HUNGARIAN JOURNAL OF INDUSTRY AND CHEMISTRY Vol. 47(2) pp. 71–75 (2019) hjic.mk.uni-pannon.hu DOI: 10.33927/hjic-2019-22 ASSESSMENT OF THE ECOTOXICITY OF NANOPLASTICS NÓRA KOVÁTS*1 , BETTINA ECK-VARANKA1 , ZSÓFIA BÉKÉSSY1 , DORINA DIÓSI1 , KATALIN HUBAI1 , AND JÁNOS KORPONAI2 1Institute of Environmental Sciences, University of Pannonia, 8200 Veszprém, Egyetem u. 10, HUNGARY 2Department of Environmental Science, Sapientia Hungarian University of Transylvania, 400193 Cluj-Napoca, Calea Turzii 4, ROMANIA The presence of micro- and nanoplastics in aquatic environments (including freshwater and marine ecosystems as well as their sediments) is becoming an increasingly serious problem worldwide. A wide range of studies have addressed the ecological effects these particles pose on biota. The main exposure pathway are food chains, e.g. under laboratory conditions these particles accumulate in the brain tissues of fish that feed on zooplankton causing brain damage. These studies, however, report mainly on the physical effects. In order to establish actual ecotoxicological effects, nanoplastics (50 nm in diameter) were assessed using the Vibrio fischeri bioluminescence inhibition bioassay (VFBIA). Our results showed that even environmentally relevant concentrations might trigger ecotoxicological effects. This study can be con- sidered to be a first screening, however, results indicate the need for more complex testing on a battery of aquatic test organisms. Keywords: nanoplastic; ecotoxicity; Vibrio fischeri ; kinetic assay 1. Introduction Aquatic environments contaminated by plastic litter are an emerging problem. Remote, pristine mountainous ar- eas are even contaminated by atmospheric microplastic deposition [1]. Polymer particles < 5 mm in diameter are defined as microplastics (MP) and may be derived directly from the use of industrial pellets or indirectly from the degradation and fragmentation of plastic parti- cles [2]. Polystyrene was proven to degrade into micro- and nanoplastics under laboratory conditions [3]. High levels of contamination have been reported in both ma- rine and freshwater habitats [4, 5]. Micro- and nanoplas- tics (NP) can float freely in bodies of water or be de- posited as sediments. The highest risk associated with these particles is their ingestion, which occurs at different levels in the aquatic food chain. Jabeen et al. [6], for example, listed approximately 150 different fish species where ingestion and accumulation have been reported. Particles can also progress upwards in the trophic levels of the food chain, i.e. fish can be exposed to the ingestion of zooplankton which is not able to discriminate between different food sources and consumes micro- and nanoplastics [7]. An experimental study showed that in fish exposed to NPs via the food chain, these particles caused brain damage and behavioural disorders as a result of accumulation in *Correspondence: kovats@almos.uni-pannon.hu brain tissues [8]. Biomagnification may also affect food safety and human health, though certain knowledge gaps exist in this field [9]. Ingestion may actually lead to starvation and even- tually the impairment of their physical condition. Un- der laboratory conditions, Daphnia magna exposed to polystyrene nanoparticles (PS-NP) exhibited reduced body size and severe alterations in terms of reproduc- tion [10]. D. magna is a widely studied species due to its key role in the aquatic food chain. It was shown to ingest nano- and microplastics (20 nm to 70 µm in diam- eter) from water [11]. In a laboratory study by Mattsson et al. [8], particles 52 nm in diameter elucidated the most severe effects. Cui et al. [12] exposed D. galeata to PS- NPs (5 mg/l, 52 nm in diameter) and detected a significant mortality rate after 2 days of exposure until the end of the study which lasted 5 days. Although a standard ecotoxi- cological test was conducted in this case, the mechanisms of mortality are still unclear: physical contact might have led to a reduction in the survival rate. In general, most ecotoxicological studies have used relatively high concentrations. Manfra et al. [13] investi- gated the impact of green fluorescently labeled carboxy- lated polystyrene nanoparticles of 40 nm in diameter with various surface charges on the marine rotifer Brachionus plicatilis. It was found that anionic PS-NPs did not elu- cidate mortality within the range of concentrations tested (0−50 µg/ml), while cationic PS-NPs caused mortality at https://doi.org/10.33927/hjic-2019-22 mailto:kovats@almos.uni-pannon.hu 72 KOVÁTS, ECK-VARANKA, BÉKÉSSY, DIÓSI, HUBAI, AND KORPONAI concentrations ≥ 2.5 µg/ml. Changes in oxidative stress enzymes were detected within the concentration range of 10−20 µg/ml in different organisms, e.g. the rotifer Bra- chionus koreanus and the marine copepod Paracyclop- ina nana [14]. The same concentration, 10 µg/ml, was reported to cause 40 % growth inhibition in the green mi- croalga Dunaliella tertiolecta [15]. In order to distinguish real (eco)toxicological effects from physical damage, a test based on the biolumines- cence inhibition of the marine bacterium Vibrio fischeri was selected. The species has been reclassified as Ali- ivibrio fischeri [16], however, as most standards and even recent papers from the literature still use the name V. fis- cheri, it will be used hereinafter. Bioluminescence is regulated by the enzyme system NAD(P)H:FMN oxidoreductase-luciferase. In toxic en- vironments, enzyme inhibition is reflected by a rapid de- crease in the luminous emittance of the bacterium. The reduction in light intensity is easy to measure as it is pro- portional to the strength of the toxicant, therefore, pro- vides a quantifiable endpoint. This test has been used in various environmental matrices [17–19]. Lappalainen et al. [20,21] developed a special version of the test which was later standardised (ISO 21338:2010: Water quality - Kinetic determination of the inhibitory ef- fects of sediment, other solids and coloured samples on the light emission of Vibrio fischeri (kinetic luminescent bacteria test)) in which bacteria are kept in suspension in direct contact with potentially toxic solid particles. Lumi- nescence readings were taken when the test commenced and the light intensity continuously monitored over the first 30 secs after the sample had been mixed with the bacteria. The light output pattern, therefore, might al- ready provide some indication of the expected toxicity of the sample [22]. The light intensity was measured once more after the pre-set exposure time (5, 15 or 30 mins as per standard). Toxicity values are normally expressed as EC50 and EC20, i.e. concentrations causing lumines- cence inhibitions of 50 and 20 % in this assay, respec- tively. 2. Materials and Methods In our experiments, the Ascent luminometer (Flash sys- tem, marketed by Aboatox, Finland) was used. A suspen- sion of the test bacteria (NRRL B-11177) was prepared in accordance with manufacturer instructions (Hach Lange GmbH). Polystyrene particles with a nominal diameter of 50 nm were used as a sample (supplier Thermo Fisher Sci- entific). As no comparative data were available on the potentially toxic concentration, a range-finding concen- tration series was set [23]. Three initial sample concen- trations were selected (1 g/l, 1 µg/l and 1 ng/l), which were further diluted, the number of dilutions was 11 (the number of concentrations the 96-multiwell plate permits) and the dilution ratio 1 : 2. Table 1: The measured EC20 values of the polystyrene nanoparticles. Concentration 1 g/l 1 µg/l 1 ng/l EC20 5.2 17.31 30.51 The Vibrio fischeri strain NRRL B-11177 was recon- stituted by adding the contents of one vial of +4 ◦C 1243- 551 Reagent Diluent. The reconstituted reagent was equi- librated at +4 ◦C for 30 min. Then the reagent was sta- bilised at +15 ◦C for 30 mins before being pipetted into the wells. Luminescence readings were taken when the test commenced, Time0, and after the pre-set exposure time of 30 mins, Time30. The luminescence inhibition of each sample was calculated as follows: CF = IC30/IC0 INH \% = 100 $-$ 100 x (IT30 / CF x IT0) where CF = correction factor IC30 = luminescence intensity of the control sample after the contact time (30 mins) in the RLU IC0 = initial luminescence intensity of the control sample in the RLU IT30 = luminescence intensity of the test sample after the contact time (30 mins) in the RLU IT0 = initial luminescence intensity of the test sample in the RLU EC20 values were calculated using the Ascent software, also developed by Aboatox Oy. 3. Results and Discussion Table 1 shows the ecotoxicity expressed in EC20, i.e. the calculated concentration of the sample that caused 20 % bioluminescence inhibition. Fig. 1 illustrates the biolumi- nescence inhibition during the first 30 secs for the sam- ples of 1 g/l and 1 ng/l in concentration. EC20 (or in some cases, EC10) are considered thresholds for the estimation of the lowest observed effec- tive concentration [24], i.e. the sample is normally con- sidered (eco)toxic if the elucidated effect exceeds 20 %. These results show that the V. fischeri bioassay de- tected a measurable degree of toxicity even at a concen- tration of 1 ng/l. Booth et al. [25] used the non-kinetic version of this bioassay (Microtox®), however, in their study, the calculated toxic concentration exceeded the range of concentrations studied (0.001−1000 mg/l). The same negative effect was reported by Casado et al. [26]. The higher degree of (detectable) toxicity in our study might be explained by the differences in the test system used. While Microtox® is a non-kinetic test, the Flash system (Ascent luminometer) was especially developed to test the toxicity of different suspensions or samples containing solid particles. The Ascent luminometer uses a 96-multiwell microplate. A specific feature of it is that Hungarian Journal of Industry and Chemistry ASSESSMENT OF THE ECOTOXICITY OF NANOPLASTICS 73 (a) (b) Figure 1: Kinetic diagram of the 1 g/l (a) and 1 ng/l (b) samples. The light output is recorded over the first 30 sec- onds. After the peak, toxicity causes a rapid reduction in the light output, on the other hand, it remains constant dur- ing the control. The two columns show the two replicates. E1-F1/G1-H1 (left): control. E2-F2/G2-H2 (right): sam- ple, maximum concentration. during luminescence readings, the microplate is continu- ously shaken by the instrument, resulting in the resuspen- sion of particles. According to our results, environmentally relevant concentrations might already pose ecotoxic effects. Ac- tual environmental concentrations are relatively difficult to compare and assess, mostly due to difficulties in sam- pling and the lack of standardized sampling methodolo- gies [27,28]. Indicative data are available: e.g. microplas- tic concentrations of 0.4 − 34 ng/l in bodies of freshwa- ter in the USA [29] or 0.51 mg/l in marine environments [10]. However, in real-world environments, even higher levels of toxicity can be expected as particles might absorb organic pollutants from the surrounding water [30], including highly toxic pesticides or polychlorinated biphenyls (PCBs) [31]. Though their bioavailability is still questionable [32], Batel et al. [33] conducted a lab- oratory study on microplastics and one polycyclic aro- matic hydrocarbon (PAH), benzo[a]pyrene (BaP). It was demonstrated that BaP adsorbed on microplastics and was transferred via an artificial food chain. These par- ticles might also possess inherent toxicity due to the use of additives during manufacturing processes [34]. 4. Conclusions It is a well-known paradigm in ecotoxicology that the sensitivity of different test organisms to a particular chemical varies, therefore, the V. fischeri test can be re- garded as a first screening. The bioluminescence inhibi- tion assay is an acute test that uses a maximum expo- sure of only 30 minutes. Naturally, chronic effects can- not be extrapolated from these results. However, the fact that the tested nanoplastics have already elucidated eco- toxicological effects in environmentally relevant concen- trations emphasises the need for more complex ecotoxi- cological testing involving a properly selected battery of test organisms. In addition to widely used aquatic test or- ganisms such as the aforementioned Daphnia magna, an ideal candidate could be the Caenorhabditis elegans test. It is a standardised bioassay using a sediment-dwelling, widely distributed nematode. 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