Ecotoxicology and Environmental Safety 113 (2015) 23–30
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Evaluation of zinc oxide nanoparticles toxicity on marine algae chlorella vulgaris through flow cytometric, cytotoxicity and oxidative stress analysis T.Y. Suman a, S.R. Radhika Rajasree a,n, R. Kirubagaran b a b
Center for Ocean Research (NIOT-SU Collaborative Research Centre), Sathyabama University, Chennai 600119, Tamil Nadu, India Marine Biotechnology, ESOO-National Institute of Ocean Technology, OSTI, Pallikaranai, Chennai 600100, India
art ic l e i nf o
a b s t r a c t
Article history: Received 26 July 2014 Received in revised form 14 November 2014 Accepted 19 November 2014
The increasing industrial use of nanomaterials during the last decades poses a potential threat to the environment and in particular to organisms living in the aquatic environment. In the present study, the toxicity of zinc oxide nanoparticles (ZnO NPs) was investigated in Marine algae Chlorella vulgaris (C. vulgaris). High zinc dissociation from ZnONPs, releasing ionic zinc in seawater, is a potential route for zinc assimilation and ZnONPs toxicity. To examine the mechanism of toxicity, C. vulgaris were treated with 50 mg/L, 100 mg/L, 200 mg/L and 300 mg/L ZnO NPs for 24 h and 72 h. The detailed cytotoxicity assay showed a substantial reduction in the viability dependent on dose and exposure. Further, flow cytometry revealed the significant reduction in C. vulgaris viable cells to higher ZnO NPs. Significant reductions in LDH level were noted for ZnO NPs at 300 mg/L concentration. The activity of antioxidant enzyme superoxide dismutase (SOD) significantly increased in the C. vulgaris exposed to 200 mg/L and 300 mg/L ZnO NPs. The content of non-enzymatic antioxidant glutathione (GSH) significantly decreased in the groups with a ZnO NPs concentration of higher than 100 mg/L. The level of lipid peroxidation (LPO) was found to increase as the ZnO NPs dose increased. The FT-IR analyses suggested surface chemical interaction between nanoparticles and algal cells. The substantial morphological changes and cell wall damage were confirmed through microscopic analyses (FESEM and CM). & 2014 Elsevier Inc. All rights reserved.
Keywords: Chlorella vulgaris Cytotoxicity Flow cytometry Oxidative stress ZnO NPs
1. Introduction Nanotechnology has come to the forefront of research in the past decade. With the advent of this technology, wide varieties of nanoparticles (NPs) with a variety of unique characteristics are manufactured and are used for a broad range of applications (Zhao and Castranova., 2011). Metal oxide NPs are among the most used engineered NPs in various commercial products, leading to concerns of their potential toxicity to human and environmental health (Aschbergeretal., 2011). Increasing use of metaloxide NPs is likely to result in the release of these particles into the aquatic environment. Adverse effects of NPs on the aquatic environment and organisms recently have drawn much special attention (Blaise etal., 2008; Farre´ et al., 2009). Among metal oxide nanomaterials, zinc oxide nanoparticles (ZnO NPs) are noted for their chemical stability and strong adsorption ability and have been extensively used in commercial n
Corresponding author. Fax: þ 44 24501270. E-mail address: [email protected] (S.R. Radhika Rajasree).
http://dx.doi.org/10.1016/j.ecoenv.2014.11.015 0147-6513/& 2014 Elsevier Inc. All rights reserved.
products like sunscreens, coatings, and paints (Osmond and McCall., 2010). ZnO has a high inherent risk of water contamination, and can reach high concentrations in surface waters posing significant threat to aquatic ecosystems (Gottschalk et al., 2009). Previous studies have demonstrated that ZnO NPs are toxic to microorganisms, cells, plants, aquatic biota and rodents (Brayner et al., 2006; Lin and Xing., 2007; Premanathan et al., 2011; Reddy et al., 2007; Wang et al., 2008a; Zhu et al., 2009). Zhu et al. (2008) found that ZnO NPs had higher acute toxicity to zebrafish embryos than nano-TiO2 and nano-Al2O3. The toxicity of ZnO NPs (96hLC50, 4.9 mg/L)to zebra fish was also much higher than that of nano-TiO2 (96 hLC50, 124.5 mg/L) (Xiong et al., 2011). Increased production of antioxidant enzymes in organisms is regarded as an early warning indicator of pollution in the environment (Song et al., 2008). Antioxidant enzyme activity can protect cells from the adverse effects of reactive oxygen species (ROS). Superoxide dismutase (SOD), catalase (CAT), and peroxidase (POD) enzymes help to check cellular levels of ROS (Weckx and Clijsters, 1996). Studies of the biotoxicity of ZnO NPs suggest several mechanisms of action of NPs, one of which is their ability to release free zinc ions which
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can synergistically enhance the production of ROS and cause oxidative damage in cells. Overproduction of ROS is believed to be a major mechanism of the toxicity of NPs (Sharifi et al., 2012). During this process, chemical reactions occur and result in increased formation of the superoxide radical (O2 ), which leads to ROS accumulation and oxidative stress (De Berardis et al., 2010). It has been reported that ZnONPs disturb the balance between oxidation and anti-oxidation processes and cause oxygen stress responses (Hao and Chen, 2012). As primary producers, phytoplankton plays a key role in aquatic ecosystems. Microalgae, being sensitive to pollutants, are excellent aquatic models and being widely prevalent in lakes and seas is easy to culture and propagate (Chen et al., 2012). The evaluation of NP effects upon marine phytoplankton is a necessary step to predict their potential impact on coastal marine food webs and on the whole ecosystems they support (Manzo et al., 2013). ZnO NPs, on entering the aquatic environment, release free metal ions in solution state. The dissolution speed of the particles mainly depends on the particle size, surface area, rough degree, etc. ZnO is slightly soluble, and can release zinc ions into the solution. Some researchers consider that dissolved zinc ions play an important role in the toxicity of ZnO NPs (Song et al., 2010). Some studies report that the toxicity of ZnO NPs was closely linked with its dissolved free ions (Franklin et al., 2007; Heinlaan et al., 2008; Wienchet al., 2009), but others have also shown that the toxicity of ZnO NPs was much higher than its ion toxicity (Nair et al., 2009; Wong et al., 2010). Both Escherichia coli and Pseudomonas fluorescens exhibited a high tolerance to zinc ions at the tested ZnO concentration (20 mg/L). The attachment of ZnO NPs to the surface of bacteria was the main cause of the ZnO toxicity of these bacterial species (Jiang et al., 2009).The point is that, at current knowledge, the observed ZnO toxicity cannot be attributed univocally to the release of zinc ions or to the nanosize as the peculiar surface interactions. Hence, the aim of the current study was to elucidate the different mechanistic modes of cytotoxicity of ZnO NPs towards marine algae, Chlorella vulgaris. The mechanistic end points included oxidative stress analyses, membrane permeability assessment, and ionic dissolution studies. This study is first of its kind in the usage of flow cytometry techniques to investigate the toxicity of ZnO NPs to cell viability in C. vulgaris and to offer a more theoretical foundation on which to evaluate the toxic effects of ZnO NPs.
2. Material and methods 2.1. Synthesis of ZnO NPs The preparation of ZnO NPs was done by the solid state pyrolytic method by mixing zinc acetate dihydrate (2.2 g, 10 mmol) and sodium bicarbonate (2.0 g, 23.8 mmol) at room temperature. The mixture was then pyrolysed at 300 °C for 3 h. Zinc acetate dihydrate gets reduced to ZnO NPs by thermal decomposition. Sodium acetate formed in the mixture is washed off with deionized water, to obtain pure ZnO NPs. 2.2. Characterization of ZnO NPs ZnO NPs were characterized by X-ray diffraction method (XRD), field emission scanning electron microscopy (FESEM) and energydispersive X-ray spectrometry (EDX). The XRD measurement of ZnO NPs was carried out on films of the respective solutions dropcoated onto a glass substrate on a Rigaku SmartLab instrument operated at a voltage of 9 kW and a current of 30 mA with CuKa radiations. The ZnO NPs were mounted on the copper stubs, and
the images were studied using FESEM with a Supra 55 (Carl Zeiss, Germany) with secondary electron detectors at an operating voltage of 5 kV. The elemental analysis was done by energy dispersive X-ray analysis (EDX) coupled to FESEM. 2.3. Dissolution of ZnO NPs The suspensions of ZnO NPs were prepared by dispersing ZnO NPs in sea water with a bath sonicator for 30 min to break aggregates as much as possible. To determine whether Zn2 þ dissolved from ZnO NPs suspensions might play a role in the observed toxicity, ZnO NPs (50–300 mg/L) were centrifuged at 10,000 rpm for 15 min and clear supernatants were carefully collected. The released Zn2 þ concentration in the supernatants were measured by Atomic Absorption Spectrometer. 2.4. Algae culture C. vulgaris was obtained from Marine Biotechnology Division, Earth System Sciences Organization – National Institute of Ocean Technology, Chennai, Tamil Nadu,India and cultured in autoclaved 1 L flask, filled with 400 ml of sea water and F/2 nutrients without trace metals and ethylenediaminetetraacetic acid (EDTA).The flasks were autoclaved at 121 °C for 20 min and allowed to cool for 24 h prior to use. The sterilized media were inoculated with cells of C. vulgaris, and the algae were cultured for 5–7 d with continuous aeration. Algae was incubated under cool white fluorescent lights (12 h light: 12 h dark) at a temperature of 20 °C. 2.5. Toxicological studies The exponential phase of C. vulgaris was used. Cells were treated with different concentrations (50, 100, 200, 300 mg/L) of the ZnO NP dispersion. An initial cell count of 4 105 cells was used for all experiments. The samples were kept in the static conditions, and intermittent mixing was carried out to avoid sticking of the cells to the walls of the container. Incubation was done under cool white fluorescent light (12 h light: 12 h dark) at a temperature of 20 °C. The samples were analyzed after an interaction period (with NPs) of 24 h and 72 h respectively. 2.6. Flow cytometric analysis In this study, fluorescence of cells stained with propidium iodide (PI) was measured to examine the cell viability. PI is a fluorescent dye that intercalates with double-stranded nucleic acids to produce red fluorescence when excited by blue light. But PI will not be able to pass through intact cell membranes of live cells. However, when the cell dies the integrity of the cell membrane fails and PI would be able to enter and stain the nucleic acids (Ormerod, 1990). PI can be used in this manner to discriminate between live nonfluorescent cells and dead fluorescent cells. Initially, cells were collected by centrifugation (3500 rpm, 10 min), washed with phosphate buffer solution (PBS, pH 7.0), and then stained with PI during an incubation period of 20 min. The final concentration of PI in the cell suspensions was 60 mM. The fluorescent emission of this compound was collected in the FACS Calibur- Becton Dickinson FL3 channel. 2.7. Cytotoxicity assay 2.7.1. MTT test In order to determine the cytotoxicity effect of ZnO NPs, 3-(4, 5-dimethylthiazol-2-yl) -2, 5-diphenyltetrazolium bromide (MTT) dye reduction assay was performed on C. vulgaris using increasing concentrations (10–300 mg/L). The result of the assay depends on
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the reduction of MTT to a blue-colored product by mitochondrial dehydrogenase, an enzyme present in the mitochondria of viable cells. After 24 and 72 h of interaction, 500 ml of the interacted cell suspension was added to 20 ml of MTT solution (5 mg MTT in 1 ml PBS) and incubated in the dark for a period of 4 h. The suspension was then centrifuged at 8000 rpm for 8 min. The pellet obtained was washed with 500 ml of seawater, and 200 ml of dimethyl sulfoxide was added. The absorbance was measured at 570 nm using a microplate reader Mosmann (1983). The cell viability was expressed considering untreated samples as 100% and the relative decrease was computed and the findings were validated through Flow cytometry. 2.7.2. Quantification of membrane damage through lactate dehydrogenase (LDH) assay About 1 ml of the ZnO NPs-interacted cell suspensions was centrifuged at 8000 rpm for 10 min. A 100 ml of the supernatant was then collected and 100 ml of 30 mM sodium pyruvate was added, followed by 2.8 ml of 0.2 M Tris–HCl. Finally, 100 ml of 6.6 mM NaDH was added just before measuring the decrease in absorbance at 340 nm as a function of 10 readings (Brown et al., 2004; Pakrashi et al., 2013) using UV–visible spectrophotometer. 2.7.3. Oxidative stress determination An enzymatic antioxidant (superoxide dismutase (SOD)), a nonenzymatic antioxidant (reduced glutathione (GSH)), and thiobarbituric acid-reactive substances (TBARS) were measured to assess the oxidative stress of ZnO NPs in the abalones. The SODspecific activities and GSH contents were determined using a spectrophotometer (Milton Roy Spec20, USA) at 550 nm and 412 nm respectively, according to the Diagnostic Reagent Kit. The SOD activity was measured based on the mechanism that the superoxides that are formed in the xanthine/xanthine oxidase superoxide generation system inhibit the formation of nitrite from hydroxylammonium chloride (Elstner and Heupel, 1976; Ji et al., 1991). One unit of SOD activity (U) is defined as the amount of enzyme that yields a 50% inhibition of nitrite formation in 1 ml of the reaction solution. GSH was studied as a non-enzymatic antioxidant. The GSH concentrations were determined using 5, 5-dithionitrobenzoic acid (Ellman, 1959). The results were expressed as mg mg 1 protein. Malonaldehyde (MDA), as an in vitro marker of lipid peroxidation, was measured by a method described by Del Rio et al. (2003). To a 200 ml of sample with a protein concentration of 2 mg/ml, 700 ml of 0.1 M HCl was added and the mixture was incubated for 20 min at room temperature. Then, 900 ml of 0.025 M thiobarbituric acid was added and the mixture was incubated for 65 min at 37 °C. Finally, 400 ml of 10 mM PBS was added. The fluorescence of MDA was recorded using a 520/549 (excitation/emission) filter. A calibration curve with MDA in the range 0.05–5 mM was used to calculate the MDA concentration. The results are expressed as nmoles of MDA/mg protein. 2.8. Surface chemical analysis through FTIR FTIR analysis was carried out to study the participation of surface groups in the interaction with ZnO NPs, which help to determine the extent of cell damage induced (Sadiq et al., 2011). After 72 h of exposure to ZnO NPs 300 mg/L, algal cells from 5 ml of suspension were harvested by centrifugation for 15 min at 10,000 rpm at an ambient temperature. The pellets were washed thrice with PBS and dried for 30 min at 3571 °C in a hot air oven. The dried cells were subjected to FTIR analysis by a potassium bromide technique using a Perkin-Elmer Spectrum.
Fig. 1. Image of dissolved Zn2 þ concentration (mg/L).
2.9. Microscopic analysis The samples treated with 300 mg/L ZnO NPs were collected after 72 h of interaction. A 250 ml of the sample was mixed with 250 ml of PI and incubated for 5 min in the dark at 25 °C followed by centrifugation at 200 rpm for 2 min at 4 °C. The pellet was washed thrice using SSC (saline sodium citrate) buffer to remove any unbound dye. About 10 ml of the resultant sample was carried onto a glass slide and sealed with cover slip for observation. These were observed using LSM 5 LIVE Duo Scan Confocal MicroscopeCarl ZEISS. 2.10. Field emission scanning electron microscopy Field emission scanning electron microscope (FESEM-Supra 55, -Carl Zeiss Germany) was used to observe the morphology of the algae and the particles. Both treated and untreated samples were drop coated on a 1 cm2 glass piece and air dried. The samples were subjected to gold sputtering and analyzed under FESEM. 2.11. Statistical analysis The experiments were carried out in replicates and the data are presented as mean 7standard error (SE). Significant differences between means were identified using one-way ANOVA, SPSS 20. Statistical significance was evaluated using significance levels at 0.05.
3. Results and discussion 3.1. Morphological characterization of ZnO NPs The XRD pattern (Fig. S1) Of ZnO NPs, which agrees with a database standard previously reported (JCPDS no. 89-1397), confirmed that the resulting product was a crystalline hexagonal – phase ZnO. The FESEM micrographs (Fig. S2A) show ZnO NPs that are spherical in shape and the size is in the range of 40–48 nm. EDAX analysis gives the additional evidence to the presence of elemental zinc (Fig. S2B). 3.2. Dissolution of ZnO NPs The dissolution rate of NPs is influenced by several factors, including particle size, surface area, surface curvature, and roughness of the particle (Farre´ et al., 2009). Zn2 þ solute from NPs can enter cells by transport and ion/voltage-gated channels (Colvin et al., 2003). Zn2 þ ions have the ability to form chelates with biomolecules or dislodge the metal ions in specific
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Table 1 A) Cell viability, B) LDH, C) SOD, D) GSH, E) LPO. Exposure period
Exposure conc ZnO NPs
A) Cell viability (%)
B) LDH (nmol/minml)
1 C) SOD (l mgprot )
1 D) GSH (mg mgpro )
1 E) LPO (nmol mgpro )
24 h
50 100 200 300 50 100 200 300
90.497 0.3 85.26 7 0.9 80.29 7 0.4 72.62 7 0.3 70.347 0.2 62.81 7 0.5 35.737 0.3 23.69 7 1.8
3.187 0.05 6.95 7 0.10 14.137 0.10 16.247 0.16 6.177 0.05 13.337 0.1 20.32 7 0.2 22.98 7 0.3
30.107 0.4 25.777 0.7 18.167 0.3 11.03 7 0.07 28.86 7 0.3 14.777 0.7 10.60 7 0.7 6.59 7 0.4
39.42 7 0.30 35.16 7 0.04 27.22 7 0.09 21.11 7 0.07 31.46 7 0.06 24.11 7 0.02 19.677 0.20 16.21 7 0.10
3.85 7 0.02 3.93 7 0.02 4.17 0.01 4.147 0.02 3.90 7 0.09 4.157 0.04 4.28 7 0.04 4.917 0.08
72 h
Fig. 2. Flow cytometry images: (A) control C. vulgaris showed nearly 100% of cell viability after 72 h; (B) 50 mg/L treated C. vulgaris viable decrease of 69.90% after 72 h; (C) 100 mg/L treated C. vulgaris viable decrease of 59.31% after 72 h; (D) 200 mg/L treated C. vulgaris viable decrease of 33.86% after 72 h and (E) 300 mg/L treated C. vulgaris viable decrease of 22.37% after 72 h.
metalloproteins, which may result in functional protein inactivation. Zn2 þ released by NPs increase their local concentration and disrupts cellular metal cation homeostasis to result in cell toxicity. Zinc oxide is nearly insoluble in pure water, where its water solubility ranges from 1.6 mg/L to 5 mg/L (PROSPEcT, 2009) whereas,
in a high ionic strength and high pH medium, such as the seawater, the solubility of the zinc oxide is significantly higher showing, in addition, some differences between the nano and bulk form (Miller et al., 2010; Miao et al., 2010; Peng et al., 2011; Wong et al., 2010). In our study to identify dissolution of NPs, the
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Fig. 3. Fourier Transform Infrared spectrum of control (A) and nanoparticle interacted (B) cells of C. vulgaris.
Fig. 4. Confocal images (A) untreated cell with intact membranes, preventing entry of PI dye, resulting in unstained cells and (B) cell treated with 300 mg/L for 72 h showing PI stained cells due to compromised membrane integrity.
concentrations of the released Zn2 þ were measured at 50– 300 mg/L of ZnO NPs. The results showed that the concentrations of releasing Zn2 þ gradually increased with increasing concentrations. The concentration of Zn2 þ dissolved from 50 mg/L of NPs was 3.96 mg/L. Even at the highest concentration (300 mg/L) of NPs, 10.68 mg/L of soluble Zn2 þ was detected in the present study (Fig. 1). Peng et al. (2011) observed a complete suppression (100% effect) of three marine diatomgrowths at 10 mg/L of nano ZnO. Some researchers considered that dissolved zinc ions played an important role in the toxicity of ZnO NPs (Song et al., 2010). 3.3. Cytotoxicity ZnO NPs were strongly cytotoxic at lower concentrations and exhibited strong protein adsorption abilities (Horie et al., 2009) which may have contributed to their cytotoxicity. ZnONPs taken up by cells may cause rapid dissolution, releasing Zn2 þ ions (Xia et al., 2008), elevated intracellular Zn2 þ induced the disruption of mitochondrial function (Bishop et al., 2007; Wiseman et al., 2007) and triggered ROS generation leading to cell death (Xia et al., 2008). The cytotoxic effect of ZnO NPs on C. vulgaris with the increasing concentration (10–300 mg/L) was evaluated using MTT assay. The ZnO NPs exerted a cytotoxic effect on C. vulgaris at lower concentrations of 50 mg/L. The viability of C. vulgaris decreased by 90.49 70.3% at 24 h upon 50 mg/L ZnO NPs, whereas upon 300 mg/L, the viability of C. vulgaris significantly reduced up to
23.697 1.8% (Table 1A). A similar phenomenon reported by Sharma et al. (2009) indicated that ZnO NPs induced cytotoxicity was concentration and time dependent. Lactate dehydrogenase (LDH) assay was carried out to confirm the intactness of the cell membrane (Jeng and Swanson, 2006). Any damage in the membrane, which leads to increased levels of lactate dehydrogenase enzyme in the test solution, can be one of the reasons for cell death. Zhang et al. (2012) have considered LDH assay as one of the single parameters for determining cellular toxicity of metal oxide nanoparticles. A significant (p o0.05) LDH leakage was observed at 24 h and 72 h on exposure of 14.13 70.14 at 200 mg/L, 16.24 70.16 at 300 mg/L of 24 h and 20.32 70.26 at 200 mg/L of 24 h, 22.74 70.33 at 300 mg/L of 48 h (p o0.05) (Table 1B). 3.4. Flow cytometric analysis of cell viability To determine the extent of cytotoxicity in C. vulgaris cells, the ZnO NP-treated cells were analyzed using flow cytometry. In flow cytometry images, upper left and right quadrants show the percentage of dead cells and lower left and right quadrants show the percentage of live cells (Fig. 4). The cell viability of the control group was almost 100% (Fig. 2A) during the 72 h of culture; nevertheless, in cultures with the various concentrations of ZnO NPs the cell viability was found to have decreased after 72 h. The percentage of viable cells decreased to 68.09%, 59.31%, 33.86% and
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Fig. 5. Scanning electron micrographs: (A) control cell shows no damage; (B) (C) extensive membrane damage upon 72 h treatment with the 50 mg/L and 100 mg /L; D and E 300 mg/L treated cell shows distorted morphological features after 72 h treatment. (For interpretation of the references to color in this figure, the reader is referred to the web version of this article.)
22.37% for 50 mg/L (Fig. 2B), 100 mg/L (Fig. 2C), 200 mg/L (Fig. 2D), and 300 mg/L (Fig. 2E) concentrations, respectively. This dose and exposure time dependence corroborate with MTT assay results as well. 3.5. Oxidative stress determination The potential of ZnO NPs to induce oxidative stress was assessed by measuring the SOD in algal cells (Table 1C). An exposure time of 72 h was found to induce SOD (6.59 7 0.4) significantly in algal cells at 300 mg/L of ZnO NPs. These results show that oxidative stress is increased by ZnO NPs. A similar phenomenon was reported by Chen et al. (2012) for C. vulgaris; the activity of
antioxidant enzymes followed a changing trend of being activated by low concentrations of ZnONPs and inhibited by higher concentrations (Hao and Chen, 2012). When the cell was exposed to ZnO NPs for 72 h, GSH level significantly decreased. When the ZnO NPs dose increased to 200 mg/L, the intracellular GSH level were almost reduced by half 19.677 0.2 compared to the control (Table 1D). The data demonstrated a significant depletion of GSH level also in ZnO NPs exposed algal cells. The GSH content decreased in the high concentration of ZnONPs stress, which may be attributed to the consumption of a large amount of the GSH in a variety of detoxification processes. At high concentrations of ZnO NPs destruction of antioxidation structures and a decrease in the level of antioxidants probably occur, which result in rapid
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accumulation of ROS. Lipid peroxidation is an indicator of oxidative damage to cell membrane lipids and has been used extensively as a biomarker for oxidative stress in vivo (Sayeed et al., 2003). It is estimated by measuring the MDA content of cells.The MDA content significantly increased at ZnO NPs concentrations higher than 200 mg/L (Table 1E). 3.6. Surface characterization by FTIR FTIR has emerged as a valuable instrument for understanding the involvement of surface charge-based attachment of NPs onto algal cells, leading to membrane damage as well as aggregate formation. Control spectra of the uninteracted cell (not treated with ZnO NPs) displayed a number of peaks (Fig. 3A). There was a variation in the intensity of bands in test samples (treated with ZnO NPs) (Fig. 3B). A control spectrum of uninteracted cell shows peak at 3390 cm 1 representing the presence of hydrogen-bonded O–H stretch. The peak at 2929.73 cm 1 and 699.70 cm 1 represents the presence of C–H stretch. The peaks at 1609.88 cm 1 and 933.33 cm 1 took place at the stretching vibration of N–H primary amine. The peaks at 1261 cm 1 and 1120 cm 1 were the stretching vibration of C–O. The treated cells with ZnO NPs showed a peak at 3334.33 cm 1 representing the presence of hydrogen-bonded O–H stretch. The surfaces of algae contain a number of functional groups with high affinity for metal ions and carry a net negative charge, due mainly to carboxylic, sulfhydryl, and phosphatic groups (Crist et al., 1990, 1994). These groups are binding sites that transport metal ions across the cell membrane and into the cell. Various studies have demonstrated that two distinct phases occur in the ingestion of metals by algal cells: a rapid assimilation that is terminated within 10 min followed by a slower, facilitated transport into the cytoplasm of the cell (Gonza ´lez Da´vila et al., 1995; Xue et al., 1988). Adsorption of metal ions to the cell surface determines the initial toxicant loading of the cells, and this is likely to be dependent on the algal cell surface area and also the nature of the binding sites (Geisweid and Urbach,1983). The peak at 1606.06 cm 1shows the stretching vibration of N–H primary amine. The peak at 1259.82 cm 1 was the stretching vibration of C–O. The peak at 542.07 cm 1 was associated with the stretching vibration of ZnO (Wahab et al., 2008). The conformation of surface chemical interaction led us to probe further for membrane disruption followed by attachment. 3.7. Microscopic analysis The compromised cellular integrity and permeability was confirmed through confocal laser scanning microscopy which allowed the visualization of stained nuclei (a nuclear-specific dye PI was used) in case of damaged cells upon ZnO NP exposure. No PI fluorescence was observed in control cells after 72 h (Fig. 4A). PI fluorescence was detected in the nucleus after 72 h (Fig. 4B). The membrane damage as a sign of cell death has also been reported by Metzler et al. (2011) in Pseudokirchneriella subcapitata upon TiO2 nanoparticle interaction.The fluorescence microscopic observation corroborated well with FESEM observation. The FESEM micrograph showed intact structural features in untreated cells and the size of the untreated cell was 3.41 mm (Fig. 5A). In contrast, the treated cell reconfirmed the cell wall (marked with red pointer) upon exposure to 50 mg/L (Fig. 5B) and 100 mg/L (Fig. 5C) ZnO NPs. In addition, a direct visualization at high resolution also revealed morphological alterations of cell structure damage with 200 mg/L and 300 mg/L (Fig. 5D and E) after 72 h and the size of the treated cell was 1.83 mm. Membrane damage as a sign of cell death has also been reported by Pakrashi et al. (2013).
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4. Conclusions To our knowledge, this is one of the first focused, systematic studies on the effect of ZnO NPs on marine microalgae C. vulgaris and first in describing through flowcytometry, cytotoxicity, oxidative stress mechanism and morphology. The results showed that C. vulgaris viability was affected by an increase of ZnO NPs concentrations. ZnO NPs were found to cause a substantial outcome on the MDA, GSH, SOD, and LDH. FTIR study of the surface chemistry of the interacted cells demonstrates probable interaction between the particles and the surface sites on the cell wall. The result showed the interaction between the ZnO NPs and algae favoring the formation of distorted morphological features after 72 h treatment. The changes observed in the activity of SOD, GSH, LPO, LDH and decrease of viable cells, thus established an end point for ZnO NPs toxicity.
Acknowledgments Authors are thankful to IFS (Sweden) (No. A/5041-1), for the financial support extended; Centre for Cellular and Molecular Platforms, NCBS, Bangalore, for their help in analyzing the sample by Flow Cytometry and Confocal Microscopy; and Centre for Nanoscience and Nanotechnology, Sathyabama University, Jeppiaar Nagar, Chennai in carrying out FESEM, XRD, and EDX.
Appendix A. Supplementary material Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.ecoenv.2014.11.015.
References Aschberger, K., Micheletti, C., Sokull-Kl ̈ uttgen, B., Christensen, F.M., 2011. Analysis of currently available data for characterizing the risk of engineered nanomaterials to the environment and human health lessons learned from four case studies. Environ. Int. 37, 1143–1156. Bishop, G.M., Dringen, R., Robinson, S.R., 2007. Zinc stimulates the production of toxic, reactive oxygen species (ROS) and inhibits glutathione reductase in astrocytes. Free Radic. Biol. Med. 42, 1222–1230. Blaise, C., Gagne´, F., Ferard, J.F., 2008. Ecotoxicity of selected nanomaterials to aquatic organisms. Environ. Toxicol. 23, 591–598. Brayner, R., Ferrari-Iliou, R., Brivois, N., Djediat, S., Benedetti, M.F., Fievet, F., 2006. Toxicological impact studies based on Escherichia coli bacteria in ultrafine ZnO nanoparticles colloidal medium. Nano Lett. 6, 866–870. Brown, D.M., Donaldson, K., Borm, P.J., Schins, R.P., Dehnhardt, M., Gilmour, P., 2004. Calcium and ROS-mediated activation of transcription factors and TNF alpha cytokine gene expression in macrophages exposed to ultrafine particles. Am. J. Physiol. – Lung Cell. Mol. Physiol. 286, 344–353. Chen, X., Zhu, X., Li, R., Yao, H., Lu, Z., Yang, X., 2012. Photosynthetic toxicity and oxidative damage induced by nano-Fe3O4 on Chlorella vulgaris in the aquatic environment. Open J. Ecol. 1, 21–28. Colvin, R.A., Fontaine, C.P., Laskowski, M., Thomas, D., 2003. Zn2 þ transporters and Zn2þ homeostasis in neurons. Eur. J. Pharmacol. 479, 171–185. Crist, R.H., Martin, J.R., Guptill, P.W., Eslinger, J.M., Crist, D.R., 1990. Interaction of Metal Ions with Acid Sites of Biosorbents Peat Moss and Vaucheria and Model Substances Alginic and Humic Acids. Environ Sci Technol 24, 337–342. Crist, R.H., Martin, J.R., Carr, D., Watson, J.R., Clarke, H.J., Crist, D.R., 1994. Interaction of metals and protons with algae. 4. Ion-exchange vs adsorption models and a reassessment of Scatchard plots; ion-exchange rates and equilibria compared with calcium alginate. Environ. Sci. Technol. 28, 1859–1866. De Berardis, B., Civitelli, G., Condello, M., Lists, P., Pozzi, R., Arancia, G., Meschini, S., 2010. Exposure to ZnO nanoparticles induces oxidative stress and cytotoxicity in human colon carcinoma cells. Toxicol. Appl. Pharmacol. 246, 116–127. Del Rio, D., Pellegrini, N., Colombi, B., Bianchi, M., Serafini, M., Torta, F., Tegoni, M., Musci, M., Brighenti, F., 2003. Rapid fluorimetric method to detect total plasma malondialdehyde with mild derivatization conditions. Clin. Chem. 49, 690–692. Ellman, G.L., 1959. Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82, 70–77. Elstner, E.F., Heupel, A., 1976. Inhibition of nitrite formation from hydroxylammoniumchloride: a simple assay for superoxide dismutase. Anal. Biochem. 70, 616–620.
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Farre´, M., Gajda-Schrantz, K., Kantiani, L., Barcelo, D., 2009. Ecotoxicity and analysis of nanomaterials in the aquatic environment. Anal. Bioanal. Chem. 393, 81–95. Franklin, N.M., Rogers, N.J., Apte, S.C., Batley, G.E., Gadd, G.E., 2007. Comparative toxicity of nanoparticulate ZnO, bulk ZnO, and ZnCl2 to a freshwater microalga (Pseudokirchneriella subcapitata): the importance of particle solubility. Environ. Sci. Technol. 41, 8484–8490. Geisweid, H.J., Urbach, W., 1983. Sorption of cadmium by the green microalgae Chlorella vulgaris, Ankistrodesmus braunii and Eremosphaera viridis. Z. Planzenphysiol. 109, 127–141. Gonza´lez Da´vila, M., Santana-Casiano, J.M., Pe´rez-Pen~a, J., Millero, F.J., 1995. Binding of Cu (II) to the surface and exudates of the alga Dunaliella tertiolecta in seawater. Environ. Sci. Technol. 29, 289–301. Gottschalk, F., Sonderer, T., Scholz, R.W., Nowack, B., 2009. Modeled environmental concentrations of engineered nanomaterials (TiO2, ZnO, Ag, CNT, fullerenes) for different regions. Environ. Sci. Technol. 43, 9216–9222. Hao, L., Chen, L., 2012. Oxidative stress responses in different organs of carp (Cyprinus carpio) with exposure to ZnO nanoparticles. Ecotoxicol. Environ. Saf. 80, 103–110. Heinlaan, M., Ivask, A., Blinova, I., Dubourguier, H., 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. Horie, M., Nishio, K., Fujita, K., Endoh, S., Miyauchi, A., Saito, Y., Iwahashi, H., Yamamoto, K., Murayama, H., Nakano, H., 2009. Protein adsorption of ultrafine metal oxide and its influence on cytotoxicity toward cultured cells. Chem. Res. Toxicol. 22, 543–553. Jeng, H.A., Swanson, J., 2006. Toxicity of metal oxide nanoparticles in mammalian cells. J. Environ. Sci. Health A Toxic Hazard. Subst. Environ. Eng. 41, 2699–2711. Jiang, W., Mashayekhi, H., Xing, B., 2009. Bacterial toxicity comparison between nano- and micro-scaled oxide particles. Environ. Pollut. 157, 1619–1625. Ji, J., Wu, Z., Liu, Q., Zhang, Y., Ye, M., Li, M., 1991. An ultramicro analytic and rapid method for determination of superoxide dismutase activity. J. Nanjing Railw. Med. Coll. 10, 27–30. Lin, D.H., Xing, B.S., 2007. Phytotoxicity of nanoparticles: inhibition of seed germination and root growth. Environ. Pollut. 150, 243–250. Manzo, S., Miglietta, M.L., Rametta, G., Buono, S., Di Francia, G., 2013. Toxic effects of ZnO nanoparticles towards marine algae Dunaliella tertiolecta. Sci. Total Environ. 445, 371–376. Miao, A.J., Zhang, X.Y., Luo, Z., Chen, C.S., Chin, W.C., Santschi, P.H., 2010. Zinc oxide engineered nanoparticles: dissolution and toxicity to marine phytoplankton. Environ. Toxicol. Chem. 29, 2814–2822. Miller, R.J., Lenihan, H.S., Muller, E.B., Tseng., N., Hanna, S.K., Keller, A.A., 2010. Impact of metal oxide nanoparticles on marine phytoplankton. Environ. Sci. Technol. 44, 7329–7334. Metzler, D.M., Li, M., Erdem, A., Huang, C., 2011. Responses of algae to photocatalytic nano-TiO2 particles with an emphasis on the effect of particle size. Chem. Eng. J. 170 (2–3), 538–546. Mosmann, T., 1983. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. Immunol. Methods 65, 55. Nair, S., Sasidharan, A., Rani, V.V.D., Menon, D., Nair, S., Manzoor, K., Raina, S., 2009. Role of size scale of ZnO nanoparticles and microparticles of toxicity toward bacteria and osteoblast cancer cells. J. Mater. Sci. Mater. Med. 20, 235–241. Ormerod, M.G., 1990. Analysis of DNA. General methods. In: Ormerod, M.G. (Ed.), Flow Cytometry. A Practical Approach. Oxford University Press, Oxford, pp. 69–87. Osmond, M.J., McCall, M.J., 2010. Zinc oxide nanoparticles in modern sunscreens: an analysis of potential exposure and hazard. Nanotoxicology 4, 15–41. Pakrashi, S., Dalai, S., Prathna, T.C., Trivedi, S., Myneni, R., Raichur, A.M., Chandrasekaran, N., Mukherjee, A., 2013. Cytotoxicity of aluminium oxide nanoparticles towards freshwater algal isolate at low exposure concentrations. Aquat. Toxicol. 133, 34–45. Peng, X., Palma, S., Fisher, N.S., Wong, S.S., 2011. Effect of morphology of ZnO nanostructures on their toxicity to marine algae. Aquat. Toxicol. 102, 186–196. PROSPEcT, 2009. Ecotoxicology test protocols for representative nanomaterials in support of the OECD sponsorship programme. 8. Premanathan, M., Karthikeyan, K., Jeyasubramanian, K., Manivannan, G., 2011. Selective toxicity of ZnO nanoparticles toward gram-positive bacteria and cancer
cells by apoptosis through lipid peroxidation. Nanomed. Nanotechnol. Biol. Med. 7, 184–192. Reddy, K.M., Feris, K., Bell, J., Wingett, D.G., Hanley, C., Punnoose, A., 2007. Selective toxicity of zinc oxide nanoparticles to prokaryotic and eukaryotic systems. Appl. Phys. Lett. 90, 2139021–2139023. Sayeed, I., Parvez, S., Pandey, S., Bin-Hafeez, B., Haque, R., Raisuddin, S., 2003. Oxidative stress biomarkers of exposure to deltamethrin in freshwater fish, Channa punctatus Bloch. Ecotoxicol. Environ. Saf. 56, 295–301. Sadiq, I.M., Pakrashi, S., Chandrasekaran, N., Mukherjee, A., 2011. Studies on the toxicity of aluminum oxide (Al2O3) nanoparticles to microalgae species Scenedesmus Sp. and Chlorella Sp. J. Nanopart. Res. 13, 3287–3299. Sharifi, S., Behzadi, S., Laurent, S., Laird Forrest, M., Stroeve, P., Mahmoudi, M., 2012. Toxicity of nanomaterials. Chem. Soc. Rev. 41, 2323–2343. Sharma, V., Shukla, R.K., Saxena, N., Parmar, D., Das, M., Dhawan, A., 2009. DNA damaging potential of zinc oxide nanoparticles in human epidermal cells. Toxicol. Lett. 185, 211–218. Song, W., Zhang, J., Guo, J., Zhang, J., Ding, F., Li, L., Sun, Z., 2010. Role of the dissolved zinc ion and reactive oxygen species in cytotoxicity of ZnO nanoparticles. Toxicol. Lett. 15, 389–397. Song, Y., Zhu, L., Wang, S., Wang, J., Liu, J.H., Xie, H., W., 2008. DNA damage and effects on antioxidative enzymes in earthworm (Eisenia fetida) induced by atrazine. Soil Biol. Biochem. 41, 905–909. Wang, B., Feng, W.Y., Wang, M., Wang, T.C., G.U., Y.Q., Zhu, M.T., Ouyang, H., Shi, J.W., Zhang, F., Zhao, Y.L., Chai, Z.F., Wang, H.F., Wang, J., 2008a. Acute toxicological impact of nano- and submicro-scaled zinc oxide powder on healthy adult mice. J. Nanopart. Res. 10, 263–276. Wahab, R., Ansari, S.G., Kim, Y.S., Dar, M.A., Shin, H.S., 2008. Synthesis and characterization of hydrozincite and its conversion into zinc oxide nanoparticles. J. Alloys Compd. 461, 66–71. Weckx, J., Clijsters, H., 1996. Oxidative damage and defense mechanisms in primary leaves of Phaseolus vulgaris as a result of root assimilation of toxic amounts of copper. Physiol. Plant. 96, 506–512. Wiseman, D.A., Wells, S.M., Hubbard, M., Welker, J.E., Black, S.M., 2007. Alterations in zinc homeostasis underlie endothelial cell death induced by oxidative stress from acute exposure to hydrogen peroxide. Am J Physiol Lung Cell Mol Physiol 292, 165–177. Wiench, K., Wohlleben, W., Hisgen, V., Radke, K., Salinas, E., Zok, S., Landsiedel, R., 2009. Acute and chronic effects of nano and non-nano-scaleTiO2 and ZnO particles on mobility and reproduction of the fresh water invertebrate Daphnia magna. Chemosphere 76, 1356–1365. Wong, S.W., Leung, P.T.Y., Djurisić, A.B., Leung, K.M., 2010. Toxicities of nano zinc oxide to five marine organisms: influences of aggregate size and ion solubility. Anal. Bioanal. Chem. 396, 609–618. Xia, T., Kovochich, M., Liong, M., Madler, L., Gilbert, B., Shi, H., Yeh, J.I., Zink, J.I., Nel, A.E., 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. Xiong, D.W., Fang, T., Yu, L.P., Sima, X.F., Zhu, W.T., 2011. Effectsofnano-scaleTiO2, ZnO and their bulk counterparts on zebrafish: acute toxicity, oxidative stress and oxidative damage. Sci.Total Environ. 409, 1444–1452. Xue, H., Stumm, W., Sigg, L., 1988. The binding of heavy metals to algal surfaces. Water Res. 22, 917–926. Zhang, H., Ji, Z., Xia, T., Meng, H., Low-Kam, C.R., Liu, S., Pokhrel, S., Lin, X., Wang, Y. P., Liao, M., Wang, L., Li, R., Rallo, R., Damoiseaux, D., Telesca, L., Mädler, Y., Cohen, J.I., Zink, A., Nel, E., 2012. Use of metal oxide nanoparticle band gap to develop a predictive paradigm for oxidative stress and acute pulmonary inflammation. ACS Nano 6, 4349–4368. Zhao, J., Castranova., V., 2011. Toxicology of nanomaterials used in nanomedicine. J. Toxicol. Environ. Health B 14, 593–632. Zhu, X.S., Zhu, L., Duan, Z.H., Qi, R.Q., Li, Y., Lang, Y.P., 2008. Comparative toxicity of several metal oxide nanoparticle aqueous suspensions in zebrafish (Daniorerio) early developmental stage. J. Environ. Sci. Health 3 (3), 278–284. Zhu, X.S., Zhu, L., Chen, Y.S., Tian, S.Y., 2009. Acute toxicities of six manufactured nanomaterial suspensions to Daphnia magna. J. Nanopart. Res. 11, 67–75.
Contents lists available at ScienceDirect
Ecotoxicology and Environmental Safety journal homepage: www.elsevier.com/locate/ecoenv
Evaluation of zinc oxide nanoparticles toxicity on marine algae chlorella vulgaris through flow cytometric, cytotoxicity and oxidative stress analysis T.Y. Suman a, S.R. Radhika Rajasree a,n, R. Kirubagaran b a b
Center for Ocean Research (NIOT-SU Collaborative Research Centre), Sathyabama University, Chennai 600119, Tamil Nadu, India Marine Biotechnology, ESOO-National Institute of Ocean Technology, OSTI, Pallikaranai, Chennai 600100, India
art ic l e i nf o
a b s t r a c t
Article history: Received 26 July 2014 Received in revised form 14 November 2014 Accepted 19 November 2014
The increasing industrial use of nanomaterials during the last decades poses a potential threat to the environment and in particular to organisms living in the aquatic environment. In the present study, the toxicity of zinc oxide nanoparticles (ZnO NPs) was investigated in Marine algae Chlorella vulgaris (C. vulgaris). High zinc dissociation from ZnONPs, releasing ionic zinc in seawater, is a potential route for zinc assimilation and ZnONPs toxicity. To examine the mechanism of toxicity, C. vulgaris were treated with 50 mg/L, 100 mg/L, 200 mg/L and 300 mg/L ZnO NPs for 24 h and 72 h. The detailed cytotoxicity assay showed a substantial reduction in the viability dependent on dose and exposure. Further, flow cytometry revealed the significant reduction in C. vulgaris viable cells to higher ZnO NPs. Significant reductions in LDH level were noted for ZnO NPs at 300 mg/L concentration. The activity of antioxidant enzyme superoxide dismutase (SOD) significantly increased in the C. vulgaris exposed to 200 mg/L and 300 mg/L ZnO NPs. The content of non-enzymatic antioxidant glutathione (GSH) significantly decreased in the groups with a ZnO NPs concentration of higher than 100 mg/L. The level of lipid peroxidation (LPO) was found to increase as the ZnO NPs dose increased. The FT-IR analyses suggested surface chemical interaction between nanoparticles and algal cells. The substantial morphological changes and cell wall damage were confirmed through microscopic analyses (FESEM and CM). & 2014 Elsevier Inc. All rights reserved.
Keywords: Chlorella vulgaris Cytotoxicity Flow cytometry Oxidative stress ZnO NPs
1. Introduction Nanotechnology has come to the forefront of research in the past decade. With the advent of this technology, wide varieties of nanoparticles (NPs) with a variety of unique characteristics are manufactured and are used for a broad range of applications (Zhao and Castranova., 2011). Metal oxide NPs are among the most used engineered NPs in various commercial products, leading to concerns of their potential toxicity to human and environmental health (Aschbergeretal., 2011). Increasing use of metaloxide NPs is likely to result in the release of these particles into the aquatic environment. Adverse effects of NPs on the aquatic environment and organisms recently have drawn much special attention (Blaise etal., 2008; Farre´ et al., 2009). Among metal oxide nanomaterials, zinc oxide nanoparticles (ZnO NPs) are noted for their chemical stability and strong adsorption ability and have been extensively used in commercial n
Corresponding author. Fax: þ 44 24501270. E-mail address: [email protected] (S.R. Radhika Rajasree).
http://dx.doi.org/10.1016/j.ecoenv.2014.11.015 0147-6513/& 2014 Elsevier Inc. All rights reserved.
products like sunscreens, coatings, and paints (Osmond and McCall., 2010). ZnO has a high inherent risk of water contamination, and can reach high concentrations in surface waters posing significant threat to aquatic ecosystems (Gottschalk et al., 2009). Previous studies have demonstrated that ZnO NPs are toxic to microorganisms, cells, plants, aquatic biota and rodents (Brayner et al., 2006; Lin and Xing., 2007; Premanathan et al., 2011; Reddy et al., 2007; Wang et al., 2008a; Zhu et al., 2009). Zhu et al. (2008) found that ZnO NPs had higher acute toxicity to zebrafish embryos than nano-TiO2 and nano-Al2O3. The toxicity of ZnO NPs (96hLC50, 4.9 mg/L)to zebra fish was also much higher than that of nano-TiO2 (96 hLC50, 124.5 mg/L) (Xiong et al., 2011). Increased production of antioxidant enzymes in organisms is regarded as an early warning indicator of pollution in the environment (Song et al., 2008). Antioxidant enzyme activity can protect cells from the adverse effects of reactive oxygen species (ROS). Superoxide dismutase (SOD), catalase (CAT), and peroxidase (POD) enzymes help to check cellular levels of ROS (Weckx and Clijsters, 1996). Studies of the biotoxicity of ZnO NPs suggest several mechanisms of action of NPs, one of which is their ability to release free zinc ions which
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can synergistically enhance the production of ROS and cause oxidative damage in cells. Overproduction of ROS is believed to be a major mechanism of the toxicity of NPs (Sharifi et al., 2012). During this process, chemical reactions occur and result in increased formation of the superoxide radical (O2 ), which leads to ROS accumulation and oxidative stress (De Berardis et al., 2010). It has been reported that ZnONPs disturb the balance between oxidation and anti-oxidation processes and cause oxygen stress responses (Hao and Chen, 2012). As primary producers, phytoplankton plays a key role in aquatic ecosystems. Microalgae, being sensitive to pollutants, are excellent aquatic models and being widely prevalent in lakes and seas is easy to culture and propagate (Chen et al., 2012). The evaluation of NP effects upon marine phytoplankton is a necessary step to predict their potential impact on coastal marine food webs and on the whole ecosystems they support (Manzo et al., 2013). ZnO NPs, on entering the aquatic environment, release free metal ions in solution state. The dissolution speed of the particles mainly depends on the particle size, surface area, rough degree, etc. ZnO is slightly soluble, and can release zinc ions into the solution. Some researchers consider that dissolved zinc ions play an important role in the toxicity of ZnO NPs (Song et al., 2010). Some studies report that the toxicity of ZnO NPs was closely linked with its dissolved free ions (Franklin et al., 2007; Heinlaan et al., 2008; Wienchet al., 2009), but others have also shown that the toxicity of ZnO NPs was much higher than its ion toxicity (Nair et al., 2009; Wong et al., 2010). Both Escherichia coli and Pseudomonas fluorescens exhibited a high tolerance to zinc ions at the tested ZnO concentration (20 mg/L). The attachment of ZnO NPs to the surface of bacteria was the main cause of the ZnO toxicity of these bacterial species (Jiang et al., 2009).The point is that, at current knowledge, the observed ZnO toxicity cannot be attributed univocally to the release of zinc ions or to the nanosize as the peculiar surface interactions. Hence, the aim of the current study was to elucidate the different mechanistic modes of cytotoxicity of ZnO NPs towards marine algae, Chlorella vulgaris. The mechanistic end points included oxidative stress analyses, membrane permeability assessment, and ionic dissolution studies. This study is first of its kind in the usage of flow cytometry techniques to investigate the toxicity of ZnO NPs to cell viability in C. vulgaris and to offer a more theoretical foundation on which to evaluate the toxic effects of ZnO NPs.
2. Material and methods 2.1. Synthesis of ZnO NPs The preparation of ZnO NPs was done by the solid state pyrolytic method by mixing zinc acetate dihydrate (2.2 g, 10 mmol) and sodium bicarbonate (2.0 g, 23.8 mmol) at room temperature. The mixture was then pyrolysed at 300 °C for 3 h. Zinc acetate dihydrate gets reduced to ZnO NPs by thermal decomposition. Sodium acetate formed in the mixture is washed off with deionized water, to obtain pure ZnO NPs. 2.2. Characterization of ZnO NPs ZnO NPs were characterized by X-ray diffraction method (XRD), field emission scanning electron microscopy (FESEM) and energydispersive X-ray spectrometry (EDX). The XRD measurement of ZnO NPs was carried out on films of the respective solutions dropcoated onto a glass substrate on a Rigaku SmartLab instrument operated at a voltage of 9 kW and a current of 30 mA with CuKa radiations. The ZnO NPs were mounted on the copper stubs, and
the images were studied using FESEM with a Supra 55 (Carl Zeiss, Germany) with secondary electron detectors at an operating voltage of 5 kV. The elemental analysis was done by energy dispersive X-ray analysis (EDX) coupled to FESEM. 2.3. Dissolution of ZnO NPs The suspensions of ZnO NPs were prepared by dispersing ZnO NPs in sea water with a bath sonicator for 30 min to break aggregates as much as possible. To determine whether Zn2 þ dissolved from ZnO NPs suspensions might play a role in the observed toxicity, ZnO NPs (50–300 mg/L) were centrifuged at 10,000 rpm for 15 min and clear supernatants were carefully collected. The released Zn2 þ concentration in the supernatants were measured by Atomic Absorption Spectrometer. 2.4. Algae culture C. vulgaris was obtained from Marine Biotechnology Division, Earth System Sciences Organization – National Institute of Ocean Technology, Chennai, Tamil Nadu,India and cultured in autoclaved 1 L flask, filled with 400 ml of sea water and F/2 nutrients without trace metals and ethylenediaminetetraacetic acid (EDTA).The flasks were autoclaved at 121 °C for 20 min and allowed to cool for 24 h prior to use. The sterilized media were inoculated with cells of C. vulgaris, and the algae were cultured for 5–7 d with continuous aeration. Algae was incubated under cool white fluorescent lights (12 h light: 12 h dark) at a temperature of 20 °C. 2.5. Toxicological studies The exponential phase of C. vulgaris was used. Cells were treated with different concentrations (50, 100, 200, 300 mg/L) of the ZnO NP dispersion. An initial cell count of 4 105 cells was used for all experiments. The samples were kept in the static conditions, and intermittent mixing was carried out to avoid sticking of the cells to the walls of the container. Incubation was done under cool white fluorescent light (12 h light: 12 h dark) at a temperature of 20 °C. The samples were analyzed after an interaction period (with NPs) of 24 h and 72 h respectively. 2.6. Flow cytometric analysis In this study, fluorescence of cells stained with propidium iodide (PI) was measured to examine the cell viability. PI is a fluorescent dye that intercalates with double-stranded nucleic acids to produce red fluorescence when excited by blue light. But PI will not be able to pass through intact cell membranes of live cells. However, when the cell dies the integrity of the cell membrane fails and PI would be able to enter and stain the nucleic acids (Ormerod, 1990). PI can be used in this manner to discriminate between live nonfluorescent cells and dead fluorescent cells. Initially, cells were collected by centrifugation (3500 rpm, 10 min), washed with phosphate buffer solution (PBS, pH 7.0), and then stained with PI during an incubation period of 20 min. The final concentration of PI in the cell suspensions was 60 mM. The fluorescent emission of this compound was collected in the FACS Calibur- Becton Dickinson FL3 channel. 2.7. Cytotoxicity assay 2.7.1. MTT test In order to determine the cytotoxicity effect of ZnO NPs, 3-(4, 5-dimethylthiazol-2-yl) -2, 5-diphenyltetrazolium bromide (MTT) dye reduction assay was performed on C. vulgaris using increasing concentrations (10–300 mg/L). The result of the assay depends on
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the reduction of MTT to a blue-colored product by mitochondrial dehydrogenase, an enzyme present in the mitochondria of viable cells. After 24 and 72 h of interaction, 500 ml of the interacted cell suspension was added to 20 ml of MTT solution (5 mg MTT in 1 ml PBS) and incubated in the dark for a period of 4 h. The suspension was then centrifuged at 8000 rpm for 8 min. The pellet obtained was washed with 500 ml of seawater, and 200 ml of dimethyl sulfoxide was added. The absorbance was measured at 570 nm using a microplate reader Mosmann (1983). The cell viability was expressed considering untreated samples as 100% and the relative decrease was computed and the findings were validated through Flow cytometry. 2.7.2. Quantification of membrane damage through lactate dehydrogenase (LDH) assay About 1 ml of the ZnO NPs-interacted cell suspensions was centrifuged at 8000 rpm for 10 min. A 100 ml of the supernatant was then collected and 100 ml of 30 mM sodium pyruvate was added, followed by 2.8 ml of 0.2 M Tris–HCl. Finally, 100 ml of 6.6 mM NaDH was added just before measuring the decrease in absorbance at 340 nm as a function of 10 readings (Brown et al., 2004; Pakrashi et al., 2013) using UV–visible spectrophotometer. 2.7.3. Oxidative stress determination An enzymatic antioxidant (superoxide dismutase (SOD)), a nonenzymatic antioxidant (reduced glutathione (GSH)), and thiobarbituric acid-reactive substances (TBARS) were measured to assess the oxidative stress of ZnO NPs in the abalones. The SODspecific activities and GSH contents were determined using a spectrophotometer (Milton Roy Spec20, USA) at 550 nm and 412 nm respectively, according to the Diagnostic Reagent Kit. The SOD activity was measured based on the mechanism that the superoxides that are formed in the xanthine/xanthine oxidase superoxide generation system inhibit the formation of nitrite from hydroxylammonium chloride (Elstner and Heupel, 1976; Ji et al., 1991). One unit of SOD activity (U) is defined as the amount of enzyme that yields a 50% inhibition of nitrite formation in 1 ml of the reaction solution. GSH was studied as a non-enzymatic antioxidant. The GSH concentrations were determined using 5, 5-dithionitrobenzoic acid (Ellman, 1959). The results were expressed as mg mg 1 protein. Malonaldehyde (MDA), as an in vitro marker of lipid peroxidation, was measured by a method described by Del Rio et al. (2003). To a 200 ml of sample with a protein concentration of 2 mg/ml, 700 ml of 0.1 M HCl was added and the mixture was incubated for 20 min at room temperature. Then, 900 ml of 0.025 M thiobarbituric acid was added and the mixture was incubated for 65 min at 37 °C. Finally, 400 ml of 10 mM PBS was added. The fluorescence of MDA was recorded using a 520/549 (excitation/emission) filter. A calibration curve with MDA in the range 0.05–5 mM was used to calculate the MDA concentration. The results are expressed as nmoles of MDA/mg protein. 2.8. Surface chemical analysis through FTIR FTIR analysis was carried out to study the participation of surface groups in the interaction with ZnO NPs, which help to determine the extent of cell damage induced (Sadiq et al., 2011). After 72 h of exposure to ZnO NPs 300 mg/L, algal cells from 5 ml of suspension were harvested by centrifugation for 15 min at 10,000 rpm at an ambient temperature. The pellets were washed thrice with PBS and dried for 30 min at 3571 °C in a hot air oven. The dried cells were subjected to FTIR analysis by a potassium bromide technique using a Perkin-Elmer Spectrum.
Fig. 1. Image of dissolved Zn2 þ concentration (mg/L).
2.9. Microscopic analysis The samples treated with 300 mg/L ZnO NPs were collected after 72 h of interaction. A 250 ml of the sample was mixed with 250 ml of PI and incubated for 5 min in the dark at 25 °C followed by centrifugation at 200 rpm for 2 min at 4 °C. The pellet was washed thrice using SSC (saline sodium citrate) buffer to remove any unbound dye. About 10 ml of the resultant sample was carried onto a glass slide and sealed with cover slip for observation. These were observed using LSM 5 LIVE Duo Scan Confocal MicroscopeCarl ZEISS. 2.10. Field emission scanning electron microscopy Field emission scanning electron microscope (FESEM-Supra 55, -Carl Zeiss Germany) was used to observe the morphology of the algae and the particles. Both treated and untreated samples were drop coated on a 1 cm2 glass piece and air dried. The samples were subjected to gold sputtering and analyzed under FESEM. 2.11. Statistical analysis The experiments were carried out in replicates and the data are presented as mean 7standard error (SE). Significant differences between means were identified using one-way ANOVA, SPSS 20. Statistical significance was evaluated using significance levels at 0.05.
3. Results and discussion 3.1. Morphological characterization of ZnO NPs The XRD pattern (Fig. S1) Of ZnO NPs, which agrees with a database standard previously reported (JCPDS no. 89-1397), confirmed that the resulting product was a crystalline hexagonal – phase ZnO. The FESEM micrographs (Fig. S2A) show ZnO NPs that are spherical in shape and the size is in the range of 40–48 nm. EDAX analysis gives the additional evidence to the presence of elemental zinc (Fig. S2B). 3.2. Dissolution of ZnO NPs The dissolution rate of NPs is influenced by several factors, including particle size, surface area, surface curvature, and roughness of the particle (Farre´ et al., 2009). Zn2 þ solute from NPs can enter cells by transport and ion/voltage-gated channels (Colvin et al., 2003). Zn2 þ ions have the ability to form chelates with biomolecules or dislodge the metal ions in specific
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Table 1 A) Cell viability, B) LDH, C) SOD, D) GSH, E) LPO. Exposure period
Exposure conc ZnO NPs
A) Cell viability (%)
B) LDH (nmol/minml)
1 C) SOD (l mgprot )
1 D) GSH (mg mgpro )
1 E) LPO (nmol mgpro )
24 h
50 100 200 300 50 100 200 300
90.497 0.3 85.26 7 0.9 80.29 7 0.4 72.62 7 0.3 70.347 0.2 62.81 7 0.5 35.737 0.3 23.69 7 1.8
3.187 0.05 6.95 7 0.10 14.137 0.10 16.247 0.16 6.177 0.05 13.337 0.1 20.32 7 0.2 22.98 7 0.3
30.107 0.4 25.777 0.7 18.167 0.3 11.03 7 0.07 28.86 7 0.3 14.777 0.7 10.60 7 0.7 6.59 7 0.4
39.42 7 0.30 35.16 7 0.04 27.22 7 0.09 21.11 7 0.07 31.46 7 0.06 24.11 7 0.02 19.677 0.20 16.21 7 0.10
3.85 7 0.02 3.93 7 0.02 4.17 0.01 4.147 0.02 3.90 7 0.09 4.157 0.04 4.28 7 0.04 4.917 0.08
72 h
Fig. 2. Flow cytometry images: (A) control C. vulgaris showed nearly 100% of cell viability after 72 h; (B) 50 mg/L treated C. vulgaris viable decrease of 69.90% after 72 h; (C) 100 mg/L treated C. vulgaris viable decrease of 59.31% after 72 h; (D) 200 mg/L treated C. vulgaris viable decrease of 33.86% after 72 h and (E) 300 mg/L treated C. vulgaris viable decrease of 22.37% after 72 h.
metalloproteins, which may result in functional protein inactivation. Zn2 þ released by NPs increase their local concentration and disrupts cellular metal cation homeostasis to result in cell toxicity. Zinc oxide is nearly insoluble in pure water, where its water solubility ranges from 1.6 mg/L to 5 mg/L (PROSPEcT, 2009) whereas,
in a high ionic strength and high pH medium, such as the seawater, the solubility of the zinc oxide is significantly higher showing, in addition, some differences between the nano and bulk form (Miller et al., 2010; Miao et al., 2010; Peng et al., 2011; Wong et al., 2010). In our study to identify dissolution of NPs, the
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Fig. 3. Fourier Transform Infrared spectrum of control (A) and nanoparticle interacted (B) cells of C. vulgaris.
Fig. 4. Confocal images (A) untreated cell with intact membranes, preventing entry of PI dye, resulting in unstained cells and (B) cell treated with 300 mg/L for 72 h showing PI stained cells due to compromised membrane integrity.
concentrations of the released Zn2 þ were measured at 50– 300 mg/L of ZnO NPs. The results showed that the concentrations of releasing Zn2 þ gradually increased with increasing concentrations. The concentration of Zn2 þ dissolved from 50 mg/L of NPs was 3.96 mg/L. Even at the highest concentration (300 mg/L) of NPs, 10.68 mg/L of soluble Zn2 þ was detected in the present study (Fig. 1). Peng et al. (2011) observed a complete suppression (100% effect) of three marine diatomgrowths at 10 mg/L of nano ZnO. Some researchers considered that dissolved zinc ions played an important role in the toxicity of ZnO NPs (Song et al., 2010). 3.3. Cytotoxicity ZnO NPs were strongly cytotoxic at lower concentrations and exhibited strong protein adsorption abilities (Horie et al., 2009) which may have contributed to their cytotoxicity. ZnONPs taken up by cells may cause rapid dissolution, releasing Zn2 þ ions (Xia et al., 2008), elevated intracellular Zn2 þ induced the disruption of mitochondrial function (Bishop et al., 2007; Wiseman et al., 2007) and triggered ROS generation leading to cell death (Xia et al., 2008). The cytotoxic effect of ZnO NPs on C. vulgaris with the increasing concentration (10–300 mg/L) was evaluated using MTT assay. The ZnO NPs exerted a cytotoxic effect on C. vulgaris at lower concentrations of 50 mg/L. The viability of C. vulgaris decreased by 90.49 70.3% at 24 h upon 50 mg/L ZnO NPs, whereas upon 300 mg/L, the viability of C. vulgaris significantly reduced up to
23.697 1.8% (Table 1A). A similar phenomenon reported by Sharma et al. (2009) indicated that ZnO NPs induced cytotoxicity was concentration and time dependent. Lactate dehydrogenase (LDH) assay was carried out to confirm the intactness of the cell membrane (Jeng and Swanson, 2006). Any damage in the membrane, which leads to increased levels of lactate dehydrogenase enzyme in the test solution, can be one of the reasons for cell death. Zhang et al. (2012) have considered LDH assay as one of the single parameters for determining cellular toxicity of metal oxide nanoparticles. A significant (p o0.05) LDH leakage was observed at 24 h and 72 h on exposure of 14.13 70.14 at 200 mg/L, 16.24 70.16 at 300 mg/L of 24 h and 20.32 70.26 at 200 mg/L of 24 h, 22.74 70.33 at 300 mg/L of 48 h (p o0.05) (Table 1B). 3.4. Flow cytometric analysis of cell viability To determine the extent of cytotoxicity in C. vulgaris cells, the ZnO NP-treated cells were analyzed using flow cytometry. In flow cytometry images, upper left and right quadrants show the percentage of dead cells and lower left and right quadrants show the percentage of live cells (Fig. 4). The cell viability of the control group was almost 100% (Fig. 2A) during the 72 h of culture; nevertheless, in cultures with the various concentrations of ZnO NPs the cell viability was found to have decreased after 72 h. The percentage of viable cells decreased to 68.09%, 59.31%, 33.86% and
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Fig. 5. Scanning electron micrographs: (A) control cell shows no damage; (B) (C) extensive membrane damage upon 72 h treatment with the 50 mg/L and 100 mg /L; D and E 300 mg/L treated cell shows distorted morphological features after 72 h treatment. (For interpretation of the references to color in this figure, the reader is referred to the web version of this article.)
22.37% for 50 mg/L (Fig. 2B), 100 mg/L (Fig. 2C), 200 mg/L (Fig. 2D), and 300 mg/L (Fig. 2E) concentrations, respectively. This dose and exposure time dependence corroborate with MTT assay results as well. 3.5. Oxidative stress determination The potential of ZnO NPs to induce oxidative stress was assessed by measuring the SOD in algal cells (Table 1C). An exposure time of 72 h was found to induce SOD (6.59 7 0.4) significantly in algal cells at 300 mg/L of ZnO NPs. These results show that oxidative stress is increased by ZnO NPs. A similar phenomenon was reported by Chen et al. (2012) for C. vulgaris; the activity of
antioxidant enzymes followed a changing trend of being activated by low concentrations of ZnONPs and inhibited by higher concentrations (Hao and Chen, 2012). When the cell was exposed to ZnO NPs for 72 h, GSH level significantly decreased. When the ZnO NPs dose increased to 200 mg/L, the intracellular GSH level were almost reduced by half 19.677 0.2 compared to the control (Table 1D). The data demonstrated a significant depletion of GSH level also in ZnO NPs exposed algal cells. The GSH content decreased in the high concentration of ZnONPs stress, which may be attributed to the consumption of a large amount of the GSH in a variety of detoxification processes. At high concentrations of ZnO NPs destruction of antioxidation structures and a decrease in the level of antioxidants probably occur, which result in rapid
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accumulation of ROS. Lipid peroxidation is an indicator of oxidative damage to cell membrane lipids and has been used extensively as a biomarker for oxidative stress in vivo (Sayeed et al., 2003). It is estimated by measuring the MDA content of cells.The MDA content significantly increased at ZnO NPs concentrations higher than 200 mg/L (Table 1E). 3.6. Surface characterization by FTIR FTIR has emerged as a valuable instrument for understanding the involvement of surface charge-based attachment of NPs onto algal cells, leading to membrane damage as well as aggregate formation. Control spectra of the uninteracted cell (not treated with ZnO NPs) displayed a number of peaks (Fig. 3A). There was a variation in the intensity of bands in test samples (treated with ZnO NPs) (Fig. 3B). A control spectrum of uninteracted cell shows peak at 3390 cm 1 representing the presence of hydrogen-bonded O–H stretch. The peak at 2929.73 cm 1 and 699.70 cm 1 represents the presence of C–H stretch. The peaks at 1609.88 cm 1 and 933.33 cm 1 took place at the stretching vibration of N–H primary amine. The peaks at 1261 cm 1 and 1120 cm 1 were the stretching vibration of C–O. The treated cells with ZnO NPs showed a peak at 3334.33 cm 1 representing the presence of hydrogen-bonded O–H stretch. The surfaces of algae contain a number of functional groups with high affinity for metal ions and carry a net negative charge, due mainly to carboxylic, sulfhydryl, and phosphatic groups (Crist et al., 1990, 1994). These groups are binding sites that transport metal ions across the cell membrane and into the cell. Various studies have demonstrated that two distinct phases occur in the ingestion of metals by algal cells: a rapid assimilation that is terminated within 10 min followed by a slower, facilitated transport into the cytoplasm of the cell (Gonza ´lez Da´vila et al., 1995; Xue et al., 1988). Adsorption of metal ions to the cell surface determines the initial toxicant loading of the cells, and this is likely to be dependent on the algal cell surface area and also the nature of the binding sites (Geisweid and Urbach,1983). The peak at 1606.06 cm 1shows the stretching vibration of N–H primary amine. The peak at 1259.82 cm 1 was the stretching vibration of C–O. The peak at 542.07 cm 1 was associated with the stretching vibration of ZnO (Wahab et al., 2008). The conformation of surface chemical interaction led us to probe further for membrane disruption followed by attachment. 3.7. Microscopic analysis The compromised cellular integrity and permeability was confirmed through confocal laser scanning microscopy which allowed the visualization of stained nuclei (a nuclear-specific dye PI was used) in case of damaged cells upon ZnO NP exposure. No PI fluorescence was observed in control cells after 72 h (Fig. 4A). PI fluorescence was detected in the nucleus after 72 h (Fig. 4B). The membrane damage as a sign of cell death has also been reported by Metzler et al. (2011) in Pseudokirchneriella subcapitata upon TiO2 nanoparticle interaction.The fluorescence microscopic observation corroborated well with FESEM observation. The FESEM micrograph showed intact structural features in untreated cells and the size of the untreated cell was 3.41 mm (Fig. 5A). In contrast, the treated cell reconfirmed the cell wall (marked with red pointer) upon exposure to 50 mg/L (Fig. 5B) and 100 mg/L (Fig. 5C) ZnO NPs. In addition, a direct visualization at high resolution also revealed morphological alterations of cell structure damage with 200 mg/L and 300 mg/L (Fig. 5D and E) after 72 h and the size of the treated cell was 1.83 mm. Membrane damage as a sign of cell death has also been reported by Pakrashi et al. (2013).
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4. Conclusions To our knowledge, this is one of the first focused, systematic studies on the effect of ZnO NPs on marine microalgae C. vulgaris and first in describing through flowcytometry, cytotoxicity, oxidative stress mechanism and morphology. The results showed that C. vulgaris viability was affected by an increase of ZnO NPs concentrations. ZnO NPs were found to cause a substantial outcome on the MDA, GSH, SOD, and LDH. FTIR study of the surface chemistry of the interacted cells demonstrates probable interaction between the particles and the surface sites on the cell wall. The result showed the interaction between the ZnO NPs and algae favoring the formation of distorted morphological features after 72 h treatment. The changes observed in the activity of SOD, GSH, LPO, LDH and decrease of viable cells, thus established an end point for ZnO NPs toxicity.
Acknowledgments Authors are thankful to IFS (Sweden) (No. A/5041-1), for the financial support extended; Centre for Cellular and Molecular Platforms, NCBS, Bangalore, for their help in analyzing the sample by Flow Cytometry and Confocal Microscopy; and Centre for Nanoscience and Nanotechnology, Sathyabama University, Jeppiaar Nagar, Chennai in carrying out FESEM, XRD, and EDX.
Appendix A. Supplementary material Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.ecoenv.2014.11.015.
References Aschberger, K., Micheletti, C., Sokull-Kl ̈ uttgen, B., Christensen, F.M., 2011. Analysis of currently available data for characterizing the risk of engineered nanomaterials to the environment and human health lessons learned from four case studies. Environ. Int. 37, 1143–1156. Bishop, G.M., Dringen, R., Robinson, S.R., 2007. Zinc stimulates the production of toxic, reactive oxygen species (ROS) and inhibits glutathione reductase in astrocytes. Free Radic. Biol. Med. 42, 1222–1230. Blaise, C., Gagne´, F., Ferard, J.F., 2008. Ecotoxicity of selected nanomaterials to aquatic organisms. Environ. Toxicol. 23, 591–598. Brayner, R., Ferrari-Iliou, R., Brivois, N., Djediat, S., Benedetti, M.F., Fievet, F., 2006. Toxicological impact studies based on Escherichia coli bacteria in ultrafine ZnO nanoparticles colloidal medium. Nano Lett. 6, 866–870. Brown, D.M., Donaldson, K., Borm, P.J., Schins, R.P., Dehnhardt, M., Gilmour, P., 2004. Calcium and ROS-mediated activation of transcription factors and TNF alpha cytokine gene expression in macrophages exposed to ultrafine particles. Am. J. Physiol. – Lung Cell. Mol. Physiol. 286, 344–353. Chen, X., Zhu, X., Li, R., Yao, H., Lu, Z., Yang, X., 2012. Photosynthetic toxicity and oxidative damage induced by nano-Fe3O4 on Chlorella vulgaris in the aquatic environment. Open J. Ecol. 1, 21–28. Colvin, R.A., Fontaine, C.P., Laskowski, M., Thomas, D., 2003. Zn2 þ transporters and Zn2þ homeostasis in neurons. Eur. J. Pharmacol. 479, 171–185. Crist, R.H., Martin, J.R., Guptill, P.W., Eslinger, J.M., Crist, D.R., 1990. Interaction of Metal Ions with Acid Sites of Biosorbents Peat Moss and Vaucheria and Model Substances Alginic and Humic Acids. Environ Sci Technol 24, 337–342. Crist, R.H., Martin, J.R., Carr, D., Watson, J.R., Clarke, H.J., Crist, D.R., 1994. Interaction of metals and protons with algae. 4. Ion-exchange vs adsorption models and a reassessment of Scatchard plots; ion-exchange rates and equilibria compared with calcium alginate. Environ. Sci. Technol. 28, 1859–1866. De Berardis, B., Civitelli, G., Condello, M., Lists, P., Pozzi, R., Arancia, G., Meschini, S., 2010. Exposure to ZnO nanoparticles induces oxidative stress and cytotoxicity in human colon carcinoma cells. Toxicol. Appl. Pharmacol. 246, 116–127. Del Rio, D., Pellegrini, N., Colombi, B., Bianchi, M., Serafini, M., Torta, F., Tegoni, M., Musci, M., Brighenti, F., 2003. Rapid fluorimetric method to detect total plasma malondialdehyde with mild derivatization conditions. Clin. Chem. 49, 690–692. Ellman, G.L., 1959. Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82, 70–77. Elstner, E.F., Heupel, A., 1976. Inhibition of nitrite formation from hydroxylammoniumchloride: a simple assay for superoxide dismutase. Anal. Biochem. 70, 616–620.
30
T.Y. Suman et al. / Ecotoxicology and Environmental Safety 113 (2015) 23–30
Farre´, M., Gajda-Schrantz, K., Kantiani, L., Barcelo, D., 2009. Ecotoxicity and analysis of nanomaterials in the aquatic environment. Anal. Bioanal. Chem. 393, 81–95. Franklin, N.M., Rogers, N.J., Apte, S.C., Batley, G.E., Gadd, G.E., 2007. Comparative toxicity of nanoparticulate ZnO, bulk ZnO, and ZnCl2 to a freshwater microalga (Pseudokirchneriella subcapitata): the importance of particle solubility. Environ. Sci. Technol. 41, 8484–8490. Geisweid, H.J., Urbach, W., 1983. Sorption of cadmium by the green microalgae Chlorella vulgaris, Ankistrodesmus braunii and Eremosphaera viridis. Z. Planzenphysiol. 109, 127–141. Gonza´lez Da´vila, M., Santana-Casiano, J.M., Pe´rez-Pen~a, J., Millero, F.J., 1995. Binding of Cu (II) to the surface and exudates of the alga Dunaliella tertiolecta in seawater. Environ. Sci. Technol. 29, 289–301. Gottschalk, F., Sonderer, T., Scholz, R.W., Nowack, B., 2009. Modeled environmental concentrations of engineered nanomaterials (TiO2, ZnO, Ag, CNT, fullerenes) for different regions. Environ. Sci. Technol. 43, 9216–9222. Hao, L., Chen, L., 2012. Oxidative stress responses in different organs of carp (Cyprinus carpio) with exposure to ZnO nanoparticles. Ecotoxicol. Environ. Saf. 80, 103–110. Heinlaan, M., Ivask, A., Blinova, I., Dubourguier, H., 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. Horie, M., Nishio, K., Fujita, K., Endoh, S., Miyauchi, A., Saito, Y., Iwahashi, H., Yamamoto, K., Murayama, H., Nakano, H., 2009. Protein adsorption of ultrafine metal oxide and its influence on cytotoxicity toward cultured cells. Chem. Res. Toxicol. 22, 543–553. Jeng, H.A., Swanson, J., 2006. Toxicity of metal oxide nanoparticles in mammalian cells. J. Environ. Sci. Health A Toxic Hazard. Subst. Environ. Eng. 41, 2699–2711. Jiang, W., Mashayekhi, H., Xing, B., 2009. Bacterial toxicity comparison between nano- and micro-scaled oxide particles. Environ. Pollut. 157, 1619–1625. Ji, J., Wu, Z., Liu, Q., Zhang, Y., Ye, M., Li, M., 1991. An ultramicro analytic and rapid method for determination of superoxide dismutase activity. J. Nanjing Railw. Med. Coll. 10, 27–30. Lin, D.H., Xing, B.S., 2007. Phytotoxicity of nanoparticles: inhibition of seed germination and root growth. Environ. Pollut. 150, 243–250. Manzo, S., Miglietta, M.L., Rametta, G., Buono, S., Di Francia, G., 2013. Toxic effects of ZnO nanoparticles towards marine algae Dunaliella tertiolecta. Sci. Total Environ. 445, 371–376. Miao, A.J., Zhang, X.Y., Luo, Z., Chen, C.S., Chin, W.C., Santschi, P.H., 2010. Zinc oxide engineered nanoparticles: dissolution and toxicity to marine phytoplankton. Environ. Toxicol. Chem. 29, 2814–2822. Miller, R.J., Lenihan, H.S., Muller, E.B., Tseng., N., Hanna, S.K., Keller, A.A., 2010. Impact of metal oxide nanoparticles on marine phytoplankton. Environ. Sci. Technol. 44, 7329–7334. Metzler, D.M., Li, M., Erdem, A., Huang, C., 2011. Responses of algae to photocatalytic nano-TiO2 particles with an emphasis on the effect of particle size. Chem. Eng. J. 170 (2–3), 538–546. Mosmann, T., 1983. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. Immunol. Methods 65, 55. Nair, S., Sasidharan, A., Rani, V.V.D., Menon, D., Nair, S., Manzoor, K., Raina, S., 2009. Role of size scale of ZnO nanoparticles and microparticles of toxicity toward bacteria and osteoblast cancer cells. J. Mater. Sci. Mater. Med. 20, 235–241. Ormerod, M.G., 1990. Analysis of DNA. General methods. In: Ormerod, M.G. (Ed.), Flow Cytometry. A Practical Approach. Oxford University Press, Oxford, pp. 69–87. Osmond, M.J., McCall, M.J., 2010. Zinc oxide nanoparticles in modern sunscreens: an analysis of potential exposure and hazard. Nanotoxicology 4, 15–41. Pakrashi, S., Dalai, S., Prathna, T.C., Trivedi, S., Myneni, R., Raichur, A.M., Chandrasekaran, N., Mukherjee, A., 2013. Cytotoxicity of aluminium oxide nanoparticles towards freshwater algal isolate at low exposure concentrations. Aquat. Toxicol. 133, 34–45. Peng, X., Palma, S., Fisher, N.S., Wong, S.S., 2011. Effect of morphology of ZnO nanostructures on their toxicity to marine algae. Aquat. Toxicol. 102, 186–196. PROSPEcT, 2009. Ecotoxicology test protocols for representative nanomaterials in support of the OECD sponsorship programme. 8. Premanathan, M., Karthikeyan, K., Jeyasubramanian, K., Manivannan, G., 2011. Selective toxicity of ZnO nanoparticles toward gram-positive bacteria and cancer
cells by apoptosis through lipid peroxidation. Nanomed. Nanotechnol. Biol. Med. 7, 184–192. Reddy, K.M., Feris, K., Bell, J., Wingett, D.G., Hanley, C., Punnoose, A., 2007. Selective toxicity of zinc oxide nanoparticles to prokaryotic and eukaryotic systems. Appl. Phys. Lett. 90, 2139021–2139023. Sayeed, I., Parvez, S., Pandey, S., Bin-Hafeez, B., Haque, R., Raisuddin, S., 2003. Oxidative stress biomarkers of exposure to deltamethrin in freshwater fish, Channa punctatus Bloch. Ecotoxicol. Environ. Saf. 56, 295–301. Sadiq, I.M., Pakrashi, S., Chandrasekaran, N., Mukherjee, A., 2011. Studies on the toxicity of aluminum oxide (Al2O3) nanoparticles to microalgae species Scenedesmus Sp. and Chlorella Sp. J. Nanopart. Res. 13, 3287–3299. Sharifi, S., Behzadi, S., Laurent, S., Laird Forrest, M., Stroeve, P., Mahmoudi, M., 2012. Toxicity of nanomaterials. Chem. Soc. Rev. 41, 2323–2343. Sharma, V., Shukla, R.K., Saxena, N., Parmar, D., Das, M., Dhawan, A., 2009. DNA damaging potential of zinc oxide nanoparticles in human epidermal cells. Toxicol. Lett. 185, 211–218. Song, W., Zhang, J., Guo, J., Zhang, J., Ding, F., Li, L., Sun, Z., 2010. Role of the dissolved zinc ion and reactive oxygen species in cytotoxicity of ZnO nanoparticles. Toxicol. Lett. 15, 389–397. Song, Y., Zhu, L., Wang, S., Wang, J., Liu, J.H., Xie, H., W., 2008. DNA damage and effects on antioxidative enzymes in earthworm (Eisenia fetida) induced by atrazine. Soil Biol. Biochem. 41, 905–909. Wang, B., Feng, W.Y., Wang, M., Wang, T.C., G.U., Y.Q., Zhu, M.T., Ouyang, H., Shi, J.W., Zhang, F., Zhao, Y.L., Chai, Z.F., Wang, H.F., Wang, J., 2008a. Acute toxicological impact of nano- and submicro-scaled zinc oxide powder on healthy adult mice. J. Nanopart. Res. 10, 263–276. Wahab, R., Ansari, S.G., Kim, Y.S., Dar, M.A., Shin, H.S., 2008. Synthesis and characterization of hydrozincite and its conversion into zinc oxide nanoparticles. J. Alloys Compd. 461, 66–71. Weckx, J., Clijsters, H., 1996. Oxidative damage and defense mechanisms in primary leaves of Phaseolus vulgaris as a result of root assimilation of toxic amounts of copper. Physiol. Plant. 96, 506–512. Wiseman, D.A., Wells, S.M., Hubbard, M., Welker, J.E., Black, S.M., 2007. Alterations in zinc homeostasis underlie endothelial cell death induced by oxidative stress from acute exposure to hydrogen peroxide. Am J Physiol Lung Cell Mol Physiol 292, 165–177. Wiench, K., Wohlleben, W., Hisgen, V., Radke, K., Salinas, E., Zok, S., Landsiedel, R., 2009. Acute and chronic effects of nano and non-nano-scaleTiO2 and ZnO particles on mobility and reproduction of the fresh water invertebrate Daphnia magna. Chemosphere 76, 1356–1365. Wong, S.W., Leung, P.T.Y., Djurisić, A.B., Leung, K.M., 2010. Toxicities of nano zinc oxide to five marine organisms: influences of aggregate size and ion solubility. Anal. Bioanal. Chem. 396, 609–618. Xia, T., Kovochich, M., Liong, M., Madler, L., Gilbert, B., Shi, H., Yeh, J.I., Zink, J.I., Nel, A.E., 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. Xiong, D.W., Fang, T., Yu, L.P., Sima, X.F., Zhu, W.T., 2011. Effectsofnano-scaleTiO2, ZnO and their bulk counterparts on zebrafish: acute toxicity, oxidative stress and oxidative damage. Sci.Total Environ. 409, 1444–1452. Xue, H., Stumm, W., Sigg, L., 1988. The binding of heavy metals to algal surfaces. Water Res. 22, 917–926. Zhang, H., Ji, Z., Xia, T., Meng, H., Low-Kam, C.R., Liu, S., Pokhrel, S., Lin, X., Wang, Y. P., Liao, M., Wang, L., Li, R., Rallo, R., Damoiseaux, D., Telesca, L., Mädler, Y., Cohen, J.I., Zink, A., Nel, E., 2012. Use of metal oxide nanoparticle band gap to develop a predictive paradigm for oxidative stress and acute pulmonary inflammation. ACS Nano 6, 4349–4368. Zhao, J., Castranova., V., 2011. Toxicology of nanomaterials used in nanomedicine. J. Toxicol. Environ. Health B 14, 593–632. Zhu, X.S., Zhu, L., Duan, Z.H., Qi, R.Q., Li, Y., Lang, Y.P., 2008. Comparative toxicity of several metal oxide nanoparticle aqueous suspensions in zebrafish (Daniorerio) early developmental stage. J. Environ. Sci. Health 3 (3), 278–284. Zhu, X.S., Zhu, L., Chen, Y.S., Tian, S.Y., 2009. Acute toxicities of six manufactured nanomaterial suspensions to Daphnia magna. J. Nanopart. Res. 11, 67–75.
