Plastic and Microplastic in the Environment. Группа авторов
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Figure 1.4 Possible exposure pathways of MPs into freshwater environments.
Routes of secondary MPs into freshwater environments are mainly from improper management of plastic wastes. This includes release during collecting, transporting, processing, and landfilling of solid waste. Another route could be the runoff of wastes through drainage canals from agricultural land. Plastic wastes on roads such as vehicle debris, tire wear particles, or fragments of road‐marking paints could overflow with storm water into the freshwater environments (Eriksen et al. 2013). Lighter plastic wastes may be transported into freshwater environments by wind (Zylstra 2013), and synthetic fibers could be carried and accumulated by atmospheric fallout (Dris et al. 2015).
1.4 Microplastic Analytical Methods in Freshwater
Until now, there has been no certain methodology for sampling and analysis of MPs in freshwater. Most researchers adopted methods from the marine environment, with modifications. However, homogeneous and standard methods for the determination of MPs are missing. This fact hampers comparability between studies both in freshwater and marine environments. Different techniques for sample collection, preparation, and analysis of MPs are summarized in Figure 1.5.
1.4.1 Sampling of Microplastic
1.4.1.1 Water Samples
As reported in previous studies, the abundance of MPs in water samples is lower than that of sediment samples (Ta and Babel 2020b). Thus, a large quantity of water is usually collected to acquire a representative sample. Volume reduced samples were commonly applied for the sampling of surface water. Until now, most studies used manta trawls or plankton nets to collect water samples. Different mesh sizes of these nets are used, ranging from 50 to 3000 μm (Tan et al. 2019; Zhang et al. 2015), while 333 μm is the most commonly used in all studies (Anderson et al. 2017; Free et al. 2014; Su et al. 2016; Ta and Babel 2020b; Wong et al. 2020). Flow meters are attached to plankton nets or manta trawls to determine the filtered water volume. This helps to normalize the number of MPs to the volume of filtered water. Another technique is the measurement of the sampling area by the Global Positioning System (GPS). Thus, results are presented as the number of MPs per surface area. Trawling speed depends on the water velocity or wave action, but usually ranges from one to five knots (Ta and Babel 2020a; Tan et al. 2019; Wong et al. 2020). Another sampling method is direct filtration of water through sieves or by the collection of batch samples (Crew et al. 2020; Yan et al. 2019). In this method, a smaller water volume can be collected, thus the sample representability is reduced (Wang et al. 2017).
Figure 1.5 Techniques reported in the literature for identifying MPs in sediment and water samples.
1.4.1.2 Sediment Samples
The sampling of sediment samples can be separated into the collection from the shoreline and bottom of the river or lake. For shore sediments, sampling strategies are transected sampling perpendicular, random sampling, and sampling in single squares or parallel to the water. Most studies applied the grid sampling method with depths of 2–5 cm on the surface layer (Jiang et al. 2019; Klein et al. 2018). Frame and corers are usually used to determine the sampling area. Non‐plastic tools such as scoop, trowels, or shovels, and non‐plastic sampling vessels are required (Alam et al. 2019; Jiang et al. 2019; Peng et al. 2018). Bottom sediment from the riverbed or lakebed can be carried out with grab samplers such as Ekman or Van Veen grabs or corers (Alam et al. 2019; Fan et al. 2019; Ta et al. 2020c; Wang et al. 2017). The sediment samples collected by grab methods are usually disturbed, therefore this is suitable for surface layer (top 5 cm) or bulk sampling. Conversely, sampling by cores allows determining MP depth profiles and undisturbed surface and depth layers. Nevertheless, the number of samples that can be collected is limited. According to Dris et al. (2018), river bottom sediments are mostly collected by grabs, while corers or grabs are used for lake bottom sediments. The number of MPs is usually normalized to the sediment volume or weight, and sampling area.
1.4.2 Sample Preparation
1.4.2.1 Extraction of Microplastics
Due to the complex nature of the sediment, MPs in the samples must be extracted from sample matrices. The density separation is widely applied to extract MPs from sediment samples. The sediment is dried prior to mixing with a concentrated salt solution. After a period of agitation, MPs and light particles float to the surface or stay suspended, whereas heavy particles settle down (Klein et al. 2018). Many studies extract MPs by using sodium chloride (NaCl) solution since this is inexpensive and environmentally friendly (Alam et al. 2019; Campanale et al. 2020; Free et al. 2014; Mani et al. 2015). However, the density of NaCl solution (~1.2 g/cm3) cannot extract some polymers such as Polyvinyl chloride (PVC), Polyethylene terephthalate (PET), polycarbonate, and polyurethane. Therefore, sodium iodide (NaI), sodium zinc chloride (ZnCl2), and sodium polytungstate (Na2WO4) are viable choices (Ballent et al. 2016; Ta and Babel 2019; Yin et al. 2020). Conversely, MPs in water samples are easily filtered and separated during the sampling step (Dris et al. 2018).
1.4.2.2 Removal of Organic Debris
The detection of MPs is hampered by organic particles that accompany MPs during collecting water samples. Thus, the removal of the organic debris is important to reduce the misidentification or underestimation of MPs. The detection step is conducted before or after the density separation. Organic debris is usually removed by strong acids (Cole et al. 2014; Imhof et al. 2016) or base solutions (Dehaut et al. 2016), or oxidation agents such as hydrogen peroxide (Ta and Babel 2019; Zhao et al. 2017). Some sensitive polymers (i.e. poly(methyl methacrylate) and polycarbonates) can be lost or damaged during the treatment (Dehaut et al. 2016; Li et al. 2018). Alternate chemicals for removing organic debris in samples are enzymes; these chemicals reduce damage to sensitive polymers. (Catarino et al. 2017; Mani et al. 2015). The method can be conducted by using proteinase K or mixtures of technical enzymes including proteinase, lipase, chitinase, amylase, and cellulose (Courtene‐Jones et al. 2017; Mani et al. 2015). The enzymatic digestion should be conducted under controlled conditions of pH and temperature. However, several disadvantages of using enzymes are reported. In comparison to chemical treatments, enzymatic treatments are expensive, time‐consuming, and may not completely remove the organic material (Courtene‐Jones et al. 2017).
1.4.3 Identification of Microplastic
1.4.3.1 Visual Sorting
In most studies, visual sorting is the first step to separate MPs from samples before identification of the polymer type. Large MPs (> 1 mm) can be recognized by the naked eye (Anderson et al. 2017), while smaller particles are identified using dissection microscopes (Faure et al. 2015; Mani et al. 2015) or scanning electron microscopy (SEM) (Eriksen et al. 2013; Su et al. 2016). This step requires experienced researchers and good optical quality of the microscope. However, identification of all particles is difficult if they are smaller than a certain size, if they are unable to be distinguished visually or cannot be managed with forceps due to their minuteness. Thus, visual sorting is time‐consuming and easy misidentification or underestimation of MPs is possible. Recently, another visual identification method using fluorescence was applied to detect and quantify small MPs. In most studies, Nile Red (NR) was used and dissolved in different solvent solution such as acetone, chloroform, and