Advertisement

Exposure to Cerium Dioxide Nanoparticles Differently Affect Swimming Performance and Survival in Two Daphnid Species

  • Ester Artells,

    Affiliations: Institut Méditerranéen de Biodiversité et d’Ecologie marine et continentale, IMBE UMR-CNRS 7263, Aix-Marseille Université, Marseille, France, Institut Méditerranéen de Biodiversité et d’Ecologie marine et continentale, IMBE UMR-CNRS 7263, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France

  • Julien Issartel,

    Affiliations: Institut Méditerranéen de Biodiversité et d’Ecologie marine et continentale, IMBE UMR-CNRS 7263, Aix-Marseille Université, Marseille, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Labex SERENADE 2012 “Safer Ecodesign Research and Education applied to NAnomaterial Development”, Aix-en-Provence, France

  • Mélanie Auffan,

    Affiliations: Centre Européen de Recherche et d’Enseignement des Géosciences de l’Environnement, CEREGE UMR-CNRS 7330, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Labex SERENADE 2012 “Safer Ecodesign Research and Education applied to NAnomaterial Development”, Aix-en-Provence, France

  • Daniel Borschneck,

    Affiliations: Centre Européen de Recherche et d’Enseignement des Géosciences de l’Environnement, CEREGE UMR-CNRS 7330, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France

  • Antoine Thill,

    Affiliations: iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Comissariat à l’énergie atomique et aux énergies alternatives, CEA Saclay, IRAMIS, UMR 3299, Laboratoire Interdisciplinaire sur l’Organisation Nanométrique et Supramoléculaire, Gif-sur-Yvette, France

  • Marie Tella,

    Affiliations: Centre Européen de Recherche et d’Enseignement des Géosciences de l’Environnement, CEREGE UMR-CNRS 7330, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Labex SERENADE 2012 “Safer Ecodesign Research and Education applied to NAnomaterial Development”, Aix-en-Provence, France

  • Lenka Brousset,

    Affiliations: Institut Méditerranéen de Biodiversité et d’Ecologie marine et continentale, IMBE UMR-CNRS 7263, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France

  • Jérôme Rose,

    Affiliations: Centre Européen de Recherche et d’Enseignement des Géosciences de l’Environnement, CEREGE UMR-CNRS 7330, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Labex SERENADE 2012 “Safer Ecodesign Research and Education applied to NAnomaterial Development”, Aix-en-Provence, France

  • Jean-Yves Bottero,

    Affiliations: Centre Européen de Recherche et d’Enseignement des Géosciences de l’Environnement, CEREGE UMR-CNRS 7330, Aix-Marseille Université, Aix-en-Provence, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Labex SERENADE 2012 “Safer Ecodesign Research and Education applied to NAnomaterial Development”, Aix-en-Provence, France

  • Alain Thiéry

    alain.thiery@imbe.fr

    Affiliations: Institut Méditerranéen de Biodiversité et d’Ecologie marine et continentale, IMBE UMR-CNRS 7263, Aix-Marseille Université, Marseille, France, iCEINT, International Consortium for the Environmental Implications of Nanotechnology, Aix-en-Provence, France, Labex SERENADE 2012 “Safer Ecodesign Research and Education applied to NAnomaterial Development”, Aix-en-Provence, France

Exposure to Cerium Dioxide Nanoparticles Differently Affect Swimming Performance and Survival in Two Daphnid Species

  • Ester Artells, 
  • Julien Issartel, 
  • Mélanie Auffan, 
  • Daniel Borschneck, 
  • Antoine Thill, 
  • Marie Tella, 
  • Lenka Brousset, 
  • Jérôme Rose, 
  • Jean-Yves Bottero, 
  • Alain Thiéry
PLOS
x
  • Published: August 15, 2013
  • DOI: 10.1371/journal.pone.0071260

Abstract

The CeO2 NPs are increasingly used in industry but the environmental release of these NPs and their subsequent behavior and biological effects are currently unclear. This study evaluates for the first time the effects of CeO2 NPs on the survival and the swimming performance of two cladoceran species, Daphnia similis and Daphnia pulex after 1, 10 and 100 mg.L−1 CeO2 exposures for 48 h. Acute toxicity bioassays were performed to determine EC50 of exposed daphnids. Video-recorded swimming behavior of both daphnids was used to measure swimming speeds after various exposures to aggregated CeO2 NPs. The acute ecotoxicity showed that D. similis is 350 times more sensitive to CeO2 NPs than D. pulex, showing 48-h EC50 of 0.26 mg.L−1 and 91.79 mg.L−1, respectively. Both species interacted with CeO2 NPs (adsorption), but much more strongly in the case of D. similis. Swimming velocities (SV) were differently and significantly affected by CeO2 NPs for both species. A 48-h exposure to 1 mg.L−1 induced a decrease of 30% and 40% of the SV in D. pulex and D. similis, respectively. However at higher concentrations, the SV of D. similis was more impacted (60% off for 10 mg.L−1 and 100 mg.L−1) than the one of D. pulex. These interspecific toxic effects of CeO2 NPs are explained by morphological variations such as the presence of reliefs on the cuticle and a longer distal spine in D. similis acting as traps for the CeO2 aggregates. In addition, D. similis has a mean SV double that of D. pulex and thus initially collides with twice more NPs aggregates. The ecotoxicological consequences on the behavior and physiology of a CeO2 NPs exposure in daphnids are discussed.

Introduction

To date, the effects of CeO2 nanoparticles (NPs) on aquatic and terrestrial environments are of growing concern since their production and uses are expected to rise in the future [1]. The CeO2 NPs are increasingly used in industry (as oxidation catalyst, gas sensor, polishing materials, UV absorber). These applications rely on the remarkable properties of Ce such as, its high affinity to oxygen, a potential redox chemistry involving Ce(III)/Ce(IV) and its unique adsorption/excitation energy bands [2]. However, the environmental release of these NPs, and subsequent behavior and biological effects are currently unclear. Consequently, since 2008 [3] CeO2 NPs have been included the OECD list of nanomaterials requesting immediate testing.

Understanding the toxic effects of these emerging xenobiotics is therefore crucial in order to anticipate the consequences of the potential degradation of ecosystems [4], [5] and their potential impact on health. The biotopes of aquatic organisms constitute the major sink for pollutants that accumulate the inputs from the surrounding hydrographic basins. Consequently, aquatic organisms, especially in the vicinity of urbanized areas, are generally considered as highly vulnerable. Studying the potential toxic effect of emerging xenobiotics of NPs on these vulnerable environments is a more than reasonable strategy.

Over the past few years, many studies have attempted to decipher the cellular toxic effects of NPs in aquatic organisms. It is now widely recognized that one of the major harmful aspects of these substances lies in the oxidative stress they induce [6]. Indeed, exposure of aquatic organisms to metallic NPs such as Fe-NPs [7] TiO2, CuO/Cu2O and Ag-NPs [8][11] as well as carbon nanomaterial such as fullerene [12], [13]; and also silica NPs [14] has been correlated to an increase in oxidative damages and to a modification of the antioxidant system [8][13]. In addition to oxidative stress markers, a large battery of other ecotoxicological endpoints has been monitored in aquatic organisms exposed to NPs. Among them, it was shown that some NPs induce the expression of varous defense cellular biomarkers such as heat shock proteins (e.g. in D. magna exposed to Cu2O NPs [9]), metallothioneins (e.g. in S. plana after a CuO NPs exposure [10]), or detoxification complexes such as CYP family isozymes (e.g. in Ag-NPs exposed medaka and C60-exposed fathead minnow [15], [16]). At a larger scale, some NPs can also induce histological abnormalities as observed in the medaka gills after exposure to Fe-NPs [7]. These fundamental sub-individual toxic effects are thought to be responsible for the time/concentration dependant-mortality observed after NPs exposure of aquatic animals. Although these case-by-case studies in highly controlled conditions are important to identify and understand the ecotoxicity mechanisms at the sub-individual scale (i.e. cellular and molecular levels), it is necessary to go further and to study the ecotoxicity of NPs at a larger biological scale. This will allow translating the toxic effects observed on sub-individual or individuals into relevant information to predict consequences at population levels. In this regards, modifications of behavior [5], [17] could be a good indicator. Indeed, behavioral parameters are accurate and reliable indicators since the behavior of an organism is the endpoint of a sequence of complex neurophysiological events (stimulation of neurons via the release of chemical messages, muscular contractions) [18][20]. Behavioral response could therefore be a very sensitive indicators of stress and very useful in obtaining a realistic picture of the effects of contaminants at the ecosystem level.

In aquatic organisms, swimming behavior responses to several environmental stimuli have been intensively investigated [21][23], especially in the case of permanently swimming zooplankters like daphnids. The swimming of these organisms is closely related to the energetic metabolism and to ecological parameters as food intake, predator escape and reproduction [24]. While the swimming of daphnids has frequently been used to test different substances such as, constituents of oral pill [25], natural cyanobacteria toxins produced by algal blooms [21], [26], [27], metals contaminants as cadmium [28], [29], copper [20], and organic xenobiotics as PCB, tributyltinchloride [30], cypermethrin [31], only few studies deal with nanoparticle effects. To our knowledge, only fullerene (nC60), TiO2 and Ag NPs were tested in relation to the swimming behavior of daphnids [32][36].

The present study is part of a series of tasks required to understand the impact of new nanotechnologies on the environment [37]. We propose to evaluate the CeO2 NPs impact on both the survival and swimming behavior of two daphnid species. To date, most of the ecotoxicity studies of NPs were performed with a single-species approach whereas a comparative multi-species approach provides a more complete and ecologically relevant overview of the impact of NPs in the ecosystem [9], [17], [32][36], [38], [39]. The Anomopod (Cladocera) Daphnia pulex (L., 1758) is an ecologically and genetically well-known organism [40], [41] and a good model to study multi-stressors in freshwater environments. For compaison with a closely related other species, the experiment was also conducted in Daphnia (Ctenodaphnia) similis (Claus, 1876), a water flea species present in temporary lakes in Provence (France).

Using an original experimental approach, our study revealed that both daphnids were differentially impacted by NPs exposure, bringing new information on the toxic effects of CeO2 NPs.

Materials and Methods

2.1. Nanoparticles Characterization

The CeO2 NPs were provided as a stable suspension at 130 g.L−1 of CeO2 by Rhodia Chemicals®. The size and crystalline structure of CeO2 NPs were determined using a Transmission Electron Microscope (TEM) JEOL® JEM 2010F URP22 equipped with an X-ray EDS-Kevex detector and an ELS-Gatan imaging filter. Samples (n = 60) were prepared by evaporating a droplet of a CeO2 NPs suspension on a carbon-coated copper grid at ambient temperature. The aggregation state of CeO2 NPs was characterized in the natural water (Cristaline®) used for daphnia cultures using the granulometer Malvern3000 (Malvern Instruments®, UK).

2.2. Organisms Breeding

Daphnia pulex (D. pulex) were collected from a permanent pond in the Paris countryside, the Forêt de Sordun, in the Seine and Marne Region, (48° 31′ 51″N, 3° 24′ 61″E, 175 m a.s.l.) and Daphnia (Ctenodaphnia) similis (D. similis) were collected, in January 2012, from a temporary pond, the Mare de Saint Maximin, in the Var Region, in Southern France (43° 26′ 16″N, 5° 52′ 19″E, 298 a.s.l.) in January 2012. No specific permissions were required for these locations. We confirm that the field studies did not involve endangered or protected species. Both species were acclimated and bred in the laboratory at 20±2°C with a natural photoperiod (10 h Light, 14 h Dark), and fed daily with the freshwater unicellular Chlorella vulgaris (Beijerinck, 1890) (AC149 strain, Algobank, France) at a concentration of 105–106 cells.mL−1. The breeding procedure was adapted from Barata [42]. The nutritive solution was the commercialized natural water (Cristaline®, France) (pH 8.5, 290 mg.L−1 HCO3, 5 mg.L−1 SO42−, 4 mg.L−1 Cl, 39 mg.L−1 Ca2+, 25 mg.L−1 Mg2+, 19 mg.L−1 Na+, 1.5 mg.L−1 K+).

2.3. Acute Toxicity Assay

The acute toxicity tests were conducted in accordance with OECD guideline number 202 [43], compatible with the procedure proposed by the US-EPA [44]. The concentrations used in this study are based on the EC50 from CeO2 exposed Daphnia magna [45]. The test medium was prepared from a 130 g.L−1 CeO2 NPs original stock solution diluted in miliQ water to obtain a final CeO2 NPs solution. To 2.5 ml of this final solution was then added to 47.5 ml of rearing Cristaline® water to obtain the experimental concentration used for the test. The bioassays were performed in septuplicate with five 8 days-old organisms. Eight days-old daphnids were chosen in order to minimize confounding effects of growth and reproduction energetic cost of younger and older stages, respectively [46]. Daphnids were placed into 50 mL of test medium and exposed for 96 h to 0, 0.1, 1, 10, 50 and 100 mg.L−1 CeO2 NPs. Immobility and mortality data were recorded each 24 h. The CeO2 NPs concentration in each chamber during toxicity test is considered constant as evaporation was negligible.

2.4. Swimming Velocity Assay

The effects of CeO2 NPs on D. pulex and D. similis swimming velocity were investigated. Both species were exposed to 0, 1, 10 and 100 mg.L−1 CeO2 NPs for 48 h in glass vials (45 mm diameter) containing 50 mL of solution. We used 3 replicates for each exposure conditions: each replicate consisted in at least 4 surviving daphnids in a vial. As both species were unable to move vertically at concentrations higher than 1 mg.L−1, only horizontal movements were measured. Before recording the daphnid movements, the volume of culture medium was slowly and carefully adjusted to 10 ml in order to limit vertical movement of daphnids. Daphnid movements were recorded using a Cam Sport® camera (China) EVO model operating at 25 frames.s−1 and high resolution 736×480 pixels; the camera was placed 15 cm above the swimming chamber. For each replicate and exposure concentration, 1 minute sequences were recorded and then transferred to a computer for analysis. Individual swimming velocities were calculated on the basis of a 10 seconds travel using ImageJ 1.46 and MTrackJ plugin, which allows calculating the distance traveled by the daphnid between two frames (i.e. 41.7 ms).

2.5. Micro-X-ray Fluorescence Analysis

The Ce spatial distribution in daphnids was determined with the XGT7000 X-ray analytical miscroscope (Horiba® Jobin Yvon) equipped with an X-ray tube producing a high-intensity beam with a 10 µm spot size (Rh X-ray source, 30 kV, 1 mA, equipped with an EDS detector). D. pulex and D. similis exposed to 10 mg.L−1 of CeO2 NPs for 48 h were analyzed using a Peltier freezing system to maintain the sample frozen during analysis. Given that the X-ray beam completely penetrates the sample, the obtained chemical images are 2D projections of a 3D sample. Elements from Na to U can be detected with a sensitivity range from about 50 mg.kg−1 to a few percent mass depending on the atomic number of the element and the nature of the matrix.

2.6. Statistical Analysis

The data obtained in these acute toxicity tests were used in order to determine the Median Effective Dose (EC50); this is done through Probit analyses using the statistical package SPSS (version 20, IBM®). For the swimming velocity statistical analysis, the normality of the data and the homogeneity of variances were verified using the Kolmogorov-Smirnov test and the Levene’s test, respectively. Differences between the mean swimming velocities of the control and the exposed groups were assessed using a one-way ANOVA. When significant differences were found, a Tukey post-hoc test was performed. Statistical analyses were performed using Statistica 6 (StatSoft Inc., Tulsa, USA). A 5% (p<0.05) significance was used in all tests.

Results

3.1. Nanoparticles Physico-chemical Behavior

By TEM, we observed well-crystallized clusters of cerianite (95–98% of purity) with a d-spacing (~3.2 Å) close to the d111 of CeO2 (dhkl). These clusters are pseudo-spherical with a diameter of 3±1 nm (n = 60) (Fig. 1). In pure water, these CeO2 NPs (100 mg.L−1) are colloidally stable with a negative zeta potential (−40±5 mV at pH 4) and an average hydrodynamic diameters of ~ 8 nm. Based on this value, the specific surface area of the CeO2 NPs was calculated to be about 110 m2.g−1.

thumbnail
Figure 1. Physico-chemical characterization of the CeO2 NPs.

TEM picture of the CeO2 in deionized water (A) and distribution of the hydrodynamic diameters within daphnia medium (B).

doi:10.1371/journal.pone.0071260.g001

The natural water (Cristalline®) used in the exposure scenario is at pH = 8.5 and elevated ionic strength. Once injected in the natural water, NPs aggregated due to the neutralization of the surface charges by the salts and the pH which is close to the isoelectric point (PIE) of our material. The PIE of these CeO2 NPs in water has previously been measured to be 7.5–8 [47]; their zeta potential measured in natural water is low, −10±2 mV (pH 8.5). Figure 1B shows the aggregate size distribution of a 100 mg.L−1 CeO2 NPs suspension in natural water measured 25 min. after NPs injection. Such a distribution of hydrodynamic diameters is not representative of the real size distribution of the NPs aggregates as the data treatment does not take into account the specific scattering properties of the NPs fractal aggregates. However, it clearly shows that CeO2 NPs form large aggregates with a maximum size larger than 300 µm.

3.2. Ecotoxicity Testing of CeO2 NPs Towards D. pulex and D. similis

The acute ecotoxicity study showed that D. similis was more sensitive to CeO2 NPs than D. pulex. For both D. pulex and D. similis, the toxic effects increased with increasing exposure duration. During the first 24 h, D. pulex was significantly more affected by CeO2 NPs than D. similis, but after 48 h an opposite trend occurred with D. similis displaying higher immobility and mortality values (Fig. 2). In the 100 mg.L−1 treatment, D. similis was more affected by CeO2 NPs than D. pulex during all test periode. The 48-h EC50 for D. similis were calculated to be 0.26 mg.L−1. For D. pulex, the 48-h EC50 (91.79 mg.L−1) obtained was 350 times higher than the 48-h EC50 of D. similis. After 72 h, surviving specimens were only observed for D. pulex in all of concentrations treatment while for D. similis in 0.1 mg.L−1 only few surviving specimens are found. This data is not sufficient to calculate the 72-h and 96-h EC50 of D. similis. The 72-h EC50 and 96-h EC50 for D. pulex were respectively 0.94 mg.L−1 and 0.78 mg.L−1.

thumbnail
Figure 2. Effect curve vs time of D. similis and D. pulex at 0.1 mg.L−1, 1 mg.L−1, 10 mg.L−1, 50 mg.L−1 and 100 mg.L−1 of CeO2 NPs.

Values are Mean EC50±SD.

doi:10.1371/journal.pone.0071260.g002

3.3. Relation Nanoparticules/Cuticle

D. similis and D. pulex present distinct morphologies. D. similis have a large distal spine (0.6–1 mm) and many small spines on the cuticle (Fig. 3C and D). On the opposite, D. pulex displays a short distal spine (0.10–0.25 mm) and only few spines on the cuticle (Fig. 3A and B). Using optical microscopy, we noticed that depending on their morphology, these daphnids were able to accumulate particles onto their shield following CeO2 NPs treatment. After a 48 h of exposure to 10 mg.L−1 of CeO2 NPs, D. similis accumulated a significant amount of particles onto the distal spine (Fig. 3D) and onto specific areas of the carapace (Fig. 3C), whereas no or only very slight accumulation was observed with D. pulex (Fig. 3A and B). This accumulation of particles formed a cloud just behind the distal spine when D. similis swam (Fig. 3E).

thumbnail
Figure 3. Representative image of distal spine (ds) and ventral margin of the shield (vms) in D. pulex and Daphnia similis exposed to 10 mg.L−1 of CeO2 NPs for 48 h.

Note the accumulation of particles onto the cuticle of D. similis. The optical image (E) represents the D. similis after 48 h exposure to 10 mg.L−1 of CeO2 NPs.

doi:10.1371/journal.pone.0071260.g003

Micro-XRF was used to identify the chemical composition of this cloud. Due to the presence of calcium and phosphorous, it is possible to observe the cuticle and the distal spine of daphnids on the Ca and P map (Fig. 4, Ca and P maps). Using the P map, we measured the length of the distal spine of D. similis to be 600 µm. This value was similar to the length measured by optical microscopy. After an incubation of 48 h, Ce was detected in a line just behind the distal spine in D. similis and on the surface in both species (Fig. 4). This CeO2 line is only visible in the case of D. similis and corresponds to the cloud observed using optical microscopy (Fig. 3).

thumbnail
Figure 4. Distribution of Ce (Lα line), P (Kα line) and Ca (Kα line) on the posterior region of D. pulex and D. similis exposed 48 h to CeO2 NPs.

Chemical map parameters: 128 pixel2 image, 1 pixel: 8 µm, total counting time 20000: sec, scale (white bar): 500 µm. Mean XRF spectra corresponding to specific area of the individual were generated from the hyperspectral map.

doi:10.1371/journal.pone.0071260.g004

3.4. Swimming Velocity

Due to the strong interactions between CeO2 NPs and the cuticle, we examined the ability of daphnids to swim in these contaminated exposure media. Figure 5 shows that the average swimming velocities (SV) were differently and significantly affected by CeO2 NPs for both species i.e. exposed daphnids swam slower than non-exposed daphnids of similar size. After 48-h exposure to 1 mg.L−1, a decrease of 30% and 40% of the SV is measured for D. pulex and D. similis, respectively. However at higher concentrations, the SV of D. similis was more impacted (60% off for 10 mg.L−1 and 100 mg.L−1) than the one of D. pulex. While the SV was significantly altered, no change of the hop frequency -i.e. number of downward thrusting of the second antennae below the helmet and then back above per minute- was observed in both species after a 48-h exposure to CeO2 NPs.

thumbnail
Figure 5. Mean swimming velocity in D. similis (A) and D. pulex (B) exposed to CeO2 NPs for 48 h.

Values are means ± SEM. Letters show significant differences established by one-way ANOVA and Tukey post-hoc (p<0.05). D. similis swimming tracks in control (C) and after a 48-h exposure to 10 mg.L−1 of CeO2 NPs (D).

doi:10.1371/journal.pone.0071260.g005

Discussion

4.1 NPs Aggregation Kinetics versus NPs/Daphnids Interaction Kinetics

The ~8 nm CeO2 NPs (hydrodynamic diameter) are introduced in a natural water at a pH close to their PIE and a ionic strength of 1.4 10−2 mol.L−1. In such physico-chemical conditions the repulsive electrostatic interactions which contribute to the colloidal stability of the CeO2 NPs are sufficiently reduced to trigger fast aggregation. Assuming a purely Brownian mechanism for the NPs collisions, it is possible to estimate the half life (t1/2) of fully destabilized NPs at a concentration of 100 mg.L−1 which depends on the temperature (T), viscosity () and initial NPs number concentration (C0) as:

This simple calculation shows that even if a significant residual stabilization is active, the NPs will aggregate very quickly. The size distribution represented on figure 1B after 25 min. is most probably reached at the very beginning of the experiment.

When the NPs interact with daphnids, the relevant collision mechanism is no longer the Brownian motion of the NPs. The active motion of the daphnids increases their collision rate with the NPs. A simple estimate of the ratio between the collision due to the Brownian motion of the NPs and those due the active swimming motion of the daphnids can be evaluated. First, Brownian collisions frequencies () involved between both the NPs and the daphnids can be written as , where is the radius of a NP and is the radius of a daphnid. As , the equation can be simplified to .

As to the collisions induced by the active motion of the daphnids, it is possible to assume that the motion of a daphnid is equivalent to a shear gradient (G) given by . Assuming this shear gradient, the collision frequency between the NPs and the daphnids reads as:

Using again the fact that rd>>rn, we have .

Using these simplified expressions, we have in the whole range of possible swimming velocities. The only important collision mechanism is thus the collisions induced by the swimming motion of the daphnids in the aggregated NPs suspension. As the size of the daphnids is the same for the two species, the difference in collision frequencies only depends on the differences of swimming velocities. Thus, we can conclude that initially D. similis collide with twice more aggregates than D. pulex.

4.2. Relation between Daphnia Morphology and the uptake of CeO2 NPs

Low levels of NPs adsorption to the exoskeleton of aquatic invertebrates has already been observed in a few previous studies (see e.g. D. magna exposed to nC60, TiO2 and Ag NPs [35], [36], [48] and Ceriodaphnia dubia exposed to Quantum Dots [49]). In a recent study, Gaiser et al. [50] observed a very slight adsorption of CeO2 NPs on D. magna neonates’ cuticles after 96 h of exposure to 10 mg.L−1. These different clinging capacities of CeO2 NPs may be due to their physico-chemical characteristics such as size, chemical nature, or surface coating [50]. The mechanisms of interaction between NPs and the cuticle are however not clear. In our case, D. pulex and D. similis display different accumulation of CeO2 NPs onto their cuticle. D. similis accumulates large aggregates whereas D. pulex is only slightly covered by small NPs or NPs aggregates. The objective of this section of the discussion is to understand the possible origin of these differences.

The interaction between the CeO2 NPs and the cuticle observed can be discussed in terms of both physico-chemical and mechanical processes. Indeed to accumulate on the cuticles of daphnids, NPs have first to undergo a collision with the cuticle; the frequency at which this occurs depends on various mechanical processes, as for example viscosity of the fluid, relative size of the aggregates and the daphnids and swimming velocities of the daphnids. Then, once on the surface of the daphnids, the NPs or the NPs aggregates can only accumulate if they adhere sufficiently strongly to resist the viscous strain induced by the daphnids active swimming motion.

A micro crustacean cuticle is mostly composed of a fibrous phase of crystalline chitin (nanofibrils with 3 nm of diameter), sugars, silk-like proteins attached through specific H-bonds, and globular proteins, which confer a net negative surface charge at neutral pH [51]. In our experimental conditions, a zeta potential of −10±2 mV was measured at the surface of the CeO2 NPs (at pH 8.5). This zeta potential value corresponds to a global negative charge which should generate a long distance repulsive potential between the NPs aggregates and the cuticle. At shorter distances, van der Waals attraction and possible surface complexation at specific CeO2 sites can be responsible for the NPs adhesion. Indeed, the surface of the CeO2 NPs being composed by a mixture of positive and negative sites, it is likely that mechanisms associating steric effects and surface complexation (with thiolated or carboxilated groups…) between the cuticle and the surface of CeO2 NPs contribute to the short distance adhesion. While, these physico-chemical interactions between CeO2 NPs and the cuticle (governed by van der Waals, steric effects and surface interaction) should be similar for both species, differences in morphology between D. similis and D. pulex are possibly responsible for different mechanical trapping of NPs or NPs aggregates. The ability to regain normal mobility after molting [39] has not been considered here as during our experiments the daphnids did not molt.

The main differences between the two daphnids species are the initial swimming velocity and the morphology of the cuticle surface. Due to its higher initial swimming velocity, the D. similis collide with NPs at an initial rate twice more important than the one of D. pulex. Moreover, the surface of D. similis is covered with several spines and has a long distal spine, while D. pulex has a short distal spine and very few spines on the cuticle. All the spines around the cuticle of D. similis and especially the distal spine generate reliefs that can act as traps for the CeO2 large NPs aggregates which dominate in the exposure media. These morphological differences may also modify the resistance of the trapped NPs aggregates against viscous strain due to the fluid motion. Furthermore, due to its smoothest surface, D. pulex will only retain the smaller aggregates.

Consequently, while D. similis is able to mechanically trap the dominating population of large NPs aggregates, D. pulex is only able to physico-chemically adsorb small aggregates. The proportion of these small agregates is not known quantitatively, but most probably it only represents a minor part of the aggregates population.

4.3. An Interspecific Sensitivity to NPs

In this study, the two different daphnids species present drastically different EC50. Interestingly, D. similis has a lower 24 h EC50 and a larger 48-h or more EC50 compared to D. pulex. D. similis also displays a large CeO2 adsorption/accumulation on its cuticle under the form of large aggregates and a high decrease of its SV. In contrast, D. pulex presents a high 24 h EC5O, a small CeO2 adsorption/accumulation under the form of smaller aggregates and a low decrease of the SV. The comparison with the EC50 values available for TiO2 NPs in the literature reveals strong interspecific survival differences in exposed daphnids (see Table 1). However, these different toxicities might be due to either different physico-chemical properties of TiO2 NPs or exposure conditions. In the current work, the same CeO2 NPs and exposure conditions were used for both species. Consequently, the different toxic effects of CeO2 NPs between D. similis and D. pulex reflect different sensibilities of each species. In daphnids, the toxicity of CeO2 NPs can be exerted via two ways: a mechanical toxicity by adsorption/accumulation of large NPs aggregates on the cuticle, and/or a metabolic toxicity by internalization of CeO2 NPs into the cells. In aquatic organisms, potential routes of internalization include entry across gills, olfactory organs or gut epithelium [14]. Although Auffan et al. (2013) showed that CeO2 NPs accumulate in the digestive tract of D. pulex [39], the metabolic toxicity of CeO2 NPs in daphnids is still unclear and, as far as we know, no direct evidence of internalization has been found in these organisms. However, in vitro studies on vertebrate cell cultures showed that the CeO2 NPs can penetrate into cells and induce oxidative stress [52], [53]. Further studies are needed to decipher the metabolic toxicity of CeO2 NPs in aquatic invertebrates.

thumbnail
Table 1. Median, maximal and minimal values of 48-h L(E)C50 of daphnids species tested with TiO2 NPs calculated from differents studies [38], [58][72].

doi:10.1371/journal.pone.0071260.t001

In our work, the higher sensibility at 48-h or more measured in D. similis can be explained by cumulative toxic effects: a mechanical toxic effect by adsorption/accumulation of large NPs aggregates due to its specific morphology and accompanied by a putative metabolic toxicity. In contrast, the lower sensibility showed by D. pulex can be explained by the metabolic toxicity alone as NPs only adsorb as small aggregates.

Consequently, we assume that the more important 48-h (or more) sensitivity of D. similis following CeO2 NPs exposure is due to the accumulation of aggregates that increase the drag force (decrease the swimming velocity). Large aggregates are however probably less efficient in inducing metabolic toxicity because these effects generally require a close proximity between the CeO2 NPs and the surface of the organism. This close proximity could explain the higher sensitivity at 24 h observed for D. pulex which only accumulates small aggregates close to the cuticle surface.

4.4. General Mechanistic Implications of CeO2 in Daphnia Physiological Functions

Among the different organism behavioral endpoints used to evaluate the risk associated to contaminants, the swimming performance of micro crustaceans is recognized particularly relevant, as this function is fundamentally correlated to numerous ecophysiological traits [33], [54]. The present work highlights that CeO2 NPs induce strong alteration of the daphnid swimming velocity related to the adsorption/accumulation of NPs onto the cuticle. Similar modifications of the swimming performance were observed in daphnids exposed to nC60, TiO2 and Ag NPs [32][35]. However, in these studies, no relationship between the NPs concentration and the alteration of the swimming behavior were measured/observed. Such concentration-response relationships were observed in studies dealing with the impact of dissolved metals and organic contaminants [20], [23], [25], [28][30]. To our knowledge, this work highlights for the first time the direct relationship existing between the decrease of the SV of daphnids and the existing concentration of NPs together with daphnid morphology effects.

Daphnids are filter feeders that are able to detect and migrate to food rich areas [55]. Thus a lower swimming capacity may directly impact their energy uptake and storage, and energetic metabolism. Our experiments showed that the hop frequency was not altered following exposure to NPs whereas the SV was dramatically decreased. This underlies that the daphnids attempt to maintain their swimming capacity but that the adsorption/accumulation of NPs onto their cuticles limit their movements through an increase of the viscous drag force. This might increase their energetic demand and lead to the organism death.

Another physiological parameter likely to be impacted by the decrease of the SV is the respiration rate. Daphnids generate a water current by swimming, this generates, through the carapace wall, gas exchange between the media and the haemolymph [56]. This water current also ensures a correct oxygenation of the eggs carried by mothers in their brood chambers [57]. An impaired capacity to swim decreases the water current, and consequently the O2 uptake by the organisms leading to anaerobiosis (i.e. a lower ATP supply).

All these sublethal effects related to swimming performance may impact survival capacities of the copepods exposed to CeO2 NPs.

Conclusions

This work investigates the acute toxicity of CeO2 NPs in two species of daphnids focusing on the survival capacities and unusual (eco)toxicity endpoint, the swimming behavior. We observed strong interspecific differences in survival, adsorption of the NPs on the cuticle and the swimming performance. This highlights how important it is to compare different species in order to thoroughly understand and anticipate the ecotoxicological effects of NPs in the environment. However, in addition to the mechanistic effect underlined in the present work, further studies should explore the metabolic toxicity of CeO2 NPs in both species, such as oxidative stress, and ionic regulation that seems to be sensitive to the morphology and surface proximity of the CeO2 aggregates.

Acknowledgments

The authors would like to thank the CNRS and the CEA for funding the iCEINT International Consortium for the Environmental implications of NanoTechnology. The authors thank the CP2M (Aix-Marseille university) for their help with the TEM analysis and Pr. William Stone for his assistance in correcting the draft of the manuscript.

Author Contributions

Conceived and designed the experiments: A. Thiéry EA. Performed the experiments: EA JI MA A. Thiéry DB. Analyzed the data: EA JI A. Thill MA. Contributed reagents/materials/analysis tools: MT LB. Wrote the paper: EA MA JI A. Thiéry. Reviewed the manuscript: EA JI MA DB A. Thill MT LB JR J-YB A. Thiéry.

References

  1. 1. Som C, Nowack B, Krug HF, Wick P (2012) Toward the development of decision supporting tools that can be used for safe production and use of nanomaterials. Acc Chem Res. DOI: 10.1021/ar3000458.
  2. 2. Lin W, Huang YW, Zhou XD, Ma Y (2006) Toxicity of cerium oxide nanoparticles in human lung cancer cells. Int J Toxicol 25: 451–457. doi: 10.1080/10915810600959543
  3. 3. Organisation for Economic Cooperation and Development (OECD) (2010) List of manufactured nanomaterials and list of endpoints for phase one of the sponsorrhip programme for the testing of manufactured nanomaterials: Revision. Series on the Safety of Manufactured Nanomaterials No. 27. Paris.
  4. 4. Auffan M, Santaella C, Thiéry A, Paillès C, Rose J, et al.. (2012) Ecotoxicity of inorganic nanoparticles: From unicellular organisms to invertebrates. In: Bhushan B, editor. Encyclopedia of Nanotechnology. 623–636.
  5. 5. Thiéry A, De Jong L, Issartel J, Moreau X, Saez G, et al. (2012) Effects of metallic and metal oxide nanoparticles in aquatic and terrestrial food chains. Biomarkers responses in invertebrates and bacteria. Inter J Nanotechnol 9: 181–203. doi: 10.1504/ijnt.2012.045326
  6. 6. Xia T, Kovochich M, Brant J, Hotze M, Sempf J, et al. (2006) Comparison of the abilities of ambient and manufactured nanoparticles to induce cellular toxicity according to an oxidative stress paradigm. Nano Lett 6: 1794–1807. doi: 10.1021/nl061025k
  7. 7. Li H, Zhou Q, Wu Y, Fu J, Wang T, et al. (2009) Effects of waterborne nano-iron on medaka (Oryzias latipes): antioxidant enzymatic activity, lipid peroxidation and histopathology. Ecotoxicol Environ Saf 72: 684–692. doi: 10.1016/j.ecoenv.2008.09.027
  8. 8. Wu Y, Zhou Q (2013) Silver nanoparticles cause oxidative damage and histological changes in medaka (Oryzias latipes) after 14 days of exposure. Environ Toxicol Chem 32: 165–173. doi: 10.1002/etc.2038
  9. 9. Fan W, Shi Z, Yang X, Cui M, Wang X, et al. (2012) Bioaccumulation and biomarker responses of cubic and octahedral Cu2O micro/nanocrystals in Daphnia magna. Water Res 46: 5981–5988. doi: 10.1016/j.watres.2012.08.019
  10. 10. Buffet PE, Tankoua OF, Pan JF, Berhanu D, Herrenknecht C, et al. (2011) Behavioural and biochemical responses of two marine invertebrates Scrobicularia plana and Hediste diversicolor to copper oxide nanoparticles. Chemosphere 84: 166–174. doi: 10.1016/j.chemosphere.2011.02.003
  11. 11. Federici G, Shaw BJ, Handy RD (2007) Toxicity of titanium dioxide nanoparticles to rainbow trout (Oncorhynchus mykiss): gill injury, oxidative stress, and other physiological effects. Aquat Toxicol 84: 415–430. doi: 10.1016/j.aquatox.2007.07.009
  12. 12. Ferreira JL, Barros DM, Geracitano LA, Fillmann G, Fossa CE, et al. (2012) In vitro exposure to fullerene C(60) influences redox state and lipid peroxidation in brain and gills from Cyprinus carpio (Cyprinidae). Environ Toxicol Chem 31: 961–967. doi: 10.1002/etc.1792
  13. 13. Klaine SJ, Alvarez PJ, Batley GE, Fernandes TF, Handy RD, et al. (2008) Nanomaterials in the environment: behavior, fate, bioavailability, and effects. Environ Toxicol Chem 27: 1825–1851. doi: 10.1897/08-090.1
  14. 14. Canesi L, Fabbri R, Gallo G, Vallotto D, Marcomini A, et al. (2010) Biomarkers in Mytilus galloprovincialis exposed to suspensions of selected nanoparticles (Nano carbon black, C60 fullerene, Nano-TiO2, Nano-SiO2). Aquat Toxicol 100: 168–177. doi: 10.1016/j.aquatox.2010.04.009
  15. 15. Zhu S, Oberdörster E, Haasch ML (2006) Toxicity of an engineered nanoparticle (fullerene, C60) in two aquatic species, Daphnia and fathead minnow. Mar Environ Res 62 Supplement 1S5–S9. doi: 10.1016/j.marenvres.2006.04.059
  16. 16. Chae YJ, Pham CH, Lee J, Bae E, Yi J, et al. (2009) Evaluation of the toxic impact of silver nanoparticles on Japanese medaka (Oryzias latipes). Aquat Toxicol 94: 320–327. doi: 10.1016/j.aquatox.2009.07.019
  17. 17. Gould P (2006) Zooplankton suffer under nanoparticle exposure: Toxicology. Nano Today 1: 19. doi: 10.1016/s1748-0132(06)70041-x
  18. 18. Lagadic L, Caquet T, Ramade F (1994) The role of biomarkers in environmental assessment (5). Invertebrate populations and communities. Ecotoxicology 3: 193–208. doi: 10.1007/bf00117084
  19. 19. Gerhardt A (1995) Monitoring behavioural responses to metals in Gammarus pulex (L.) (Crustacea) with impedance conversion. Environ Sci Pollut Res 2: 15–23. doi: 10.1007/bf02987506
  20. 20. Untersteiner H, Kahapka Jr, Kaiser H (2003) Behavioural response of the cladoceran Daphnia magna Straus to sublethal Copper stress–validation by image analysis. Aquat Toxicol 65: 435–442. doi: 10.1016/s0166-445x(03)00157-7
  21. 21. Hamza W, Ruggiu D (2000) Swimming behaviour of Daphnia galeata x hyalina as a response to algal substances and to opaque colours. Int Rev Hydrobiol 85: 157–166. doi: 10.1002/(sici)1522-2632(200004)85:2/3<157::aid-iroh157>3.3.co;2-p
  22. 22. Larsen PS, Madsen CV, Riisgard HU (2008) Effect of temperature and viscosity on swimming velocity of the copepod Acartia tonsa, brine shrimp Artemia salina and rotifer Brachionus plicatilis. Aquat Biol 4: 47–54. doi: 10.3354/ab00093
  23. 23. Garaventa F, Gambardella C, Di Fino A, Pittore M, Faimali M (2010) Swimming speed alteration of Artemia sp. and Brachionus plicatilis as a sub-lethal behavioural end-point for ecotoxicological surveys. Ecotoxicology 19: 512–519. doi: 10.1007/s10646-010-0461-8
  24. 24. Ebert D (2005) Introduction to Daphnia Biology. In: Bethesda (MD) National Center for Biotecnology, editor. Ecology, Epidemiology, and Evolution of Parasitism in Daphnia [Internet]. Avaiable from: http://www.ncbi.nlm.nih.gov/books/NBK203​6/. Accessed 2013 Mar 3.
  25. 25. Goto T, Hiromi J (2003) Toxicity of 17 a-ethynylestradiol and norethindrone, constituents of an oral contraceptive pill to the swimming and reproduction of cladoceran Daphnia magna, with special reference to their synergetic effect. Mar Pollut Bull 47: 139–142. doi: 10.1016/s0025-326x(03)00052-3
  26. 26. Haney JF, Sasner JJ, Ikawa M (1995) Effects of products released by Aphanizomenon flos-aquae and purified saxitoxin on the movements of Daphnia carinata feeding appendages. Limnol Oceanogr 40: 263–272. doi: 10.4319/lo.1995.40.2.0263
  27. 27. Ferrão-Filho AdS, Costa SMd, Ribeiro MGL, Azevedo SMFO (2008) Effects of a saxitoxin-producer strain of Cylindrospermopsis raciborskii (cyanobacteria) on the swimming movements of cladocerans. Environ Toxicol 23: 161–168. doi: 10.1002/tox.20320
  28. 28. Wolf G, Scheunders P, Selens M (1998) Evaluation of the swimming activity of Daphnia magna by image analysis after administration of sublethal Cadmium concentrations. Comp Biochem Physiol A 120: 99–105. doi: 10.1016/s1095-6433(98)10016-8
  29. 29. Baillieul M, Blust R (1999) Analysis of the swimming velocity of cadmium-stressed Daphnia magna. Aquat Toxicology 44: 245–254. doi: 10.1016/s0166-445x(98)00080-0
  30. 30. Schmidt K, Steinberg CEW, Staaks GBO (2005) Influence of a xenobiotic mixture (PCB and TBT) compared to single substances on swimming behavior or reproduction of Daphnia magna. Acta hydrochim hydrobiol 33: 287–300. doi: 10.1002/aheh.200400579
  31. 31. Christensen BT, Lauridsen TL, Ravn HW, Bayley M (2005) A comparison of feeding efficiency and swimming ability of Daphnia magna exposed to cypermethrin. Aquat Toxicol 73: 210–220. doi: 10.1016/j.aquatox.2005.03.011
  32. 32. Lovern SB, Strickler JR, Klaper R (2007) Behavioral and physiological changes in Daphnia magna when exposed to nanoparticle suspensions (Titanium dioxide, Nano-C60, and C60HxC70Hx). Environ Sci Technol 41: 4465–4470. doi: 10.1021/es062146p
  33. 33. Brausch KA, Anderson TA, Smith PN, Maul JD (2011) The effect of fullerenes and functionalized fullerenes on Daphnia magna phototaxis and swimming behavior. Environ Toxicol Chem 30: 878–884. doi: 10.1002/etc.442
  34. 34. Tao X, He Y, Zhang B, Chen Y, Hughes JB (2011) Effects of stable aqueous fullerene nanocrystal (nC60) on Daphnia magna: Evaluation of hop frequency and accumulations under different conditions. J Environ Sci 23: 322–329. doi: 10.1016/s1001-0742(10)60409-3
  35. 35. Asghari S, Johari SA, Lee JH, Kim YS, Jeon YB, et al. (2012) Toxicity of various silver nanoparticles compared to silver ions in Daphnia magna. J Nanobiotechnology 10: 14. doi: 10.1186/1477-3155-10-14
  36. 36. Dabrunz A, Duester L, Prasse C, Seitz F, Rosenfeldt R, et al. (2011) Biological surface coating and molting inhibition as mechanisms of TiO2 nanoparticle toxicity in Daphnia magna. PLoS One 6: e20112. doi: 10.1371/journal.pone.0020112
  37. 37. Saez G, Moreau X, De Jong L, Thiéry A, Dolain C, et al. (2010) Development of new nano-tools: Towards an integrative approach to address the societal question of nanotechnology? Nano Today 5: 251–253. doi: 10.1016/j.nantod.2010.06.002
  38. 38. Klaper R, Crago J, Barr J, Arndt D, Setyowati K, et al. (2009) Toxicity biomarker expression in daphnids exposed to manufactured nanoparticles: Changes in toxicity with functionalization. Environ Pollut 157: 1152–1156. doi: 10.1016/j.envpol.2008.11.010
  39. 39. Auffan M, Bertin D, Chaurand P, Paillès C, Dominici C, et al. (2013) Role of molting on the biodistribution of CeO2 nanoparticles within Daphnia pulex. Water Res. 47: 3921–3930. doi: 10.1016/j.watres.2012.11.063
  40. 40. Colbourne JK, Pfrender ME, Gilbert D, Thomas WK, Tucker A, et al. (2011) The ecoresponsive genome of Daphnia pulex. Science 331: 555–561. doi: 10.1126/science.1197761
  41. 41. Shaw J, Colbourne J, Davey J, Glaholt S, Hampton T, et al. (2007) Gene response profiles for Daphnia pulex exposed to the environmental stressor cadmium reveals novel crustacean metallothioneins. BMC Genomics 8: 477. doi: 10.1186/1471-2164-8-477
  42. 42. Barata C, Baird DJ (2000) Determining the ecotoxicological mode of action of chemicals from measurements made on individuals: results from instar-based tests with Daphnia magna Straus. Aquat Toxicol 48: 195–209. doi: 10.1016/s0166-445x(99)00038-7
  43. 43. Organisation for Economic Cooperation and Development (OECD) (2004) Guidelines for the testing of chemicals. Test No. 202: Daphnia sp. Acute immobilisation test. Avaiable: http://www.oecd-ilibrary.org/environment​/. Accessed 2013 Jan 14.
  44. 44. US-EPA (2002) Methods for measuring the acute toxicity of effluents and receiving waters to freshwater and marine organisms, 5th ed; Agency E-RUSEP, editor. Washington, DC.
  45. 45. Van Hoecke K, Quik JT, Mankiewicz-Boczek J, De Schamphelaere KA, Elsaesser A, et al. (2009) Fate and effects of CeO2 nanoparticles in aquatic ecotoxicity tests. Environ Sci Technol 43: 4537–4546. doi: 10.1021/es9002444
  46. 46. Arzate-Cárdenas MA, Martínez-Jerónimo F (2011) Age-altered susceptibility in hexavalent chromium-exposed Daphnia schodleri (Anomopoda: Daphniidae): integrated biomarker response implementation. Aquat Toxicol 105: 528–534. doi: 10.1016/j.aquatox.2011.08.006
  47. 47. Diot M-A (2012) Étude du vieillissement et de l’altération de nanocomposites de la vie courante. Aix-en-Provence: Aix-Marseille Université. 232 p.
  48. 48. Baun A, Sørensen SN, Rasmussen RF, Hartmann NB, Koch CB (2008) Toxicity and bioaccumulation of xenobiotic organic compounds in the presence of aqueous suspensions of aggregates of nano-C60. Aquat Toxicol 86: 379–387. doi: 10.1016/j.aquatox.2007.11.019
  49. 49. Ingle T, Alexander R, Bouldin J, Buchanan R (2008) Absorption of semiconductor nanocrystals by the aquatic invertebrate Ceriodaphnia dubia. Bull Environ Contam Toxicol 81: 249–252. doi: 10.1007/s00128-008-9481-y
  50. 50. Gaiser BK, Biswas A, Rosenkranz P, Jepson MA, Lead JR, et al. (2011) Effects of silver and cerium dioxide micro- and nano-sized particles on Daphnia magna. J Environ Monitoring 13: 1227–1235. doi: 10.1039/c1em10060b
  51. 51. Julian VFV (2002) Arthropod cuticle: a natural composite shell system. Composites Part A: App Sci Manufac 33: 1311–1315. doi: 10.1016/s1359-835x(02)00167-7
  52. 52. Xia T, Kovochich M, Liong M, Mädler L, Gilbert B, et al. (2008) Comparison of the mechanism of toxicity of zinc oxide and cerium oxide nanoparticles based on dissolution and oxidative stress properties. ACS Nano 2: 2121–2134. doi: 10.1021/nn800511k
  53. 53. Park EJ, Yi J, Chung KH, Ryu DY, Choi J, et al. (2008) Oxidative stress and apoptosis induced by titanium dioxide nanoparticles in cultured BEAS-2B cells. Toxicol Lett 180: 222–229. doi: 10.1016/j.toxlet.2008.06.869
  54. 54. Lagergren R, Lord H, Stenson JAE (2000) Influence of temperature on hydrodynamic costs of morphological defences in zooplankton: experiments on models of Eubosmina (Cladocera). Funct Ecol 14: 380–387. doi: 10.1046/j.1365-2435.2000.00433.x
  55. 55. Jensen KH, Larsson P, Högstedt G (2001) Detecting food search in Daphnia in the field. Limnol Oceanogr 46: 1013–1020. doi: 10.4319/lo.2001.46.5.1013
  56. 56. Pirow R, Wollinger F, Paul RJ (1999) The sites of respiratory gas exchange in the planktonic crustacean Daphnia magna: an in vivo study employing blood haemoglobin as an internal oxygen probe. J Exp Biol 202: 3089–3099.
  57. 57. Seidl MD, Pirow R, Paul RJ (2002) Water fleas (Daphnia magna) provide a separate ventilatory mechanism for their brood. Zoology 105: 15–23. doi: 10.1078/0944-2006-00050
  58. 58. Lovern SB, Klaper R (2006) Daphnia magna mortality when exposed to titanium dioxide and fullerene (C60) nanoparticles. Environ Toxicol Chem 25: 1132–1137. doi: 10.1897/05-278r.1
  59. 59. Hund-Rinke K, Simon M (2006) Ecotoxic effect of photocatalytic active nanoparticles (TiO2) on algae and daphnids. Environ Sci Pollut R 13: 225–232. doi: 10.1065/espr2006.06.311
  60. 60. Warheit DB, Hoke RA, Finlay C, Donner EM, Reed KL, et al. (2007) Development of a base set of toxicity tests using ultrafine TiO2 particles as a component of nanoparticle risk management. Toxicol Lett 171: 99–110. doi: 10.1016/j.toxlet.2007.04.008
  61. 61. Griffitt RJ, Luo J, Gao J, Bonzongo JC, Barber DS (2008) Effects of particle composition and species on toxicity of metallic nanomaterials in aquatic organisms. Environ Toxicol Chem 27: 1972–1978. doi: 10.1897/08-002.1
  62. 62. Heinlaan M, Ivask A, Blinova I, Dubourguier HC, Kahru A (2008) Toxicity of nanosized and bulk ZnO, CuO and TiO2 to bacteria Vibrio fischeri and crustaceans Daphnia magna and Thamnocephalus platyurus. Chemosphere 71: 1308–1316. doi: 10.1016/j.chemosphere.2007.11.047
  63. 63. Hall S, Bradley T, Moore JT, Kuykindall T, Minella L (2009) Acute and chronic toxicity of nano-scale TiO2 particles to freshwater fish, cladocerans, and green algae, and effects of organic and inorganic substrate on TiO2 toxicity. Nanotoxicology 3: 91–97. doi: 10.1080/17435390902788078
  64. 64. Lee S-W, Kim S-M, Choi J (2009) Genotoxicity and ecotoxicity assays using the freshwater crustacean Daphnia magna and the larva of the aquatic midge Chironomus riparius to screen the ecological risks of nanoparticle exposure. Environl Toxicol Phar 28: 86–91. doi: 10.1016/j.etap.2009.03.001
  65. 65. Wiench K, Wohlleben W, Hisgen V, Radke K, Salinas E, et al. (2009) Acute and chronic effects of nano- and non-nano-scale TiO2 and ZnO particles on mobility and reproduction of the freshwater invertebrate Daphnia magna. Chemosphere 76: 1356–1365. doi: 10.1016/j.chemosphere.2009.06.025
  66. 66. Zhu X, Zhu L, Chen Y, Tian S (2009) Acute toxicities of six manufactured nanomaterial suspensions to Daphnia magna. J Nanopar Res 11: 67–75. doi: 10.1007/s11051-008-9426-8
  67. 67. Zhu X, Chang Y, Chen Y (2010) Toxicity and bioaccumulation of TiO2 nanoparticle aggregates in Daphnia magna. Chemosphere 78: 209–215. doi: 10.1016/j.chemosphere.2009.11.013
  68. 68. Kim KT, Klaine SJ, Cho J, Kim SH, Kim SD (2010) Oxidative stress responses of Daphnia magna exposed to TiO2 nanoparticles according to size fraction. Sci Total Environ 408: 2268–2272. doi: 10.1016/j.scitotenv.2010.01.041
  69. 69. García A, Espinosa R, Delgado L, Casals E, González E, et al. (2011) Acute toxicity of cerium oxide, titanium oxide and iron oxide nanoparticles using standardized tests. Desalination 269: 136–141. doi: 10.1016/j.desal.2010.10.052
  70. 70. Menard A, Drobne D, Jemec A (2011) Ecotoxicity of nanosized TiO2. Review of in vivo data. Environ Pollut 159: 677–684. doi: 10.1016/j.envpol.2010.11.027
  71. 71. Amiano I, Olabarrieta J, Vitorica J, Zorita S (2012) Acute toxicity of nanosized TiO2 to Daphnia magna under UVA irradiation. Environ Toxicol Chem 31: 2564–2566. doi: 10.1002/etc.1981
  72. 72. Marcone GP, Oliveira AC, Almeida G, Umbuzeiro GA, Jardim WF (2012) Ecotoxicity of TiO2 to Daphnia similis under irradiation. J Hazard Mater 211–212: 436–442. doi: 10.1016/j.jhazmat.2011.12.075