Methodological Review

For this literature review, original peer-reviewed research articles, gray literature and conference proceedings from the 1970s to July 2016 were examined. Literature referring to the extraction of microplastics from marine, freshwater and terrestrial biota using Web of Knowledge, Science Direct Scopus and Google Scholar were identified. The authors also mined the journals Marine Pollution Bulletin, Environmental Pollution and Environmental Science and Technology, owing to the regularity with which they publish relevant material. Analysis of microplastic specific studies was expanded to include historical literature that did not necessarily have microplastics as the central theme of the research, such as studies which used fluorescent latex beads as a tracer for feeding and retention experiments. Of the 120 papers included in this meta-analysis, 58.3% of studies were conducted in the laboratory, 38.3% focused on organisms collected from the wild and 3.4% involved both laboratory exposure and field collection (Figure 8.2). There were 96 studies wholly focused on marine organisms, 21 on freshwater, two studies on both marine and freshwater organisms and one published study on a terrestrial species.

Studies of biota interactions with microplastic in the laboratory and field

FIGURE 8.2 Studies of biota interactions with microplastic in the laboratory and field.

Sampling

Field-Collected Organisms

Observations of microplastic uptake by environmentally exposed organisms have now been reported in a range of habitats, including the sea surface, water column, benthos, estuaries, beaches and aquaculture [4]. The diversity of the organisms studied and the habitats from which they are sampled require a range of collection techniques [123]: the sampling method employed 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 [34,41], and planktonic and nektonic invertebrates by way of manta and bongo nets [10,12,14,16]. Fish species are generally recovered in surface, midwater and benthic trawls, depending on their habitats [69-92]. Gill nets have been used in riverine systems [102]. Some species are collected from the field by hand; this is common practice for bivalves, crustaceans and annelids [21,35,37,42,56]. Another method is direct collection from shellfish or fish farms [15,55,56] or from commercial fish markets, where the capture method is often unknown [58,103]. Avoiding contamination and biases during sampling and sample analysis is paramount, and mitigation protocols are described below.

8.2.1.1.1 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 are varied, ranging from as little as 30 minutes for decapod crustaceans (N. Welden, personal observations), <2 hours for calanoid copepods [10], 10-52 hours for fish [68,124] to over 150 hours in larger lobsters [125]. Therefore, some animals might egest microplastic debris prior to analysis [41]. 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. This will reduce the potential for microplastic regurgitation; the frequency with which animals expel consumed plastics during sampling is unknown. The copepod Eurytemora ajfinis [126] and some fish species have been observed regurgitating their stomach contents [127]. The main cause of regurgitation in fish is thought to be related to the expansion of gas in the swim bladder: this causes the compression of the stomach and may, in extreme cases, result in total stomach inversion [128]. Compression of a catch in the cod end might induce regurgitation in fish [129]. 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 [128,130]. As such, regurgitation may bias the stomach content estimation, affecting consumption estimates and the presence of plastic debris.

8.2.1.1.2 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 natura is complicated as determining whether the resulting accumulation has occurred naturally, or as a by-product of the sampling regimen, is prohibitive. While most studies focus on the consumption of plastic, any research considering external adherence of microplastics should be aware that observed entanglement may have occurred during sampling and may be unrepresentative of microplastic-biota interactions at large. A similar interaction may occur with organisms feeding on microplastics during capture in nets; this is particularly a concern when the mesh size of the net is capable of collecting microplastics, for example, in manta nets (common mesh size 0.33 mm) [69]. Control of microplastic contamination is discussed in Section 8.2.4.

8.2.1.1.3 Sample Storage

Consideration should be given to the storage of biotic samples. Choice of preservation technique will largely depend on the research question being considered; for example, will the fixative affect the structure, microbial surface communities, chemical composition, color or analytical properties of any microplastics within the sample? Four percent formaldehyde and 70% ethanol are commonly used fixatives; however, consultation of resistance tables suggests these preservatives, albeit at higher concentrations, can damage some polymers; for example, 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 [12] and freezing [41,77,83,89].

Laboratory-Exposed Organisms

Laboratory studies have been implemented 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 (e.g., gills, intestinal tract, liver). Fluorescently labeled plastics, either purchased or dyed in the lab [17], allow visualization of microplastics in organisms with transparent carapaces [10,15,30], circulatory fluids [47,49] or histological sections [105]. Where dissection is prohibitive (e.g., mussels), fluorescent microplastics can be quantified by physically homogenizing tissues followed by microscopic analysis of subsampled homogenate [35]. 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 [10,35]. Bio-imaging techniques, however, are not feasible with field-sampled biota, as environmental plastics do not fluoresce and may be obscured by tissues or algal fluorescence.

 
Source
< Prev   CONTENTS   Source   Next >