| Processing Steps |
- Parameter or Variable: microplastic concentration (measured); Units: count of MPs/individual; Observation Category: in situ; Sampling Instrument: dredge; Sampling and Analyzing Method: Six molluscs species were selected among those most widely and frequently consumed by the Catalan population. Wedge clams, Donax trunculus (WC), marine snail, Bolinus brandaris (S), razor clams Ensis siliqua (RC), fine clams Tapes decussatus (FC), big oyster Crassostrea gigas (BO) and mussels Mytilus galloprovincialis (M) were included in this survey. Molluscs species were sampled in October–November 2017 from biggest anglers’ cooperatives along the Catalan coast (377 km). The Catalan areas selected for the study were the following: Barcelona (Central), Girona (North) and Tarragona (South). Between 2 and 4 kg of each sample type of molluscs were purchased and acquired by donation fisherman cooperatives. The samples were wrapped in aluminum foils and kept refrigerated during the transport to the laboratory. Subsequently, samples were rinsed with ultrapure water (distillate water filtrate to 0.45 μm through filters), being external byssus threads removed. For representability, >50 individuals were analyzed for each sample of mussels, razor clams, fine clams, wedge clams, and snail according to depuration conditions and catchment zones. In turn, up to 28 individuals were analysed for oysters. A number of soft tissues sub-samples (50–70 g) were prepared for analysis, being these sub-samples lyophilized (Telstar Cryodos-50/80) during a period of 48 h in a clean and fiber free beaker. All subsamples were kept at -20 ◦C until analysis. A general protocol for MPs ≥ 20 μm extraction in 50–70 g subsamples of soft tissues was partially -or completely- applied, depending on molluscs sample characteristics. The removal of organic and inorganic matter from the sample was carried out by three phases and an additional density separation step. The first phase required 4 days for complete procedure, while the second one 8 days, and the third one 4 days, with a total of 16 days. The density separation step took one additional day. The first phase (KS) is a combination of alkaline hydrolysis (KOH 2 M) (K) and surfactants (SDS 10 % (S) at 40 ◦C with agitation. The second phase (E þ H), consisted in enzymatic hydrolysis with protease, lipases, and celluloses (E), followed by an oxidation with hydrogen peroxide 33–35 % (H), at 40 ◦C and gently agitation. The third step (F þ Ch) involved wet peroxide oxidation (Fenton processes, oxidation in presence of a Fe (II) catalyst to digest labile organic matter) (F), and enzymatic hydrolysis with Chitinase (Ch) at 40 ◦C and agitation. Finally, in the case of presence of inorganic matter, density separation (DS) with ZnCl2 solution (1.8 g/mL) was carried out. Each phase should be applied based on the amount of organic matter removed in the previous phase through decision steps. Once the organic matter removal has been achieved (based on interference grade for easy particles detection and identification), the process stops. The sample is then ready for density separation steps, or for identification, thereby it is not necessary to complete all phases. The application of alkaline, enzymatic hydrolysis and Fenton processes in phases and sequential steps offer an effective MPs extraction and their different morphologies at less time with less airborne fibers contamination, avoiding significant damages on surface of particles, and reducing costs for chemical reagents. The percentage of removal of organic matter obtained with the application of the three phases is up to 98 %. The chemical reagents and filters were the following: absolute ethanol (Scharlau, >99,9 %, Barcelona-Spain), potassium hydroxide (Sigma-Aldrich, 99,9 %, Madrid-Spain), hydrogen peroxide stabilized (PamReac AppliChem ITW Reagents 33 % w/v Barcelona-Spain), hydrogen peroxide aqueous solution stabilized (Thermo-Scientific Acros 33–35 % w/v, Madrid-Spain), zinc chloride (Acros Organics,>98,0 %, Madrid-Spain), zinc chloride anhydrous (Sigma-Aldrich, ≥98 %, Madrid-Spain), SDS (PamReac AppliChem ITW Reagents, 99 %, Barcelona-Spain), iron (II) sulphate heptahydrate (Sigma-Aldrich, >99,0 %, Madrid-Spain), nitric acid (Scharlau, 2 N, Barcelona-Spain), sodium acetate anhydrous (Scharlau, 99 %, Barcelona-Spain), acetic acid (Scharlau, 96 %, Barcelona-Spain), Tris-HCl (Scharlau, 99 %, Barcelona-Spain), PTFE 5, 10 μm pore size filters (Sartorius, Madrid-Spain). The enzymes used were Chitinase 1000 U/mL, Celullase TXL >1000 U/mL, Lipase FE-01 > 18,000 U/mL, Protease A-01 > 1000 U/ mL provide by ASA Spezialenzyme GmbH (Wolfenbüttel-Germany). All particles suspected of being MPs were totally quantified on PTFE filters of each subsample using both, a stereoscopic (LEICA MZ10 coupled to FLEXACAM C1) camera, and an optical microscope (Olympus CX41). Microplastics were classified according to the morphology as fibers (including filaments and fishing line), films (particles with a two-dimensional shape), fragments (particles with a three-dimensional shape), and pellets (solid spheres; solids foam spheres). The biggest and medium particles were placed on a clean PTFE filter, and the smaller ones (<0.5 mm) into a square of 1 cm2 area on calcium fluoride (CaF2) slide. For smaller MPs and abundant particles ≤50 μm, the original filters were divided in four quadrants. In each section, four fields from 40 to 80× magnification were randomly chosen, being particles of each morphology and sizes measured. The CaF2 slide and filter were photographed and the particles length was measured with ImageJ software. Polymeric composition analysis was carried out for all photographed and measured particles. Fragments and films between 0.5 and 5 mm for each subsample were grouped and placed on clean PTFE filters. Fibers, fragments and films from 0.02 to 0.5 mm were arranged on calcium fluoride (CaF2) slides. Particles size larger than 0.5 mm were analysed by infrared spectroscopy technologies with a Perkin Elmer Frontier instrument with a Spectrum Software and an Attenuated Total Reflectance (ATR) accessory. For the identification of each polymer, the spectra obtained with the ATR-FTIR and μFTIR techniques were performed, being the unknown spectra compared with OMIC software libraries database. It includes HR Nicolet Sampler Library, Hummel Polymer Sample Library, Polymer Laminate Films, Wizard Library, and an own library with >80 IR spectra. Only match spectra with major -or equal- than 75 % of similarity with reference spectra were accepted. The rejected items were counted as the temporary unidentified category. The unidentified spectra were also identified comparing with BIO-RAD IR from University of Barcelona (Barcelona, Spain) spectral databases. The matches of similarity major or equal than 75 % were accepted. For microplastics measurement per gram of dry weight (MPs/g d.w.), data is reported as a mean for each molluscs type.; Data Quality Method: To prevent contamination, distilled water (MilliQ® water), and all solutions prepared, were filtered before the use through sterilized nitrocellulose membrane filters GF/F 0.45 μm (Whatman 47 mm diameter, Maidstone, United Kingdom), and stored in glass bottles. All glassware, sieves and fine tools as scalpels, and stainless-steel tweezers, were washed with ultrapure water (distilled water filtrate through 0.45 μm sterilized membrane filters) and alcohol 70 % after washing. All materials were wrapped in aluminum foil and stored in a clean fume hood. All laboratory countertops were cleaned with alcohol 96 % and fiber-free napkins. Clean cotton laboratory coats and latex gloves were used during all processes. All glass material was immediately covered to reduce MPs airborne contamination in extraction and lyophilization procedures. Samples processing and digestion procedures were conducted in clean fume hood and laminar flow cabinet. Blanks were used in every step of samples processing and digestion. Five and 10 cleaned Petri dishes were placed during the samples processing and during MPs visual sorting, respectively, in order to calculate the deposition of airborne fibers. Eleven procedural blanks, with the same reagents volume that the samples, were run with every set of samples in digestion processes fulfilling the methodology, step by step and by phases. Finally, three blanks in lyophilization processes were applied; one for every set of sub-samples. The results were corrected by fibers and particles contaminations. After processing and preparations of sub-samples, lyophilization procedures and visual sorting of MPs analysis, airborne fibers and particles contamination, were not detected in any blank. However, after the digestion process fibers were found in 91 % of the controls. The fibers in controls assigned to every set were classified by color, being subtracted from the total quantity found in the respective subsamples. The numbers of fibers in controls ranged from 3 to 17 and the colors were the following: blue, yellow, red, black, grey, green, and transparent. To ensure methodology effectiveness in the MPs extraction from the samples, recovery rates were calculated in parallel with samples analysis. Samples of three molluscs were randomly chosen and spiked with colored virgin plastic spherical particles of polyethylene (PE) spanning from 53 to 500 μm diameter with density 1.02–1.06 g/cm3 (purchased from Cospheric Inc., California, USA). Subsamples around 50–55 g of soft tissues of molluscs were dissected for spiking. The tissues were carefully cut and inoculated for different parts with colored PE mix spheres ranging from 53 to 500 μm. The recovery rates were 60 % for 53–63 μm, 60 % for 125–150 μm, 74 % for 250–300 μm, and 99 % for 425–500 μm. Neither damages by agitation (mechanic forces), nor chemical attacks, were found in spheres..
- Parameter or Variable: microplastic concentration (measured); Units: count of MPs/g w.w.; Observation Category: in situ; Sampling Instrument: dredge.
- Parameter or Variable: microplastic concentration (measured); Units: count of MPs/g d.w.; Observation Category: in situ; Sampling Instrument: dredge.
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