Section IV: Sampling and Analysis


: Sampling, Isolating and Digesting of Microplastics

Leo M. L. Nollet CONTENTS

  • 7.1 Sampling 103
  • 7.1.1 Field-Collected Organisms 103
  • Microplastic Losses during Field Sampling 104
  • Microplastic Accumulation during Field Sampling 104
  • Sample Storage 104
  • 7.1.2 Laboratory-Exposed Organisms 104
  • 7.2 Isolating Microplastics 105
  • 7.2.1 Dissection 105
  • 7.2.2 Depuration 105
  • 7.3 Digestion 106
  • 7.3.1 Nitric Acid 107
  • 7.3.2 Other Acids 107
  • 7.3.3 Alkalis 107
  • 7.3.4 Oxidizing Agents 108
  • 7.3.5 Sodium Hypochlorite 108
  • 7.3.6 Enzymes 109
  • 7.3.7 Effect of Temperature 109
  • 7.4 Filtering Digestants 109
  • 7.5 Density Separation 111

References 111

A. L. Lusher et al. [1] published an interesting and valuable article on sampling, isolating and identifying microplastics ingested by fish and invertebrates.


Field-Collected Organisms

Micro- and nanoplastics are taken up by a wide range of organisms in a diverse range of habitats, including the sea surface, water column, benthos, estuaries, beaches and aquaculture [2]. The diversity of the organisms and habitats where they live and are sampled require a range of collection techniques [3-5]. The sampling method is determined by the research question, available resources, habitat and target organism. Benthic invertebrate species such as Nephrops norvegicus may be collected in grabs, traps and creels, or by bottom trawling [6,7]. Planktonic and nektonic invertebrates are collected by way of manta and bongo nets [8-11]. Fish species are generally recovered in surface, midwater and benthic trawls, depending on their habitats. Gill nets have been used in riverine systems [12]. Some species are collected from the field by hand; this is common practice for bivalves, crustaceans and annelids [13-17]. Another method is direct collection from shellfish or fish farms [18-20] or from commercial fish markets, where the capture method is often unknown [21-22].

Microplastic Losses during Field Sampling

Handling stress, physical movement and the physiological and behavioral specificities of the sampled organism may result in the loss of microplastics prior to animal preservation. Gut evacuation times for animals range from minutes for decapod crustaceans to several hours for calanoid copepods [10] and fish [23,24] to days in larger lobsters [25]. Some animals might egest microplastic debris prior to analysis [7]. In such cases, the time between sample collection and the preservation of the animal must be as short as possible. Care must also be taken to minimize handling stress or physical damage.

The copepod Eurytemora affinis [26] and some fish species have been observed regurgitating their stomach contents [27]. Compression of a catch in the cod end might induce regurgitation in fish [29]. The likelihood of regurgitation increases with depth of capture, and gadoids are more prone to regurgitation than flatfish. Piscivorous predators are prone to regurgitation owing to their large distensive esophagus and stomach [28, 30].

Microplastic Accumulation during Field Sampling

Laboratory studies have identified that nano- and microplastics can adhere to external appendages of marine copepods [10]. Cataloguing such interactions in nature is complicated as determining whether the resulting accumulation has occurred naturally or as a by-product of the sampling regimen is difficult. A similar interaction may occur with organisms feeding on microplastics during capture in nets; this is particularly of concern when the mesh size of the net is capable of collecting microplastics, for example, in manta nets (common mesh size 0.33 mm) [23].

Sample Storage

It is important how biotic samples are stored. The choice of the preservation technique will largely depend on the following analysis. It is important to know if the fixative will affect the structure, microbial surface communities, chemical composition, color or analytical properties of microplastics within the sample. Four percent formaldehyde and 70% ethanol are commonly used fixatives; however, these preservatives at higher concentrations can damage some polymers. Polyamide is only partially resistant to 10% formaldehyde solution, while polystyrene can be damaged by 100% alcohol. Alternative methods for storage of organisms include desiccation [9] and freezing [7,31-33].

Laboratory-Exposed Organisms

Laboratory studies are used to better understand the interactions between microplastics and biota. Controlled laboratory exposures facilitate monitoring of the uptake, movement and distribution of synthetic particles in whole organisms and excised tissues. Fluorescently labeled plastics, either purchased or dyed in the lab [34], allow visualization of microplastics in organisms with transparent carapaces, [10,18,35] circulatory fluids, (36,37) or histological sections [38]. Where dissection is prohibitive (e.g., mussels) fluorescent microplastics can be quantified by physically homogenizing tissues followed by microscopic analysis of sub-sampled homogenate [14]. Coherent anti-Stokes Raman scattering (CARS) has also been used to visualize non-fluorescent nano- and microplastics in intestinal tracts and those adhered to external appendages of copepods and gill lamellae of crabs [10,14]. Bioimaging techniques, however, are not feasible, with field sampled biota as environmental plastics do not fluoresce and may be obscured by tissues or algal fluorescence.

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