Next Article in Journal
Mercury in Soil and Forage Plants from Artisanal and Small-Scale Gold Mining in the Bombana Area, Indonesia
Next Article in Special Issue
‘The Plastic Nile’: First Evidence of Microplastic Contamination in Fish from the Nile River (Cairo, Egypt)
Previous Article in Journal
Evaluation of Existing Models to Estimate Sorption Coefficients for Ionisable Pharmaceuticals in Soils and Sludge
Previous Article in Special Issue
Enchytraeus crypticus Avoid Soil Spiked with Microplastic
 
 
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Communication

Microplastics Exposure Causes Negligible Effects on the Oxidative Response Enzymes Glutathione Reductase and Peroxidase in the Oligochaete Tubifex tubifex

1
Faculty of Biological and Environmental Sciences Ecosystems and Environment Research Programme, University of Helsinki, Niemenkatu 73, Lahti FI-15140, Finland
2
Korea Institute of Science & Technology (KIST Europe) Environmental Safety Group. Joint Laboratory of Applied Ecotoxicology Campus E 7.1 66123 Saarbrücken, Germany
3
Helsinki Institute of Sustainability (HELSUS), Fabianinkatu 33, 00014 Helsinki, Finland
4
Department of Chemistry “Ugo Schiff”, University of Florence, Sesto Fiorentino, 50019 Florence, Italy
5
Department of Chemistry “Ugo Schiff”, University of Florence, and Consorzio Interuniversitario per lo Sviluppo dei Sistemi a Grande Interfase (CSGI), Sesto Fiorentino, 50019 Florence, Italy
*
Author to whom correspondence should be addressed.
Submission received: 23 January 2020 / Revised: 10 February 2020 / Accepted: 12 February 2020 / Published: 15 February 2020
(This article belongs to the Special Issue Prevalence, Fate and Effects of Plastic in Freshwater Environments)

Abstract

:
Microplastics (MPs) are emerging pollutants, which are considered ubiquitous in aquatic ecosystems. The effects of MPs on aquatic biota are still poorly understood, and consequently, there is a need to understand the impacts that MPs may pose to organisms. In the present study, Tubifex tubifex, a freshwater oligochaete commonly used as a bioindicator of the aquatic environment, was exposed to fluorescent polyethylene microspheres (up to 10 µm in size) to test whether the oxidative stress status was affected. The mortality rate of T. tubifex, as well as the activities of the oxidative stress status biomarker enzymes glutathione reductase and peroxidase, were assessed. In terms of oxidative stress, no significant differences between the exposure organisms and the corresponding controls were detected. Even though the data suggest that polyethylene MPs and the selected concentrations did not pose a critical risk to T. tubifex, the previously reported tolerance of T. tubifex to environmental pollution should be taken into account and thus MPs as aquatic pollutants could still represent a threat to more sensitive oligochetes.

1. Introduction

Plastic pollution is one of the primary environmental concerns we are facing today [1]. Microplastics (MPs), typically referred to as pieces smaller than 5 mm in any dimension [2], have been found on beaches, in oceans, seas, rivers, and lakes [2,3,4,5,6,7,8]. One of the biggest concerns regarding MP pollution is that marine and freshwater biota can mistake MP particles for food. MPs can be ingested by benthic and pelagic organisms belonging to different trophic levels [9], including mussels [10], lugworms [11], amphipods [12], zooplankton [13], and fish [14]. Some studies showed that with ingestion possible internal damages and blockages [9,15] may occur. Ingested MPs can act as vectors for transferring chemicals, additives, and other persistent organic compounds (such as Polybrominated diphenyl ethers (PBDEs) and Polychlorinated biphenyls (PCBs)) adsorbed from surrounding waters to biota [16,17,18,19]. However, the effects of MPs on exposed biota have not been extensively investigated, and the physiological effects remain poorly understood. Thus, there is a current need to gather data to deepen our understanding of their impacts. A few studies have demonstrated that MPs could induce oxidative stress in the organisms able to ingest them [20,21]. Browne et al. [11] studied the effect of MPs on the oxidative status of lugworms (Arenicola marina) demonstrating that animals exposed to polyvinyl chloride (PVC) microparticles were more susceptible to oxidative damage by up to 30%. Jeong et al. [20] showed that in response to microplastic-induced reactive oxygen species (ROS), antioxidant enzymes such as glutathione reductase (GR), glutathione peroxidase (GPx), glutathione-S-transferase (GST), and superoxide dismutase (SOD) were activated in the monogonont rotifer (Brachionus koreanus). The authors exposed the monogonont rotifer to three different polystyrene MP sizes (0.05, 0.5, and 6 µm) and observed that the toxicity was size-dependent, and the smaller the particles, the more toxic. More recently, Lu et al. [21] showed that the freshwater fish Danio rerio (zebrafish) exhibited a higher oxidative stress status after seven days of exposure to polystyrene MP, revealed by increased SOD and catalase (CAT) activities. Deng et al. [22] tested the effect of fluorescent polystyrene MP in mice, revealing increased activities for SOD and GPx and decreased CAT activity, signifying the potential health risk MPs represent to biota. Nevertheless, only a few of these studies report on the physiological effects of MP exposure in biota from aquatic environments, where MP abundance is most often reported [3,4,5,6,7,8].
The present study, therefore, investigated the toxic effects of MPs in Tubifex tubifex, which inhabits sediments of lakes and rivers. T. tubifex (also referred to as sludge or sewerage worm) lives in the uppermost sediment layers of these freshwater systems usually feeding in the sediment fraction smaller than 63 µm [23]. T. tubifex plays a key role in the decomposition of organic matter and bioturbation [24], dwelling in the sediment with the anterior part and keeping the tail outside, undulating it in the water to enable cutaneous respiration [6]. It is considered to be a model organism to perform sediment toxicity experiments [25,26] due to its high pollution tolerance [27] and is thus often one of the last species to disappear from a polluted habitat [24,28]. Tubifex worms are known to accumulate heavy metals [29,30,31] and organic pollutants [32] and are able to ingest MPs under natural conditions [33].
We, therefore, selected T. tubifex as a test subject to investigate the possible effects of MP pollution it may be exposed to in its environment. The aim of the present study was to understand whether exposure to MPs could affect the oxidative stress status of T. tubifex. Tubifex worms were exposed to fluorescent polyethylene (PE) MP particles (10 µm diameter) to evaluate the survival of the worms and the potential oxidative stress induced by MPs. PE was chosen because it is one of the most abundant polymers identified among samples collected from aqueous environments [7]. The size was selected within the size range of particles the worms could consume.
The enzyme activity of the oxidative stress biomarker enzymes GR and peroxidase (POD) were measured to check for alterations. Both enzymes, GR and POD, are known to be able to counteract the damages caused by ROS species to cells [34,35,36,37] and are thus induced in response to an increased oxidative stress status. GR plays a vital role in the antioxidant system [37,38], acting as a reductant in oxidation-reduction processes catalyzing the reduction of glutathione disulfide (GSSG) to glutathione (GSH) using NADPH as a cofactor. POD is known for playing a pivotal role in preventing H2O2 causing damage to DNA, proteins, and cell membranes [34,37] and has often been used in monitoring stress induced by contaminants [35,36].

2. Materials and Methods

T. tubifex was cultured in 1 L beakers in a synthetic medium at a constant temperature of 20 ± 1 °C and low light (18 μmol photons/m2s) with a photoperiod of 16 h light to 8 h dark (permanent continuous culture at the University of Helsinki) for several weeks before experimentation. The synthetic medium (artificial freshwater) was exchanged every three to five days, and it consisted of de-ionized water, CaCl2 [240 μg/L], KCl [6 μg/L], MgSO4.7H2O [123 μg/L], and NaHCO3 [55 μg/L]. The T. tubifex worms were fed with dry fish food (Sera, Mikropan, analytical constituents: crude protein 47.6%, crude fat 8.7%, crude fiber 3.3%, moisture 6.0%, crude ash 11.6%) daily.

2.1. Exposure to Microplastics

To establish the mortality of the worms in response to MP exposure, the T. tubifex worms were exposed to polyethylene MP particles (up to 10 µm in diameter) in four treatments; i.e., (1) T. tubifex in artificial freshwater containing 2 g/L MP (w/v) without sediment, (2) T. tubifex in artificial freshwater and sediment with the sediment containing 2 mg/g MP (w/w), (3) T. tubifex in artificial freshwater and sediment with the media spiked with 2 g/L MP (w/v), and (4) T. tubifex in artificial freshwater and sediment with both the media (2 g/L w/v MP) and the sediment (2 mg/g w/w MP) containing MP. The surviving worms were counted after 24 h, 48 h, and 120 h. For each experiment (mortality and enzyme assays, for all treatments), negative controls without MPs were conducted in replicates of five in parallel.
In order to examine the effects on the oxidative stress status, T. tubifex worms in sediment and artificial freshwater were exposed to the polyethylene MP particles (up to 10 µm in diameter) for 24 h in two different scenarios, i.e., MP-contaminated soil (2 mg/g w/w MP) vs. MP-contaminated media (2 g/L w/v MP). After 24 h, the treated and control worms were removed from the glass beakers and washed three times each with artificial freshwater, and the surviving worms counted.
In each experimental setup, five replicates of six adult T. tubifex worms per replicate were used. Each independent replicate was set up in a glass beaker containing either contaminated non-sterile sediments from Vesijärvi Lake (Lahti, Finland) or contaminated artificial freshwater or both as specified above.

2.2. Oxidative Stress Status: Enzyme Assays

After rinsing the worms with artificial freshwater, the enzymes were extracted using a shortened protocol modified from Pflugmacher [39]. T. tubifex worms were homogenized in 20 mM sodium phosphate buffer (pH 7.0) and stirred for 20 min followed by centrifugation at 9.000× g for 20 min. The supernatant (S-9 fraction) was collected and the enzyme activities were assayed immediately.
The activity of peroxidase (POD, EC 1.11.1.7) was analyzed spectrophotometrically using guaiacol as substrate [40], which is oxidized in the presence of hydrogen peroxide (H2O2) to octahydrotetraguaiacol. Changes in color as absorbance were measured at 436 nm over 3 min at 30 °C.
Glutathione reductase (GR, EC 1.8.1.7) activity was analyzed according to Schaedle and Bassham [41] following the glutathione disulfide-dependent NADPH oxidation at 340 nm for 3 min.
Each spectrophotometric enzyme assay was performed in triplicate per independent replicate (15 readings) using an Infinite 200 Pro plate reader (Tecan). The enzyme activity was normalized against protein content, which was determined according to Bradford [42]. Enzymatic activities are reported in nkat/mg protein, where 1 kat is the conversion rate of 1 mol of substrate per sec.
During all steps of the laboratory work, the possibility of self-contamination was taken into consideration and therefore fleece clothing or other personal items which could serve as a source of MP contamination were avoided [43].

2.3. Data Analysis

Statistical analysis was performed using IBM SPSS Statistics version 25 (2017). All data were checked for normality and homogeneity by Shapiro-Wilks and Levene’s test. A normal distribution of the data was analyzed using one-way analysis of variance (ANOVA) followed by Tukey’s test. For not normally distributed data, the Kruskal-Wallis test was employed to identify differences amongst the treatments, followed by the Mann-Whitney test if necessary. To assess whether survival/mortality changed with time, a repeated-measures ANOVA was performed. The results were considered significant at an alpha value of 0.05. All values are reported as mean ± standard error.

3. Results

3.1. Mortality

Contamination of the artificial water, sediment, or both water and sediment with MP (Figure 1) did not significantly affect the survival of T. tubifex in comparison to controls with time (p > 0.05). With contaminated artificial water and non-contaminated sediment, 95% ± 5% of the worms survived and with both sediment and water contaminated 90% ± 10% survived after five days. For all other treatments, 100% survived.

3.2. Oxidative Stress Status

Exposure of T. tubifex to fluorescence PE microspheres did not result in significant (p > 0.05) changes in the GR activity in comparison to the controls (Figure 2a). The mean GR activities in the worms with MP contaminated water (S + (L + MP)) and contaminated sediment (L + (S + MP)) were 0.005 ± 0.001 and 0.006 ± 0.001 nkat/mg protein, respectively, compared to the control GR activity of 0.005 ± 0.001 nkat/mg protein.
Peroxidase activity with exposure of T. tubifex to fluorescence PE microspheres, as observed for GR, did not result in significant (p > 0.05) changes in POD activity in comparison to controls (Figure 2b). Mean POD activities with the water (S + (L + MP)) and sediment (L + (S + MP)) contaminated respectively, were 0.120 ± 0.002 nkat/mg protein and 0.146 ± 0.002 nkat/mg protein, respectively, and the control POD activity was 0.130 ± 0.003 nkat/mg protein.

4. Discussion

Oxidative stress has been investigated intensively in many organisms [44,45,46] and it has been shown to cause damages to DNA, lipids, and proteins, potentially leading to an alteration in vital functions [47,48,49,50]. However, studies regarding oxidative stress and the alterations in antioxidant enzyme activities due to exposure to MPs are still limited [20,21].
In this study, T. tubifex was exposed to fluorescent PE microspheres to evaluate the potential damages caused by MPs as percentage mortality or survival and on a physiological level as induced oxidative stress. Mortality percentage did not show any difference between treatment and control samples. These results are in accordance with Redondo-Hasselerharm et al. [51], where no effects on the mortality rate of five freshwater benthic macroinvertebrates (including a Tubifex sp.) were found after exposure to polystyrene MPs for 28 days.
No significant differences were evident in terms of GR activity of T. tubifex exposed to MP via contaminated water and sediment. Similarly, POD activity did not differ significantly in comparison to controls in both of the experiments. These data show that the selected MP concentrations are not particularly critical for T. tubifex and they do not cause oxidative stress to the worms. Nevertheless, our results do not imply that MPs are not representing a threat to the biodiversity of aquatic environments. Contrary to our findings, Jeong at al., [20] and Lu et al., [21] showed that exposure to PS MPs caused oxidative stress in the monogonont rotifer (Brachionus koreanus) and Zebrafish (Danio rerio), respectively. However, we have to take into consideration the different targeted organisms and the type and size of MPs used in the studies. Either the chosen test subjects may have been more susceptible than T. tubifex to MPs exposure, or the selection of a different polymer could explain the discrepancy between the results. Even though exposure to PE MPs seems not to increase the production of ROS in T. tubifex, it does not mean that MP occurrence is not able to negatively affect freshwater environments. T. tubifex is considered to be a pollution-tolerant species [24,28], and the effects of MPs could pose a major risk to more sensitive organisms [11,20,21]. Furthermore, the concentration of MPs is expected to increase worldwide [52] representing a possible threat to the biodiversity of marine and freshwater environments. Further research is required to understand the effects in more sensitive organisms. In this study, no estimate on how many PE MPs were ingested has been made; a follow-up study should be performed repeating the tests changing the level of MP exposure and performing fluorescence microscopy analyses to examine the MP ingestion rate.

Author Contributions

Conventionalization and experimental design: S.P., C.S., M.E.; Analysis: C.S., M.E.; Manuscript preparation: C.S., M.E., A.C. All authors have read and agreed to the published version of the manuscript.

Funding

Open access funding provided by University of Helsinki including Helsinki University Central Hospital.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Brandon, J.A.; Jones, W.; Ohman, M.D. Multidecadal increase in plastic particles in coastal ocean sediments. Sci. Adv. 2019, 5. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Martellini, T.; Guerranti, C.; Scopetani, C.; Ugolini, A.; Chelazzi, D.; Cincinelli, A. A snapshot of microplastics in the coastal areas of the Mediterranean Sea. Trends Anal. Chem. 2018, 109, 173–179. [Google Scholar] [CrossRef]
  3. Cole, M.; Lindeque, P.; Halsband, C.; Galloway, T.S. Microplastics as contaminants in the marine environment: A review. Mar. Pollut. Bull. 2011, 62, 2588–2597. [Google Scholar] [CrossRef]
  4. Cincinelli, A.; Martellini, T.; Guerranti, C.; Scopetani, C.; Chelazzi, D. A potpourri of microplastics in the sea surface and water column of the Mediterranean Sea. Trends Anal. Chem. 2018, 110, 321–326. [Google Scholar] [CrossRef]
  5. Cincinelli, A.; Scopetani, C.; Chelazzi, D.; Lombardini, E.; Martellini, T.; Katsoyiannis, A.; Fossi, M.C.; Corsolini, S. Microplastic in the surface waters of the Ross Sea (Antarctica): Occurrence, distribution and characterization by FTIR. Chemosphere 2017, 175, 391–400. [Google Scholar] [CrossRef]
  6. Liu, T.; Diao, J.; Di, S.; Zhou, Z. Bioaccumulation of isocarbophos enantiomers from laboratory-contaminated aquatic environment by tubificid worms. Chemosphere 2015, 124, 77–82. [Google Scholar] [CrossRef]
  7. Klein, S.; Dimzon, I.K.; Eubeler, J.; Knepper, T.P. Analysis, occurrence, and degradation of microplastics in the aqueous environment. In Freshwater Microplastics; Wagner, M., Lambert, S., Eds.; Springer: Heidelberg, Germany, 2018; Volume 58, pp. 51–67. [Google Scholar]
  8. Scopetani, C.; Chelazzi, D.; Cincinelli, A.; Esterhuizen-Londt, M. Assessment of microplastic pollution: Occurrence and characterisation in Vesijärvi lake and Pikku Vesijärvi pond, Finland. Environ. Monit. Assess 2019, 191, 652. [Google Scholar] [CrossRef] [Green Version]
  9. Eerkes-Medrano, D.; Thompson, R.C.; Aldridge, D.C. Microplastics in freshwater systems: A review of the emerging threats, identification of knowledge gaps and prioritisation of research needs. Water Res. 2015, 75, 63–82. [Google Scholar] [CrossRef]
  10. Farrell, P.; Nelson, K. Trophic level transfer of microplastic: Mytilus edulis (L.) to Carcinus maenas (L.). Environ. Pollut 2013, 177, 1–3. [Google Scholar] [CrossRef] [PubMed]
  11. Browne, M.A.; Niven, S.J.; Galloway, T.S.; Rowland, S.J.; Thompson, R.C. Microplastic moves pollutants and additives to worms, reducing functions linked to health and biodiversity. Curr. Biol. 2013, 23, 2388–2392. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  12. Ugolini, A.; Ungherese, G.; Ciofini, M.; Lapucci, A.; Camaiti, M. Microplastic debris in sandhoppers. Estuar. Coast. Shelf Sci. 2013, 129, 19–22. [Google Scholar] [CrossRef]
  13. Cole, M.; Lindeque, P.; Fileman, E.; Halsband, C.; Goodhead, R.; Moger, J.; Galloway, T.S. Microplastic ingestion by zooplankton. Environ. Sci. Technol. 2013, 47, 6646–6655. [Google Scholar] [CrossRef] [PubMed]
  14. Lusher, A.L.; McHugh, M.; Thompson, R.C. Occurrence of microplastics in the gastrointestinal track of pelagic and demersal fish from the English Channel. Mar. Pollut. Bull. 2013, 67, 94–99. [Google Scholar] [CrossRef] [PubMed]
  15. Wright, S.L.; Thompson, R.C.; Galloway, T.S. The physical impacts of microplastics on marine organisms: A review. Environ. Pollut. 2013, 178, 483–492. [Google Scholar] [CrossRef] [PubMed]
  16. Mato, Y.; Isobe, T.; Takada, H.; Kanehiro, H.; Ohtake, C.; Kaminuma, T. Plastic resin pellets as a transport medium for toxic chemicals in the marine environment. Environ. Sci. Technol 2001, 35, 318–324. [Google Scholar] [CrossRef]
  17. Ogata, Y.; Takada, H.; Mizukawa, K.; Hirai, H.; Iwasa, S.; Endo, S.; Mato, Y.; Saha, M.; Okuda, K.; Nakashima, A.; et al. International pellet watch: Global monitoring of Persistent Organic Pollutants (POPs) in coastal waters. 1. Initial phase data on PCBs, DDTs, and HCHs. Mar. Pollut. Bull 2009, 58, 1437–1446. [Google Scholar] [CrossRef] [PubMed]
  18. Lithner, D.; Larsson, A.; Dave, G. Environmental and health hazard ranking and assessment of plastic polymers based on chemical composition. Sci. Total Environ. 2011, 409, 3309–3324. [Google Scholar] [CrossRef]
  19. Scopetani, C.; Cincinelli, A.; Martellini, T.; Lombardini, E.; Ciofini, A.; Fortunati, A.; Pasquali, V.; Ciattini, S.; Ugolini, A. Ingested microplastic as a two-way transporter for PBDEs in Talitrus saltator. Environ. Res. 2018, 167, 411–417. [Google Scholar] [CrossRef]
  20. Jeong, C.B.; Won, E.J.; Kang, H.M.; Lee, M.C.; Hwang, D.S.; Hwang, U.K.; Zhou, B.; Souissi, S.; Lee, S.J.; Lee, J.S. Microplastic size-dependent toxicity, oxidative stress induction, and p-JNK and p-p38 activation in the Monogonont rotifer (Brachionus koreanus). Environ. Sci. Technol. 2016, 50, 8849–8857. [Google Scholar] [CrossRef]
  21. Lu, Y.; Zhang, Y.; Deng, Y.; Jiang, W.; Zhao, Y.; Geng, J.; Ding, L.; Ren, H. Uptake and accumulation of polystyrene microplastics in zebra fish (Danio rerio) and toxic effects in liver. Environ. Sci. Technol. 2016, 50, 4054–4060. [Google Scholar] [CrossRef]
  22. Deng, Y.; Zhang, Y.; Lemos, B.; Ren, H. Tissue accumulation of microplastics in mice and biomarker responses suggest widespread health risks of exposure. Sci. Rep. 2017, 7, 46687. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. Rodriguez, P.; Martinez-Madrid, M.; Arrate, J.A.; Navarro, E. Selective feeding by the aquatic oligochaete Tubifex tubifex (Tubificidae, Clitellata). Hydrobiologia 2001, 463, 133–140. [Google Scholar] [CrossRef]
  24. Mosleh, Y.Y.; Paris-Palacios, S.; Biagianti-Risbourg, S. Metallothioneins induction and antioxidative response in aquatic worms Tubifex tubifex (Oligochaeta, tubificiadae) exposed to copper. Chemosphere 2005, 64, 121–128. [Google Scholar] [CrossRef] [PubMed]
  25. Chapman, P.M. Utility and relevance of aquatic oligochaetes in ecological risk assessment. In Aquatic Oligochaete Biology VIII; Rodriguez, P., Verdonschot, P.F.M., Eds.; Springer: Dordrecht, The Netherlands, 2001; Volume 158, pp. 149–169. [Google Scholar]
  26. Chapman, K.K.; Benton, M.J.; Brinkhurst, R.O.; Scheuerman, P.R. Use of the aquatic oligochaetes Lumbriculus variegatus and Tubifex tubifex for assessing the toxicity of copper and cadmium in a spiked-artificial-sediment toxicity test. Environ. Toxicol. 1999, 14, 271–278. [Google Scholar] [CrossRef]
  27. Wiederholm, T.; Dave, G. Toxicity of metal polluted sediments to Daphnia magna and Tubifex tubifex. Hydrobiologia 1989, 176–177, 411–417. [Google Scholar] [CrossRef]
  28. Lucan-Bouché, M.L.; Biagianti-Risbourg, S.; Arsac, F.; Vernet, G. An original decontamination process developed by the aquatic oligochaete Tubifex tubifex exposed to copper and lead. Aquat. Toxicol. 1999, 45, 9–17. [Google Scholar] [CrossRef]
  29. Bouché, M.L.; Habets, F.; Biagianti-Risbourg, S.; Vernet, G. Toxic effects and bioaccumulation of cadmium in the aquatic oligochaete Tubifex tubifex. Ecotoxicol. Environ. Saf. 2000, 46, 246–251. [Google Scholar] [CrossRef]
  30. Hare, L.; Tessier, A.; Warren, L. Cadmium accumulation by invertebrates living at the sediment-water interface. Environ. Toxicol. Chem. 2001, 20, 880–889. [Google Scholar]
  31. Méndez-Fernández, L.; Martínez-Madrid, M.; Rodriguez, P. Toxicity and critical body residues of Cd, Cu and Cr in the aquatic oligochaete Tubifex tubifex (Müller) based on lethal and sublethal effects. Ecotoxicology 2013, 22, 1445–1460. [Google Scholar] [CrossRef]
  32. Egeler, P.; Römbke, J.; Meller, M.; Knacker, T.; Franke, C.; Studinger, G.; Nagel, R. Bioaccumulation of lindane and hexachlorobenzene by tubificid sludgeworms (Oligochaeta) under standardised laboratory conditions. Chemosphere 1997, 35, 835–852. [Google Scholar] [CrossRef]
  33. Hurley, R.R.; Woodward, J.C.; Rothwell, J.J. Ingestion of microplastics by freshwater tubifex worms. Environ. Sci. Technol. 2017, 51, 12844–12851. [Google Scholar] [CrossRef] [PubMed]
  34. Di Giulio, R.T.; Washburn, P.C.; Wenning, R.J.; Winston, G.W.; Jewell, C.S. Biochemical response in aquatic animals: A review of determinants of oxidative stress. Environ. Toxicol. Chem. 1989, 8, 1103–1123. [Google Scholar] [CrossRef]
  35. Scalet, M.; Federico, R.; Guido, M.; Manes, F. Peroxidase activity and polyamine changes in response to ozone and simulated acid rain in Aleppo pine needles. Environ. Exp. Bot. 1995, 35, 417–425. [Google Scholar] [CrossRef]
  36. Mitrovic, S.M.; Pflugmacher, S.; James, K.J.; Furey, A. Anatoxin-a elicits an increase in peroxidase and glutathione S-transferase activity in aquatic plants. Aquat. Toxicol. 2004, 68, 185–192. [Google Scholar] [CrossRef] [PubMed]
  37. Monferran, M.V.; Wunderlin, D.A.; Nimptsch, J.; Pflugmacher, S. Biotransformation and antioxidant response in Ceratophyllum demersum experimentally exposed to 1,2- and 1,4-dichlorobenzene. Chemosphere 2007, 68, 2073–2079. [Google Scholar] [CrossRef]
  38. Mishra, S.; Srivastava, S.; Tripathi, R.D.; Kumar, R.; Seth, C.S.; Grupta, D.K. Lead detoxification by coontail (Ceratophyllum demersum L.) involves induction of phytochelatins and antioxidant system in response to its accumulation. Chemosphere 2006, 65, 1027–1039. [Google Scholar] [CrossRef]
  39. Pflugmacher, S. Promotion of oxidative stress in the aquatic macrophyte Ceratophyllum demersum during biotransformation of the cyanobacterial toxin microcystin-LR. Aquat. Toxicol. 2004, 70, 169–178. [Google Scholar] [CrossRef]
  40. Drotar, A.; Phelps, P.; Fall, R. Evidence for glutathione peroxidase activities in cultured plant cells. Plant Sci. 1985, 42, 35–40. [Google Scholar] [CrossRef]
  41. Schaedle, M.; Bassham, J.A. Chloroplast glutathione reductase. Plant Physiol. 1977, 59, 1011–1012. [Google Scholar] [CrossRef] [Green Version]
  42. Bradford, M.M. A rapid and sensitive method for the quantification of microgram quantities of proteins utilising the principal of protein-dye binding. Anal. Biochem. 1976, 72, 248–254. [Google Scholar] [CrossRef]
  43. Scopetani, C.; Esterhuizen-Londt, M.; Chelazzi, D.; Cincinelli, A.; Setälä, H.; Pflugmacher, S. Self-contamination from clothing in microplastics research. Ecotoxicol. Environ. Saf. 2020, 189, 110036. [Google Scholar] [CrossRef] [PubMed]
  44. Cochón, A.C.; Della Penna, A.B.; Kristoff, G.; Piol, M.N.; San Martín de Viale, L.C.; Verrengia Guerrero, N.R. Differential effects of paraquat on oxidative stress parameters and polyamine levels in two freshwater invertebrates. Ecotoxicol. Environ. Saf. 2007, 68, 286–292. [Google Scholar] [CrossRef] [PubMed]
  45. Antunes, S.C.; Freitas, R.; Figueira, E.; Gonçalves, F.; Nunes, B. Biochemical effects of acetaminophen in aquatic species: Edible clams Venerupis decussate and Venerupis philippinarum. Environ. Sci. Pollut. Res. 2013, 20, 6658–6666. [Google Scholar] [CrossRef] [PubMed]
  46. Esterhuizen-Londt, M.; Schwartz, K.; Pflugmacher, S. Using aquatic fungi for pharmaceutical bioremediation: Uptake of acetaminophen by Mucor hiemalis does not result in an enzymatic oxidative stress response. Fungal Biol. 2016, 120, 1249–1257. [Google Scholar] [CrossRef]
  47. Rikans, L.E.; Hornbrook, K.R. Lipid peroxidation, antioxidant protection and aging. Biochim. Biophys. Acta 1997, 1362, 116–127. [Google Scholar] [CrossRef] [Green Version]
  48. Halliwell, B.; Gutteridge, J.M.C. Free Radicals in Biology and Medicine, 5th ed.; Oxford University Press: Oxford, UK, 1999; p. 905. [Google Scholar]
  49. Barata, C.; Varo, I.; Navarro, J.C.; Arun, S.; Port, C. Antioxidant enzyme activities and lipid peroxidation in the freshwater cladoceran Daphnia magna exposed to redox cycling compounds. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 2005, 140, 175–186. [Google Scholar] [CrossRef]
  50. Esterhuizen-Londt, M.; Wiegand, C.; Downing, T.G. ß-N-Methylamino-L-alanine (BMAA) uptake by the animal model, Daphnia magna and consequent oxidative stress. Toxicon 2015, 100, 20–26. [Google Scholar] [CrossRef]
  51. Redondo-Hasselerharm, P.E.; Falahudin, D.; Peeters, E.T.H.M.; Koelmans, A.A. Microplastic effect thresholds for freshwater benthic macroinvertebrates. Environ. Sci. Technol. 2018, 52, 2278–2286. [Google Scholar] [CrossRef]
  52. Barboza, L.G.A.; Gimenez, B.C.G. Microplastics in the marine environment: Current trends and future perspectives. Mar. Pollut. Bull. 2015, 97, 5–12. [Google Scholar] [CrossRef]
Figure 1. Tubifex tubifex mortality after exposure to microplastics (MPs) in media (L + MP (2 g/L)) only, in sediment and media with the media contaminated with MPs (S + (L + MP 2g/L), in sediment and media with the sediment contaminated (L + (S + MP 2 mg/g)), and sediment and media both contaminated with MPs ((S + MP 2 mg/g) + (L + MP 2 g/L)).
Figure 1. Tubifex tubifex mortality after exposure to microplastics (MPs) in media (L + MP (2 g/L)) only, in sediment and media with the media contaminated with MPs (S + (L + MP 2g/L), in sediment and media with the sediment contaminated (L + (S + MP 2 mg/g)), and sediment and media both contaminated with MPs ((S + MP 2 mg/g) + (L + MP 2 g/L)).
Toxics 08 00014 g001
Figure 2. Peroxidase (a) and glutathione reductase (b) activities in T. tubifex exposed to fluorescent polyethylene (PE) microspheres through the media (S + (L + MP)) and the sediment (L + (S + MP). Values are expressed as mean enzyme activity ± standard error (n = 5). * denotes significance compared to the control (p < 0.05).
Figure 2. Peroxidase (a) and glutathione reductase (b) activities in T. tubifex exposed to fluorescent polyethylene (PE) microspheres through the media (S + (L + MP)) and the sediment (L + (S + MP). Values are expressed as mean enzyme activity ± standard error (n = 5). * denotes significance compared to the control (p < 0.05).
Toxics 08 00014 g002

Share and Cite

MDPI and ACS Style

Scopetani, C.; Esterhuizen, M.; Cincinelli, A.; Pflugmacher, S. Microplastics Exposure Causes Negligible Effects on the Oxidative Response Enzymes Glutathione Reductase and Peroxidase in the Oligochaete Tubifex tubifex. Toxics 2020, 8, 14. https://0-doi-org.brum.beds.ac.uk/10.3390/toxics8010014

AMA Style

Scopetani C, Esterhuizen M, Cincinelli A, Pflugmacher S. Microplastics Exposure Causes Negligible Effects on the Oxidative Response Enzymes Glutathione Reductase and Peroxidase in the Oligochaete Tubifex tubifex. Toxics. 2020; 8(1):14. https://0-doi-org.brum.beds.ac.uk/10.3390/toxics8010014

Chicago/Turabian Style

Scopetani, Costanza, Maranda Esterhuizen, Alessandra Cincinelli, and Stephan Pflugmacher. 2020. "Microplastics Exposure Causes Negligible Effects on the Oxidative Response Enzymes Glutathione Reductase and Peroxidase in the Oligochaete Tubifex tubifex" Toxics 8, no. 1: 14. https://0-doi-org.brum.beds.ac.uk/10.3390/toxics8010014

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop