientific literatureavailable, samples are usually collected resorting to specially devised air pumps.For example, Dris and colleagues collected indoor air at a rate of 8 L�min�1, and,after, the air was then filtered through a 1.6 mm filter (Dris et al. 2017). The authors

Introduction to the Analytical Methodologies for the Analysis of. . . 15

found that microplastics comprised a third of all indoor fibers and that the concen-trations were highly site-dependent. In another study, smaller filters were used, but,again, a high variability, ranging from 0 to over 32,000 particles filtered during30-min air sampling periods, was observed. The authors noted that the number ofmicroplastics in the atmosphere varied substantially depending on time, space, andseasonal conditions (Tunahan Kaya et al. 2018). Modified rain samplers have alsobeen used to collect atmospheric fallout and to determine the concentration ofmicroplastics in the atmosphere. This was done, for example, by Allen and col-leagues, whom, using this method, reported daily counts of 249 fragments, 73 films,and 44 fibers per square meter in a remote location at the French Pyrenees (Allenet al. 2019). A different approach was used by Vianello et al. (Vianello et al. 2019).These authors used a breathing thermal manikin, which has long been used to assessindoor environment, validate the computational fluid dynamic predictions of airflow,and evaluate occupants’ thermal comfort and perceived air quality. The researchersfound that all samples were contaminated with microplastics and that concentrationsvaried between 1.7 and 16.2 particles.m�3. Interestingly, considering the indoorsampling settings, the authors noted that synthetic fragments and fibers accountedapproximately 4% of the total identified particles but that these were, generally,smaller than non-synthetic particles, leading to the potential of inhalation of thesematerials by humans (Vianello et al. 2019). More recently, a method has beendescribed for the sampling of particles from air samples (Prata et al. 2020). Thoughgreatly improving analysis time, the method relies on the use of an adapted air filterfor the assessment of particulate matter (PM10, defined as inhalable particles, withdiameters that are generally 10 micrometers and smaller), thus underscoring the needto develop specific methods for the collection and analysis of microplastics in air.

Sample Handling

As with any experimental protocol, following sample collection, sample handlingmust be judiciously evaluated. Nonetheless, this aspect is of particular relevancewhen determining the presence of microplastics in the environment, as most sam-pling methods are nonselective. As such, subsequent steps for the quantification andcharacterization of microplastics include isolation/purification steps, often insucceeding fashion (Löder and Gerdts 2015; Pinto da Costa et al. 2019). Therefore,special care must be taken to avoid any potential contamination, which can arisefrom numerous sources, including field gear, clothing, sieves, and nets, as well asstoring containers, plastic ware, and air conditioning systems during laboratoryhandling and analysis. As noted by Costa and Duarte, any sampling process involvesnot only the capture of the plastic materials by the sampling tool but also isolationfrom the matrix, whether soil, water, or tissue, before proceeding to the followingsteps of identification, characterization, and quantification (Costa and Duarte 2017).Based on the scientific literature, a set of recommendations for processingmicroplastic-containing samples in the laboratory is highlighted in Box 2. However,the sampling matrix must not be ignored, as it is key in understanding and

16 J. P. da Costa and A. C. Duarte

interpreting the obtained results and to contextualize the presence – if any – ofmicroplastics within the matrix. Ideally, a biogeochemical characterization of thesampling matrices should be conducted, and, because the matrix removal procedureis usually destructive, samples should be taken in excess (Costa and Duarte 2017),not only for these analyses but also for the eventual contamination of any of thesample replicates. It is also important to consider sample storage for future reference,including the use of more sensitive methods for the analysis of microplastics thatmay meanwhile develop. As long-term contamination is a common route of con-tamination and transformation of environmental samples, a dedicated storage loca-tion is recommended, though not always possible.

Box 2 Contamination Prevention and Assessment Protocol. The Guidelines areBased from Numerous Publications and Adapted from Crawford and Quinn(2017)1. Preparation

A clean cotton (natural fiber) lab coat should be worn at all times. Gloves(nitrile) should only be used when manipulating chemicals (such as H2O2)and if the user exhibits any cuts or bruises. Otherwise, the use of gloves isnot recommended, and thorough hand washing should be done beforesample handling. Ideally, the user should wear clothing composed ofnatural fibers only.

Air currents should be minimized. All doors and windows should beclosed during sample handling. The laboratory should also be cleaned andfree from dust. Working space should be away from any overhead fixturesthat may lead to the accumulation of dust. All equipment should bethoroughly cleaned with 70% ethanol. Material used should also be cleanedwith ethanol and then rinsed three times with distilled water. Ideally, thiswater should be previously filtered and stored in glass containers, asopposed to the typically used plastic containers and squirters. After, coverall equipment and unused material with aluminum foil. Immediately beforework, working surfaces should be cleaned with ethanol.

2. Solid particulate surface monitoringThe process should be carried out before and after all analyses.

The adhesive side of a section (e.g., 5 cm2) of transparent high tackadhesive tape should be pressed three times onto the work surface and thencollected. Any solid particulate will adhere to the tape. These sections canthen be adhered to a clean piece of cellulose acetate film, in which date andtime are noted. The collected sections of adhesive tape are then examined,and the putative presence of microplastics, including fibers, is assessed.If present, these materials should be identified by, for example, infraredspectroscopy and subsequently excluded from the sample of interest.

(continued)

Introduction to the Analytical Methodologies for the Analysis of. . . 17

Box 2 Contamination Prevention and Assessment Protocol. The Guidelines areBased from Numerous Publications and Adapted from Crawford and Quinn(2017) (continued)3. Solid airborne particulate monitoring

Prior to initiating the work, clean pieces of dampened filter paper should beplaced in standard ∅90 cm Petri dishes, and the filter paper should coverthe entire internal area of the dish. These are then placed around the worksurface, where they remain for the duration of the laboratory work in theimmediate vicinity of the user. Upon completion of the lab work, the filtersare then examined for the presence of microplastics. Alternatively, the Petridishes may be covered with the respective glass lid and later analyzed.Again, found particulates should be identified and excluded from thestudied samples.

Extraction/Separation of Microplastics

Following collection, samples should be processed, and existing microplasticsshould be separated and clean for subsequent quantification, identification, andcharacterization. The most widely used techniques for the separation of micro-plastics include density flotation, sieving, filtration, and digestion protocols. Thechoice of method depends upon the characteristics of the sampled matrix, andpotential hazards and costs vary accordingly.

The most commonly used method is density separation or flotation. This tech-nique has been used in numerous types of media (Nguyen et al. 2019). Based on thedifferences of density between plastics and non-synthetic materials, it is frequentlyused as an eco-friendly and inexpensive technique, frequently relying on the use of asaturated NaCl solution. However, NaCl concentrated solution has a density of1.2 g�cm�3, which is insufficient to allow for the flotation of all polymers. Thus,alternative salts have been used with added benefits, although some trade-offsbetween costs and/or health hazards have to be reached. It has been suggested thatthe apparent inherent higher costs and/or hazards of separating plastics of higherdensity, such as PVC (density of rigid PVC is 1.45 g�cm�3), may be reduced byimplementing a two-step method, in which a NaCl solution is used in a “pre-extraction” step and a NaI solution (1.80 g�cm�3) is then used for a second flotationstep (Nuelle et al. 2014). Recently, Liu and colleagues summarized the features ofcommonly used solutions for flotation methods, and these are reproduced in Table 3.

A potential alternative to NaCl has been the use of CaCl2. Although the efficiencyof the latter may be higher than that of NaCl, given the higher density of CaCl2, someresearchers have pointed out that the divalent Ca ions may result in the formationof aggregates with the present organic material, thus impacting the subsequent

18 J. P. da Costa and A. C. Duarte

identification process (Wang et al. 2020a). Solutions of higher density may beobtained using ZnCl2 or NaI (Table 3). Nonetheless, not only are these solutionsconsiderably more expensive, they pose a significant risk to the environment andhave been shown to have embryotoxic effects (Koda-Kimble 2007). Alternatesolutions, such as NaBr and ZnBr2, have also been used for the separation ofmicroplastics (Quinn et al. 2017), and, in the case of the former, given that it is ofrelatively low cost and environmentally friendly and shows relatively high efficiencyin the separation of microplastics, its use may be more widespread in the future,particularly, in soils. Some of these separation solutions have also been implementedwithin specially developed instruments aimed at facilitating the density separationprocess. These include, for example, circulation separation devices (Liu et al. 2019),air-induced overflow method (Nuelle et al. 2014), volume reduction by elutriation(Claessens et al. 2013), or a “so-called” Munich Plastic Sediment Separator (Imhofet al. 2012). These are interesting approaches that may in the future curtail thedifficulties associated with the high-throughput analysis of samples, though, pres-ently, they remain very much independently used by the researchers responsible fortheir development. Additionally, the reported extraction efficiencies vary greatlyamong the used techniques, but such disparities depend not only on the developedmethod but also on the characteristics of the sampled particles, including shape, size,and type of polymer.

Table 3 Commonly used salts in density separation methods of microplastics. (Adapted from Liuet al. 2020)

Salt

Saturatedsolution density(g�cm�3)

Approximated costsper 100 mLof solutiona Potential hazardsb Type of sample

NaCl 1.19 $0.21 – Water;sediment; soil;biota; air

NaI 1.89 $26.49 H315-H319-H400 Water; air;sediment; soil;biota

NaBr 1.55 $1.63 H303-H313-H320 Sediment; soil

CaCl2 1.42 $0.59 H319 Sediment; soil

ZnCl2 1.68 $6.21 H302-H314-H410 Water;sediment; soil;biota

KBr 1.58 $13.3 – Water;sediment; biota

Na2WO4 1.4 $4.51 H303 Water;sediment; biota

Oil – – No Sediment; soilaCost estimation based on the commercial prices. Price is US dollarsbThe hazard code used follows the guidelines established in the Globally Harmonized System ofClassification and Labeling of Chemicals (OSHA 2013)“H” stands for “hazard” and “3” and “4” for health and environmental hazards, respectively

Introduction to the Analytical Methodologies for the Analysis of. . . 19

Following density separation, samples are commonly subjected to filtration.Generally, to avoid any cross-contamination, stainless steel sieves of glass fiberfilters are used. Naturally, rinsing with distilled water is always necessary before,between, and after each sieving/filtration. Depending on the type of samples,including matrix complexity, an array of sieves is often implemented (Mai et al.2018), and a 1 mm mesh size has been recommended and is often used (Nuelle et al.2014). Subsequently, a 500 μm mesh size is frequently used, so that fractions largerand smaller than this size are obtained. The main reason for this is that 500 μm isbroadly considered as the threshold for visual identification. Researchers haveresorted to different degrees of size fractioning, and, although this may result in adetailed description of microplastics present in a sample and an exhaustive sizecategory classification, with the procedural advantage of reduced rate of clogging,it also represents a lower frequency of microplastics per size range. Hence, astandardized series of size fractioning is needed so that, again, data comparison isfeasible. Electrostatic metal separators have also been used, which sort material intononconductor, mix, and conductor fraction (Felsing et al. 2018). Microplastics maybe recovered from the mix and nonconductor fractions, which are combined. Thoughan interesting approach towards the high-throughput analysis of microplastics inenvironmental samples, the method requires extensive validation, as the reportednumber of replicates was limited and recovery values were not subject to statisticalvalidation. Alternatively, oil has also been used as the density separation media(Crichton et al. 2017). This method yielded very good recoveries (96.1% � 7.4), butthe inclusion of post-extraction cleanup steps to remove traces of oil for polymeridentification (in this case, FTIR) has rendered this method of limited use.

The choice of filter is also of paramount importance. Microplastic studies areoften conducted using glass fiber membranes, cellulose acetate/nitrate membranes,polycarbonate membrane, or alumina membrane. Some researchers have reportedtheir findings using nylon filters (e.g., Yang et al. 2019), but their use is discouraged,as it may lead to the release of fibers that result in the overestimation of fibers presentin the analyzed samples. A similar issue has been observed with glass fiber filtermembranes, often used (e.g., Prata et al. 2019), although these may be distinguishedfrom microplastics, including fibers, with relative ease. Their surface, however, israther rough and may hinder identification when resorting to infrared spectroscopy(Liu et al. 2020). On the other hand, materials such as polycarbonate exhibitrelatively high absorbance in the near and infrared regions, which may disturb thesignal in Fourier transform infrared spectroscopy (FTIR) studies. Cellulose filters,contrarily, show a smooth surface, as alumina’s. Both types of filter membranes aresuitable for conventional filtration experiments, and the latter is transparent to IRabove the 1250 cm�1 wavenumber. Pore size is dependent on the nature of samplesand the research questions. Broadly, aquatic samples are filtered using membraneswith pore sizes varying between 1 μm and 0.5 mm, while biological samples tendto be filtered using membranes of 0.22 μm to 5 μm pores (Mai et al. 2018).Nonetheless, irrespective of the type of membrane and pore size, the accurateidentification and quantification of microplastics on the membrane remain a chal-lenge, though research on the field continues to evolve with the development of

20 J. P. da Costa and A. C. Duarte

algorithms, for example (Prata et al. 2019), that may contribute, in the long run, tothe development of methods amenable to semiautomatic analyses.

Sample purification is also frequently necessary. As environmental samplescontain organic matter to the different extent that is not during the density separationand filtration steps, an additional purification step, commonly referred to as diges-tion, is needed. Acidic, alkaline or enzymatic digestion procedures are commonlyused, as well as oxidizing treatments. Table 4 summarizes the advantages anddisadvantages associated with these treatments.

The choice of the digestion or purification method is of the utmost importance forthe accurate identification of the isolated microplastics. Inadequate removal ofassociated organic matter, for example, will render subsequent analyses extremelydifficult, as they will affect identification results, such as infrared or Raman spec-troscopies. Although the degree to which these methods may impact the chemicaland physical characteristic of microplastics, it remains undetermined how the poten-tially adsorbed chemicals, such as POPs and plasticizers, are affected by thesedifferent approaches. Nonetheless, Fenton reaction-based purification methodolo-gies will undoubtedly fully degrade any present organics, as this is a frequentstrategy used for such purposes (Soltani and Lee 2019).

Identification of Microplastics

The most-reported technique used in the detection of microplastics consists in thevisual identification of likely plastic particles. This is followed by confirmationmethodologies through chemical composition analyses. The combination of opticaland spectroscopic methods allows for the minimization of the occurrence of falsenegative and/or positives (Silva et al. 2018). Nonetheless, this strategy is hindered bythe frequent misidentification of a considerable percentage of particles, up to 20%(Eriksen et al. 2013). As noted by Lenz and colleagues (Lenz et al. 2015), nearly athird (32%) of visually counted particles <100 mm were not confirmed whensubsequently analyzed with micro-Raman (μRaman). When using FTIR

Table 4 Commonly used digestion protocols and related advantages and disadvantages. (Adaptedfrom Kershaw et al. 2019)

Method Reagents Advantages Disadvantages

Aciddigestion

HNO3, HCl, HNO3, HClO4 Fast May degrade somepolymers

Alkalinedigestion

NaOH, KOH Minimal damage tomost polymers

Cellulose acetate isdegraded

Oxidativetreatment

H2O2,Fe salts (Fenton reaction)

Inexpensive Requires temperaturecontrol; multiple cyclesare often needed

Enzymaticdigestion

Tested enzymes includeproteinase K, lipase, cellulose,trypsin, and protease A-01

Highly effective;minimal damage tomost polymers

Expensive; time-consuming

Introduction to the Analytical Methodologies for the Analysis of. . . 21

spectroscopic analyses, this value reached, in some cases, 70% of erroneous iden-tification, as reviewed by the team led by Thiel (Hidalgo-Ruz et al. 2012). Opticalidentification may also be achieved by resorting to optical microscopy. Surfacetexture, color, and morphological characterization of the suspected particles maybe gathered, effectively permitting the accurate identification of microplastics.Whether visual or microscope-assisted, visual identification is a fast, simple, andcheap technique that has the added advantage of being amenable to in situ identifi-cation. However, reporting on the gathered data must not be restricted to visualidentification methods, as this has been demonstrated to be a method highly prone tomismatching errors (da Costa et al. 2017). For example, Silva and colleagues (Silvaet al. 2018) describe a study in which the authors identified blue fibers as micro-plastics that were later identified as cotton (indigo) (Dyachenko et al. 2017) viamicro-FTIR (μFTIR). The use of complementary techniques, as spectroscopic-basedmethods, is not always available, and alternatives, as the use of a hot needle, havebeen suggested to confirm the identity of suspected microplastics (Campbell et al.2017). This is, however, considerably difficult for smaller particles. Naturally,optical microscopy is limited to the maximum resolution of these devices, consid-ered to be ~1 μm (with a 10� magnification and a numerical aperture of 0.3).However, it is frequent to enumerate particles that are up to 10 mm, and, althoughsmaller particles may be accounted for, these are often not included unless brightlycolored and hence reliably identified (Covernton et al. 2019). Considering theaforementioned details regarding visual identification, it emerges that this approachshould only be used as an initial assessment when quantifying microplastics presentin a given sample. Chemical composition must be carried out, so that microplasticsare not only counted but also identified regarding their polymeric nature, as this willactively contribute to an analysis of the potential sources of these materials in thestudied samples. Electron microscopy, and, more specifically, scanning electronmicroscopy (SEM) is also a reoccurring analysis technique used for the identificationof microplastics. It provides highly resolved images with extreme clear and definedmorphological characterization of the imaged particles, including features that maybe indicative of weathering and/or (bio)degradation (da Costa et al. 2018). Coupledto energy dispersive X-ray spectroscopy (EDS, SEM-EDS), it may be used forevaluating the presence of plasticizers and/or adsorbed contaminants (Shikuku2019). Contrary to the perceived usefulness of SEM-EDS; however, this techniqueshould not be used to differentiate between polymers, as the backbone of manypolymers is identical and the vacuum under which the samples are analyzed is notcomplete, meaning that atmospheric components (e.g., H, O, N, and C) may accountfor the elements (semiquantitatively) identified. SEM or SEM-EDS is, however, anexpensive and lengthy technique and involves laborious sample preparation. Addi-tionally, it should only be used for obtaining representative details of sampledmicroplastics, as its use in routine and, above all, high-throughput analysis, isunfeasible. Another limitation of this technique is that colors are not identifiableunder SEM, and this is a recommended descriptor and class of categorization ofidentified microplastics.

22 J. P. da Costa and A. C. Duarte

The de facto identification of polymers is based on their chemical composition,which may be achieved through spectroscopic methodologies, as Raman and FTIRspectroscopies. Optimized variations of FTIR, as attenuated total reflectance FTIT(FTIR-ATR), μFTIR, and focal plane array detector (FPA) FTIR are often usedin microplastic identification (da Costa 2018). In the case of FTIR and their varia-tions, however, the choice of technique is perhaps more dependent on the type ofplastic than on the sample. As underscored by Mai and colleagues (Mai et al. 2018),FTIR-ATR has been shown to yield better results than μFTIR when analyzingmicroplastics of irregular shape, though, contrary to μFTIR, unsuitable for particlessmaller than 500 μm. This limitation may be overcome resorting to FPA-FTIR,which may successfully identify particles as small as 20 μm within a relatively largesurface area (>10 mm in diameter), allowing for the complete analysis of ∅47 mmfilters in under 9 h (Löder and Gerdts 2015). FTIR analyses are also suited toinvestigate the degree of weathering, as this process has a demonstrated effect inthe collected spectra of polymers; higher degrees of (bio)degradation result in theformation of peaks at known wavenumbers (carbonyl and hydroxyl groups) ofhigher intensity that, when divided by that of peaks that do not change (e.g., thepeak associated to the symmetric stretching vibration of the methylene group),allows the determination of a “rate” or “index” of degradation (Silva et al. 2018).This is, however, mostly used to PE and PP, and further studies are requiredto validate such an index of degradation for all available synthetic polymers.Nonetheless, whenever possible, this is certainly an assessment that should beincluded in any set of results.

Raman spectroscopy, much like IR spectroscopy, offers the possibility of accu-rately identifying microplastics based on their unique IR spectra. FTIR and Ramanspectroscopies are based in complementary principles: molecular vibrations that areactive in IR and inactive in Raman and vice versa (Koenig 2001). Also a nonde-structive method, Raman may be coupled to optical microscopy techniques(μRaman), which can theoretically achieve a higher resolution than μFTIR, below1 μm (Löder and Gerdts 2015; Silva et al. 2018). However, despite this theoreticalhigher resolution, real-world applicability is yet to be demonstrated. Raman spec-troscopy, when complemented with confocal laser-scanning microscopy, is espe-cially suited for the analysis of biological samples, allowing for the localization ofpolymers with subcellular precision (Li et al. 2020). Another variation is couplingRaman spectroscopy with hyperspectral imaging, which has been used to identifythe tissue localization of polymeric particles in zebra fish and shrimps (Gallowayet al. 2017). There, nonetheless, limitations to the use of Raman spectroscopy, asfluorescent samples are excited by the laser and remaining biological residues, forexample, can severely impact the interpretation of generated Raman spectra. Thismay be overcome or partially minimized using higher wavelengths (>1000 nm), butthe lower laser energy yields lower signals, and a compromise must be reached(Löder and Gerdts 2015). Hence, thorough purification steps prior to Raman ana-lyses should be included in the experimental protocols. This highlights the need toexpand the research on the optimal conditions under which studies of differentpolymers, in specific matrices, should be performed.

Introduction to the Analytical Methodologies for the Analysis of. . . 23

Less used, destructive techniques such as pyrolysis or thermodesorption gaschromatography with mass spectrometric detection (Pyr-GC/MS and TDS-GC/MS, respectively) are two nonvisual methods that have been applied to the identi-fication and quantification of polymers in different types of samples. These tech-niques are based on thermoanalytical principles that have the inherent limitation ofnot offering the possibility of determining the number of particles or their morphol-ogy present in the analyzed sample. Though it may be inferred that such techniquesare not limited to any specific size limit, it has been suggested that, for obtainingclear chromatograms/mass spectra, particles, the quantity, or size of plastic particlesnecessary should be >100 μm (Zhang et al. 2020). Recent advances, particularly, inTDS-GC/MS methods, may enable the thorough identification and quantificationof small microplastics and plastic quantities, opening the possibility for the analysesof even atmospheric particles, frequently present as smaller particles. This techniqueis also more suited for processing larger sample masses, even of more complexmatrices, thus opening the possibility of semiautomation. Furthermore, thesemethods have the added benefit of allowing for both the identification andquantification of both additives and adsorbed organic contaminants, thus providingkey information for an extensive ecotoxicological assessment of present particles.However, such strategies should always be used in conjunction with visual-basedtechniques, as enumeration, color, and morphologies are essential aspects whenevaluating the presence of microplastics in environmental samples.

Recommendations, Knowledge Gaps, and Future Venuesof Research

There is an extensive and growing body of research related to the presence, fate, andeffects of microplastics in the environment. Nonetheless, from the previous sections,it emerges that much of the gathered data may be of limited use and clear and definedmethod for this evaluation are urgently needed, including sampling and datareporting harmonization, as well as quality control assessment and even inclusionof cross-contamination methods. Far from exhaustive, a summary of research gapsand needs may be found below, and these may be considered as indicative andsuggestions of the immediate routes for research in microplastic-associated studies.

Choosing the appropriate sampling location, sample strategy, and the number ofneeded replicates, namely, as a function of the subsequent analyses to be performed,is essential. However, as noted, this is also dependent on the research question.Broadly, for soil and sediment sampling, choice of sampling locations is ofspecial relevance, as it may affect the accuracy of representativeness. Informationsuch as sampling depth must be included and, ideally, standardized. For marinewaters, manta trawls or nets are highly recommended, given the large areas thesetools allow to sample. Depending on the mesh size used, smaller microplasticsmay be sampled resorting to the grab sampling technique. These strategiesshould always be accompanied by key information, such as wind and currents,and the use of at least one, and ideally, two, flowmeter(s), is recommended.

24 J. P. da Costa and A. C. Duarte

Techniques for sampling microplastics in biota are less amenable to standardization,as the biological materials are as varied and biota itself, particularly regardingreplicates, are also highly dependent on the availability of the specimens. For airsampling, there is an urgent need of developing fit-for-purpose methods, and futuretrends may very well lie within the classification of microplastics as particulatematter and their inclusion in the PM2.5 and PM10 categories, though this is, to date,conjectural. However, considering the available scientific literature, there is also theneed to extend sampling protocols, irrespective of the sampled matrix, for anadequate temporal and spatial assessment on the prevalence of microplastics in theenvironment.

Sample preparation should be carried out using density separation, particularly incomplex samples, such as soils and sediments. This step should include a sizefractioning step, and resorting to the size cut-off defined by Hidalgo-Ruz andcolleagues (> and <500 μm) is suggested (Hidalgo-Ruz et al. 2012), so that resultsof different studies may be directly compared. Nonetheless, given the vast array ofsolutions and reagents used for this purpose, there is a clear need to identify suitedand cost-effective methods that may include the evaluation of alternative separationagents, such as surfactants. Hence, more research is needed in this context, and suchstudies should be promoted, and the assignment of standard operating procedures(SOP) that vary according to the sampling matrix should be implemented. Whenneeded, removal of organic matter should be conducted resorting to Fenton reaction,though it is concomitantly necessary to include replicates that undergo an alternative,milder treatment and that do not degrade any potentially present adsorbed contam-inants and plasticizers, whenever these are the focus of research. Enzymatic diges-tion is the better option, but its inherent high costs do not allow for its universal use,and, therefore, including this strategy in any SOP is not encouraged.

Although visual identification remains, in many cases, as the benchmark methodfor the identification of microplastics, the inclusion of spectroscopy-based tech-niques is essential, particularly for particles <500 μm. Ideally, for an accurateassessment of all classes of sizes of microplastics present, μFTIR and/or μRamantechniques should be used. The advances made in recent years regarding the use ofmodified versions of these methods, such as FPA detectors, have yielded promisingresults, particularly when considering their relatively short time of analysis, butSOPs are not yet available. The correct identification and quantification of particles<1 μm remains a challenge, and, although methodologies such as X-ray diffraction(Mai et al. 2018) have been suggested as a potential alternative for circumventingthis limitation, this is yet to be validated within the context of environmentalanalysis. Ultimately, addressing these methodological issues may result in thedevelopment and implementation of techniques that would permit an automated,or, at least, semiautomated, methods for the identification and quantification ofmicroplastics in the different compartments of the environment, effectively reducingexperimental errors. Only then will it be possible to:

(a) Improve the understanding of the pathways of microplastic distribution in bothland and aquatic systems.

Introduction to the Analytical Methodologies for the Analysis of. . . 25

(b) Determine the apportionment of microplastics, between sources and sinks.(c) Understand the dynamics and fate of microplastics in the environment as a

function of their chemical and physical characteristics.(d) Develop mathematical models that may be used to predict distribution and

accumulation patterns and consequently devise plans of action.

Combining efforts will allow successfully achieving these goals and, ultimately,may lead to the needed reduction of microplastic contamination, though, consideringthe prevalence and pervasiveness of these materials already in the environment,elimination of such contamination is, in all likelihood, impossible.

Acknowledgments Thanks are due to FCT/MCTES for the financial support toCESAM (UID/AMB/50017/2019), through national funds. This work also received fundingfrom national funds (OE), through FCT, in the scope of the framework contract foreseen inthe numbers 4, 5, and 6 of Article 23, of the Decree-Law 57/2016, of August 29, changedby Law 57/2017, of July 19. This work is funded by project MicroPlasTox, with the referencePOCI-01-0145-FEDER-028740; by FEDER, through COMPETE2020 – Programa OperacionalCompetitividade e Internacionalizacão (POCI); and by national funds (OE), through FCT/MCTES.

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