Microplastic Extraction from Marine Vertebrate Digestive Tracts, Regurgitates and Scats: A Protocol for Researchers from All Experience Levels

Materials and Reagents

Note: All materials and reagents can be adapted depending on laboratory resources. For example, specific forceps and glass equipment are not required, but some specific precautions may be performed. See Table 1 for options.

1. Filter papers (e.g., glass microfibre filters, GF/D, 47 mm, pore size 0.47 microns, Whatman, catalog number: 1823-047)
2. Aluminum foil lids cut to the required size or non-plastic lids
3. Laboratory gloves (e.g., blue nitrile gloves, VWR, catalog number: 112-2373)
4. Mesh (e.g., 200 µm monofilament Nylon fibre mesh) 
5. Filtered water (e.g., Milli-Q Filtered water, 0.22 μm membrane filter)
6. Ethanol absolute > 99% (v/v) (VWR, catalog number: 20821.310E) 
7. Potassium hydroxide, KOH, 85.0%-100.5%, pellets (VWR, catalog number: 26668.296)
8. Sodium chloride, NaCl, 99.5%-100.5%, pellets (VWR, catalog number: 27810.364)
9. Sodium iodide, NaI, > 99.5% (VWR, catalog number: 27913.260)
10. Sodium polytungstate solution, SPT 85% (v/v) (Sigma-Aldrich, catalog number: 80656)
    

Equipment

1. 500 ml beakers (Corning, Pyrex^®, catalog number: 1395-500)
2. Magnetic stirrer (Sigma-Aldrich, Benchmark, catalog number: Z742550)
3. Vacuum filtering flask assembly for 47 mm filters with funnel, filter folder (e.g., 2,000 ml, VWR, catalog number: 511-0264H and Fisher Scientific, Kimble^TM, catalog number: K953771-0000) 
4. Vacuum pump (e.g., mini diaphragm vacuum pump, VP86, VWR, catalog number: 181-0065H)
5. Stainless steel sieves: 1 mm, 0.5 mm, 0.25 mm, and smaller sieves when a lower size limit is required
6. Cotton and white laboratory clothes
7. Non-plastic dissection tools (including scissors, forceps and scalpel)
8. Stereomicroscope with a camera (e.g., Leica Microsystems, model: MZ6)
9. FT-IR instrumentation (or similar equipment where possible for chemical analysis) 
10. Fridge
11. Incubator and shaker (e.g., New Brunswick Scientific, model: Innova 26R)
    

Procedure

1. Sample collection
   Here, we describe a step-by-step procedure for collecting samples from the field (Steps A1-A4, Figure 1). We understand that the methods used for collection can change depending on the samples targeted. The importance of contamination control steps is included as this information is required when collecting samples.
   
   
   Figure 1. Procedural steps for extracting plastics and microplastics from vertebrate digestive tracts, scats and regurgitates
   
   
   1. Collect digestive tracts following the standard protocols for animal necropsies (e.g., Kuiken and Garcia-Hartmann, 1991). Digestive tracts must be secured with a bridle or a cord to avoid loss of contents. If digestive tracts are not processed immediately, they should be stored in a durable container. It is recommended that when a plastic bag is used, that is made of a bright color. Therefore, in case of accidental contamination such plastic piece will be easy to identify.
   2. Scats and regurgitates are collected in transparent plastic zipped bags or transparent glass containers when possible.
   3. It is recommended that white gloves and white cotton laboratory coats/clothes be used during the collection of all samples. Any other color and polymeric material used must be noted for reference to take potential contamination into account when carrying out analysis.
   4. Samples should be processed fresh, but when this is not possible, samples can be frozen at -20 °C or lower until required.
   
   
2. Reagent preparation
   Caution: KOH is a caustic and irritant solvent. All researchers must wear laboratory gloves and eye protection at all times. If the solution touches skin, the affected area should be washed immediately with running water.
   
   1. To make a 10% KOH solution, add 100 g KOH to 1,000 ml of water. Solution volume can be adjusted depending on the sample processing requirements. It is recommended to prepare the solution in a beaker or conical flask with a lid.
   2. Close the lid and gently mix until pellets are dissolved, this can be carried out by shaking by hand or using a magnetic stirrer depending on the volume being prepared. Once pellets have been dissolved, the solution will have turned colorless, leave for at least 15 min to cool.
   3. The final solution should be filtered using a Buchner filter or similar (Table 1) to ensure that there are no residual microplastics in the solution, either from the KOH pellets, water or from procedural contamination.
      
   
   
3. Laboratory preparation
   
   1. The laboratory should be scrubbed before use to remove any sources of airborne contamination. There should be limited access to reduce external contamination.
   2. Researchers in the laboratory should wear white cotton laboratory coat and gloves. Any other color and polymeric material used must be noted for reference to take potential contamination into account when carrying out analysis.
   3. All equipment must be rinsed at least three times with pre-filtered water immediately before use. Glass or metal equipment is recommended, however, if unavoidable see Table 1. The water supply used for flushing the samples must be filtered using a sieve with a mesh size smaller than target microplastics. Any variation on the color of the equipment should be noted.
   4. Set up nested metal sieves in a sink connected to the pre-filtered water source. The sieve at the top will have the largest mesh size (e.g., 1 mm) and the bottom one the smaller one (e.g., 50 µm). Using consecutive sieves will allow removal of larger pieces of plastics, food remains and parasites. If researchers wish to target smaller plastics, they can add additional sieves. This allows for size differentiation and speeds up processing in later steps. The sieves should be covered with a lid when not in use to prevent contamination.
      
      Table 1. Troubleshooting advice and alternative methodological approaches
      
      
   
   
4. Sample preparation
   
   1. If samples have been frozen, it is recommended that samples be defrosted slowly in a fridge (24-36 h), to avoid the fast decomposition of the tissues. It is also possible to defrost samples at room temperature in a clean laboratory.
      Note: Depending on the sample collected there are different initial processing steps required. For digestive tract samples, proceed with Steps D2-D10, for regurgitate and scat samples proceed from Steps D11-13.
   
   Digestive tracts
   
   2. Digestive tracts must be rinsed externally before processing and removing the ties to remove external contamination.
      Note: Ties are used to stop contents leaking.
   3. Digestive tracts of marine organisms vary in length and number of compartments (esophagus, stomachs and intestines). Each compartment should be processed and rinsed independently (e.g., Hernandez-Milian et al., 2018).
   4. If different sections of fish digestive tracts cannot be differentiated, it can be analyzed without divisions.
   5. For large intestines (> 1 m, i.e., mammals), we recommend separating into smaller, equally sized pieces. The most appropriate number is 20 equal pieces. The division of this part of the digestive tracts will allow the researcher to investigate if microplastics tend to concentrate in any specific area as some parasites do (Lawlor et al., 1990), as well as make feasible the analysis of a large sample.
   6. Each compartment or intestine section must be opened using clean and sterilized scissors and forceps.
   7. A subsample for genetic analysis should be collected from the middle of the bulk of prey remains and stored frozen or in 70% pre-filtered ethanol.
   8. Transfer the remaining material to the nested sieve column avoiding touching the mucosal surface.
   9. Examination of the gastric and intestinal mucosal surface should be carried out with care to prevent damage to the surface by rubbing it with fingers or other material. If any pathology is detected, a sample of that area should be collected and stored with the fixed chemical required following the standard protocols.
   10. Compartments should be processed individually under the filtered water supply through nested sieves following Step E1 onwards.
       
       
   Scats and regurgitates (solid samples)
   
   11. The container where scats and regurgitates are stored must be rinsed externally with pre-filtered water before processing. This will avoid procedural contamination.
   12. A subsample from the middle of the sample will be collected and frozen or stored in 70% ethanol for genetic analysis.
   13. The rest of the samples should be poured and processed in the nested sieves following Step E1 onwards. If the scat or the regurgitated sample is not fluid, they can be poured into a container with pre-filter water for few hours to hydrate before processing. Samples must always be covered with a lid to prevent procedural contamination.
   
   
5. Sample processing
   
   1. Rinse each sample or compartment into the nested sieves separately (Figure 1).
   2. Transfer the non-plastic material collected in both top sieves to containers with 70% ethanol for between 2 and 24 h to sterilize the material and prevent mold and odor.
   3. Store hard food remains (e.g., otoliths, bones, shells) dry in bags or containers, while cephalopod beaks, and other soft remains (e.g., crustaceans, worms, parasites) should be stored in 70% ethanol for further identification (Figure 1).
   4. Rinse the plastic material (> 1 mm) found in the larger sieve with pre-filtered water and store dry. Material kept in the smaller sieve will be transferred to a glass container with pre-filtered water to obtain a suspension for the microplastic sample (Figure 1).
   5. The suspension should be as concentrated as possible. This will reduce the volume of KOH solution required in the following step.
   Note: Eye protection must be worn from this step.
   
   6. Add 10% KOH solution to the suspension in a ratio of 3:1 (KOH:suspension) (Figure 1).
   7. Cover samples loosely with aluminum foil, or screw top lids to prevent contamination and evaporation.
   8. Incubate samples at 60 °C for 24 h with continuous agitation at 125 rpm; alternatively, heat can be forgone but the reaction will take longer (e.g., 3 weeks, Foekema et al., 2013).
   9. The solution will turn transparent/slight yellow in coloration when all biological material has been digested.
   10. Remove samples from the incubator and leave to cool before further processing.
   11. After cooling, filter solution under vacuum using equipment such as a glass Buchner filter with a microfiber filter (GF/D or alternative). Alternatively, if this equipment is unavailable, glass funnels with a microfiber filter covered during the filtration can be used.
       Note: Filter papers must be checked under a microscope for contamination before use.
   12. When large amounts of undigested organic material (e.g., bones) remain after filtering, density flotation can be used to separate undissolved organic material from low density plastics which will float. Great care is required when using density separation as there are some less dense dietary remains that can float, such as crustacean carapaces. NaCl is the recommended density separation solution (1.2 g cm^-3). More costly solutions include NaI (1.8 g cm^-3) and SPT (2.8 g cm^-3).
   13. Another option to remove undigested organic matter involves rinsing the solution through a sieve once more before filtering; this reduces the likelihood of filter papers clogging.
       
   
   
6. Visual characterization
   Visual characterization is valuable when researchers are categorizing and sorting samples. Visual characterization of plastics must be used with other more robust identification techniques such as chemical analysis (e.g., Steps F4-F5, G1-G7), even if large plastics can be recognized. If more robust techniques are not available, Steps F4-F5 can aid researchers in reducing the likelihood of misidentification.
   
   1. Use a stereomicroscope to investigate particles retained on the filter paper. It is recommended that a camera be attached to the microscope to allow researchers to record visual images of all particles.
   2. Visually inspect each filter paper for potential plastic microparticles.
   3. Carry out visual characterization following existing criteria (Lusher et al., 2014):
      
      1. Measured longest dimension (mm) and smallest aperture lengths.
      2. Record shape and color (Figure 1).
         
         1. Shape categories include: fiber, fragment and spherical (beads). Fragment can be further divided into films and foams depending on research objectives, but this division is not always necessary.
         2. Color is subjective and therefore not recommended as a stand-alone classification, but it does help researchers when categorizing samples.
   4. Particles observed need to be visually inspected for the following characteristics, otherwise they should be rejected or tested with other chemical techniques (Lusher et al., 2014):
      
      1. Homogeneous color.
      2. No natural structures (such as cells).
      3. Unnatural bending.
      4. Fibers should have a consistent thickness throughout length.
      5. There should be no fraying at ends of fibers.
   5. A hot needle can be used during visual characterization to aid in the presence of plastic particles, which in case of plastic will react to the heat through bending or melting. This method has been recommended by National Oceanographic and Atmospheric Administration (NOAA) and the European Union under the Marine Strategy Framework Directive (EU-MSFD) however the hot needle is not suitable for semi-synthetic fibers as they do not react. Therefore, some research groups are advised to take this step with caution.
   
   
7. Chemical characterization
   Note: Chemical characterization should only be carried out on clean and dry suspected plastic particles.
   
   1. Chemical characterization should be carried out on a representative subsample. MSFD recommends 10% although as many particles as possible should be analyzed chemically to reduce identification errors.
   2. The specific instrumentation used for chemical classification will depend on the research facilities available. Here we describe FT-IR because this instrumentation is available at most of research laboratories (Figure 1).
   3. Trained personnel should conduct chemical analysis. Sufficient knowledge of polymer identification and spectra interpretation is required.
   4. Spectral analysis should follow the protocols available at individual laboratories to produce an output spectrum.
   5. All output spectra should be compared to a polymer library database.
   6. Only polymers which matched reference spectra with a high level of certainty (> 70% match) should be accepted as correct identification.
   7. Researchers with sufficient knowledge of polymers may also consider visual inspection of produced spectra that have a lower level of certainty (60%-70%). Caution should be taken for such interpretation. One example is water absorbance which can alter spectra and that have clear polymeric characteristics.

Data analysis

This protocol is a standard protocol for the collection of microplastics. Data analysis is subject to the specific user requirements of the protocol. It is recommended that researchers assess the number of particles in each identified compartment (stomach, intestine section or individual scat and/or regurgitate). Values obtained can be compared with other samples from the same research area or further afield. Specifically dividing the digestive tract and intestines allows researchers to compare microplastic levels between and within species (Lusher et al., 2018).

Acknowledgments

The authors obtained no external funding for the elaboration of this protocol, A.L.L. and G.H.M. acknowledge their previous funders which supported their earlier investigations. A.L.L. was funded by an Irish Research Council Postgraduate Scholarship and a GMIT 40th anniversary studentship between 2012-2015. G.H.M. was supported by a Beaufort Ecosystem Approach to Fisheries Management award, as part of the Irish Government’s National Development Plan (NDP) between 2013-2015. Both authors thank National Parks and Wildlife Service (NPWS) for reporting the strandings to the Irish Whale and Dolphin Group (IWDG).
Author Contributions: A.L.L. and G.H.M. conceived and designed the protocol; performed the experiments, analyzed the data; and wrote the paper equally.
This protocol is the culmination of adaptions from previous work by both the authors.

Competing interests

The authors declare no conflict of interest.

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