Occurrence of microplastics in edible aquatic insect Pantala sp. (Odonata: Libellulidae) from rice fields

Study area

Samples were taken from a rice field in the Kasetsart University Kamphaeng Saen campus, Kamphaeng Saen district, Nakhon Pathom province, central Thailand (N14°00′32.2474 E99°58′54.1744) (Fig. 1).

Collection and identification of samples

Aquatic insects were sampled at random along the edge of the rice plots in October 2020 using an aquatic dip net (dimensions 30 × 30 cm, 250 m mesh, 50 cm length). According to our preliminary samplings, several aquatic insects were discovered towards the margin of the rice plot, due to the greater volume of water. As a result, this section of the rice plot was sampled. For such a rice plot, samples were taken along 3 m on each side. Aquatic insect collections were made in each plot by dragging a dip net down the ground for 3 m along the margin of the rice plot. Because aquatic insects have high water content, aquatic insects caught in the net were collected and preserved in vials containing 95% alcohol. The standard keys were used to identify aquatic insects up to the lowest taxonomic level (Dudgeon, 1999; Yule & Yong, 2004). The nymph of Pantala sp. (Odonata, Libellulidae) was used in this research as an aquatic insect for MP analysis (Fig. 2). In addition, more Pantala sp. were collected in the same rice field for the MPs’ investigation in October 2021.

Preparation of Pantala sp. larvae

Only the Pantala sp. taxa were used for the microplastic investigation. For the study, a total of 180 Pantala sp. specimens of comparable weight were analyzed. Three times distilled water was used to wash the fresh preserved specimens. To assess MPs from Pantala sp., 5 specimens (×18 replicates) of the whole body, 5 specimens (×18 replicates) of the gastrointestinal tract (GT) alone, and 5 specimens (×18 replicates) of the body without the GT were pooled. The specimens were dissected individually using scissors and forceps. All replicates were transferred to a 25 mL erlenmeyer flask. The entire body, the GT, and the body without the GT weight were all measured for wet weight. The results were presented as a mean ± standard deviation.

H₂O₂ treatment

Each pooled sample was placed in one erlenmeyer flask and eighteen replicates were prepared for each sample type. To digest the organic materials, 10 mL of a hydrogen peroxide solution (30% H₂O₂) was added to each flask (Ehlers, Manz & Koop, 2019). After that, the flasks were wrapped in parafilm and stored at room temperature for 7 days.

Floatation and filtration

Following tissue disintegration, potassium formate (99%) was employed for density of separation the resulting dissolved liquid from soft tissues (Ehlers, Manz & Koop, 2019). Each sample was placed in a glass separatory funnel, which was then filled with approximately 16 g of potassium formate and shaken to separate the solution. Because of its less dense form, saturated potassium formate solution facilitates the separation of the microplastic layer after about 4 h. The undissolved inorganic residues were then drained, and the supernatant was vacuum filtered onto nylon membrane filters (pore size of 0.45 µm; diameter of 47 mm). The filter was then placed in clean glass petri dishes with aluminum foil covers and dried for two days in a 50 °C drying cabinet.

Contamination control

To avoid contamination from airborne MPs, all containers and equipment were cleaned with distilled water and covered with aluminum foil when not in use. Exclusive gloves (nitrile), steel, and glass devices were always used at the laboratory. Before beginning labwork, the scissors and forceps were rinsed three times with deionized water. Lab surfaces were thoroughly cleaned with 70% ethanol. At every stage of the analysis, blanks were run without tissues in parallel with the same procedure used for the samples. All the experimental procedures were finished as soon as possible.

Microplastic observation and polymer identification

Under a stereomicroscope (Leica EZ4E) with 35x magnification, the filters were visually examined, and photos were obtained at various magnifications to identify MP particle based on their color and type. The MP particles were recorded. A PerkinElmer Spectrum-Fourier transform infrared spectrometer (FT-IR) in attenuated total reflection (ATR) mode was used to verify selected particles (range size 400–500 µm). The spectral range was 4,000 to 500 cm⁻¹, with a 32 cm⁻¹ spectral resolution and 16 co-scans for each measurement. The characterization of functional groups and the analysis of polymer types were compared to the Bruker spectrum library. Considering the spectrum analysis, the matching degree of spectra with a quality index ≥0.7 was accepted (Woodall et al., 2014).

Data analysis

The abundance, types, and colors of MPs were counted. Non-parametric analyses were applied as the abundance of MPs was not a normal distribution among the three sample types. Kruskal–Wallis H test was used to test for differences in the mean abundance of MP between sample types, using the Statistical Package for the Social Sciences (SPSS) version 19. Statistical significance was defined as a p-value of less than 0.05 (p < 0.05).

Abundance, type and color of MP in edible aquatic insects

The mean wet weights of Pantala sp. whole body, gastrointestinal (GT) tract only, and body without the GT were 0.3098 ± 0.0795, 0.0399 ± 0.0133 and 0.2445 ± 0.0707 g, respectively. In the controls, no MP particles were found in the blanks. The total number of particles were 121, 95, and 66 in all eighteen replicates of pooled samples (Figs. 3A–3C), with mean abundance per individual of 1.34 ± 1.11, 1.06 ± 0.77 and 0.73 ± 0.51 items in the whole body, gastrointestinal (GT) tract, and body without the GT, respectively. Kruskal-Wallis H test revealed no significant differences in the mean abundance of MP particles among sample types (chi-squared = 2.774, df = 2, p = 0.250) (Table 1). Different types of MPs identified in three samples were fragment, fiber, and rod (Figs. 3D–3F). The colors of MPs were shown in five different shades of red, green, blue, violet, and orange (Figs. 3G–3I).

Fragments and fibers were the most common particle types in edible aquatic insects (Odonata), which was similar to prior observations in Nigerian freshwater insects (Akindele, Ehlers & Koop, 2020). They were found in different sample types in this study, indicating that edible aquatic insects may be vulnerable to MP pollution. The whole bodies had more microplastic items than the other two samples because that is the structure where food and other ingested materials are deposited. In the case of contamination in the body without GT, microplastics could be retained in the exoskeleton of the insect body. Ingestion of MPs, on the other hand, is likely to have different effects on an organism depending on its size, shape, concentrations, and exposure time (Redondo-Hasselerharm et al., 2018).

Identification of microplastic polymers by FT-IR

A selected 52 plastic-like particles (about 18% of total MPs) were identified by a Fourier Transformed Infrared Spectroscope (FT-IR). Some particles were confirmed as polymethyl methacrylate (PMMA), polyethylene terephthalate (PET), and polypropylene (PP) (Table 2). For the 52 selected particles, 46.1% were identified as microplastics, 15.4% as non-microplastics, and 38.5% as unidentified particles. The spectral characteristics of these polymers are shown in Figs. 4A–4C.

The findings suggest evidence of detecting MPs in aquatic insects such as Pantala sp. (Odonata: Libellulidae) that humans eat from rice fields. Previously, there are limited research on plastic contamination in Thailand, particularly on aquatic insects in freshwater environments. Recent research (Windsor et al., 2019) revealed data on the occurrence of MP particles in aquatic macroinvertebrates in a riverine valley in South Wales, UK, with the presence of MPs in almost half of the samples (0.14 MPs/mg tissue). Ehlers, Manz & Koop (2019) conducted a similar study and discovered that MPs (e.g., polypropylene, polyethylene, and polyvinyl chloride) were present in the biological structure of freshwater organisms. Microplastics can also pass through mosquito life stages (i.e., larva, pupa, and adult) and spread throughout aquatic systems (Rhodes, 2019). Furthermore, ingestion of MP polymers was reported in Chironomus sp. (Diptera) (i.e., styrene ethylene butylene styrene, acrylonitrilebutadiene styrene (ABS), chlorinated polyethylene, polypropylene (PP), and polyester), Siphlonurus sp. (Ephemeroptera) (i.e., polyester and ABS) and Lestes viridis (Odonata) (i.e., polyester and PP) from Ogun and Osun Rivers, Nigeria (Akindele, Ehlers & Koop, 2020). As larger plastic debris breaks into smaller plastic bits, Cole et al. (2011) discovered that MPs were more likely to be fragments. Also, fibers, and rods were generated from original MPs. Secondary MPs make extrapolating results from single-species and virgin MP investigations challenging (Rummel et al., 2016). One of three ways that freshwater systems become contaminated by MPs are via effluent discharge, overflow of wastewater sewers during high rain events, and run-off from sludge, which all occur in rice fields (Eriksen et al., 2013). Storms and extreme weather conditions, according to can aggravate the flow of MPs from land to water (Cole et al., 2011).

In terms of trophic transfer and the potential for effects across multiple trophic levels, D’Souza et al. (2020) showed that plastics can be transported from invertebrate consumers to predators in natural freshwater ecosystems, including humans. Plastic is contaminating practically every area of the world and its ecosystems, which is a startling fact. However, ocean pollution and plastic deposition on marine animals and sea birds have received the most attention so far. As a result, it is past time for us to focus on freshwater sources as well.