Nano-titanium dioxide induces genotoxicity and apoptosis in human lung cancer cell line,A549

Abstract

The increased inhaled application of titanium dioxide nanoparticles (TiO₂ NPs) increases the potential pulmonary health risks. The present investigations were carried out to study the TiO₂ NPs-induced apoptosis, oxidative stress and genotoxicity in the human lung cancer cell line, A549, a widely used cell system for pulmonary toxicity studies. Tetrazolium bromide salt and lactate dehydrogenase release assays were used to study the cytotoxicity. The genotoxicity studies were carried out using cytokinesis block micronucleus assay. Apoptosis was confirmed by the formation of apoptotic bodies and altered expression (messenger RNA (mRNA) and protein) of markers such as P⁵³, P²¹, Bax, Bcl₂ and cleaved caspase-3. Cells exposed to TiO₂ NPs (10 and 50 μg/ml) for 6–24 h shows significant induction in oxidative stress, that is, the production of reactive oxygen species and malondialdehyde and decrease in the activity of catalase and glutathione. TiO₂ NPs exposure also induces the formation of apoptotic bodies and micronucleus as marker of genotoxicity. A significant up-regulation in the expression of apoptosis markers such as P⁵³, P²¹ and cleaved caspase-3 was observed, while the levels were down-regulated for Bcl₂ at both mRNA and protein levels. TiO₂ NPs exposure could not pose significant effects on Bax expression. Data indicate that nano-TiO₂ induces oxidative stress, genotoxicity and apoptosis in human lung cancer cell line, A549. Our result also identifies the mechanisms involved in TiO₂ NP-induced changes in A549 cells. Perhaps, reporting for the first time, the association of TiO₂ NPs-induced genotoxicity and apoptosis at transcriptional and translational level in the human lung cancer cell line, A549 cells.

Keywords

A549 apoptosis genotoxicity oxidative stress

Introduction

Nanotization of particles may enhance adverse toxic responses upon the biological interactions.¹ Release of such nanoparticles (NPs) in the environment is known to induce damaging effect in the respiratory tract² and lungs.³ Among the possible exposure routes, inhalation and dermal exposure are considered as most important for NPs. Exposure of NPs through inhalation and translocation from lung to other organs has been documented.^4,5 In general, interaction of NPs with cell membrane is known to generate reactive oxygen species (ROS) and lipid peroxidation (LPO), which subsequently leads the DNA lesion, cellular damages, apoptosis and finally cell death.^6

–10 Agglomeration/aggregation state of NPs also affects the kinetics of cellular binding/uptake and the ability to induce cytotoxic responses.¹¹ Physicochemical characterization is a critical factor for correlating the NPs characteristics with any measured biological/toxicological outcome.^12,13 It is demonstrated recently that the physicochemical properties (such as size, shape, surface area, phase and composition) of titanium dioxide (TiO₂) and zinc oxide NPs play a crucial role in their toxicity.¹⁴ Although these unique physiochemical properties of TiO₂ NPs make them to extensively use in cosmetic products, paint industries, dusting powder, varnishes, enamels, lacquers and paper coatings, their exposure at occupation setups and consumers end is well reported.¹⁵ TiO₂ NPs were also found to induce oxidative stress and apoptosis under both in vivo ^16,17 and in vitro ^18,19 conditions in cells of human and animal origin. This may leads to sequential events of signal transduction that can induce the accumulation of P53, a tumor suppressor protein, which block the cellular proliferation by inducing cell cycle arrest and apoptosis in response to cellular stress such as ROS. Hussain et al.²⁰ investigated the molecular pathway of apoptosis induced by carbon black and TiO₂ NPs in bronchial epithelial cells by biochemical estimation and immunofluorescence staining of apoptosis marker proteins. However, the altered expression (messenger RNA (mRNA) and protein level) of markers of genotoxicity and apoptosis at transcriptional (mRNA) and translational (protein) levels in cells of respiratory system, lung exposed to TiO₂ NPs has not been explored so far. Thus, the present investigations were carried out to study TiO₂ NPs-induced alterations in the expression (mRNA and protein) of selected markers of apoptosis and their association with oxidative stress and genotoxicity in human lung cancer cells, A549, a widely used cell line for inhalation/pulmonary toxicity studies.

Methods

Reagents and consumables

All the specified chemicals, reagents and diagnostic kits were purchased from Sigma Chemical Company Pvt. Ltd (St. Louis, Missouri, USA) unless otherwise stated.

Cell culture, particle characterization and exposure

Human lung cancer cell line, A549, used in the study was originally procured from National Centre for Cell Sciences (Pune, India) and since then it has been maintained at In Vitro Toxicology Laboratory, Indian Institute of Toxicology Research (Lucknow, India) following the standard protocol.²¹ Prior to using in the experiments, cells were assessed for viability and batches showing more than 95% viability were only used in the study. TiO₂ NPs (d < 25 nm, specific surface area 200–220 m²/g) without any coating were purchased from Sigma Aldrich (St. Louis, Missouri, USA, Cat no. 637254). Sterilization of the NPs were carried out by heating at 120°C for 2 h and then suspended in complete DMEM-F12 medium.²¹ Stock solution of TiO₂ NPs (1 mg/ml) was prepared in complete DMEM-F12 medium containing 10% serum and then sonicated intermittently for 5 min with each stretch of 1 min at 100 Hz. The experimental concentrations were prepared by diluting the stock solution in the same medium. Size distribution and zeta potential (ζ) of TiO₂ were determined using dynamic light scattering and phase analysis light scattering in a Zetasizer Nano-ZS, Model ZEN3600 equipped with 4.0 mW, 633 nm laser (Malvern instruments Ltd., Worcestershire, UK). The well-adhered monolayer of cells was exposed to TiO₂ NPs (1–100 μg/ml) for 6–48 h as per the demand of the experiments.

Cytotoxicity assessment

MTT assay

Percentage cell viability was assessed using the 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyl tetrazolium bromide (MTT) assay as described earlier.²² Briefly, cells (1 × 10⁴) were allowed to adhere for 24 h under high humid environment in 5% CO₂ at 37°C in 96-well culture plates. After the respective exposure, MTT (5 mg/ml of stock in phosphate buffered saline (PBS)) was added (10 μl/well in 100 μl of cell suspension), and plates were incubated for 4 h. At the end of incubation period, the reaction mixture was carefully taken out and 200 μl of dimethyl sulfoxide was added to each well by pippeting up and down several times until the contents were mixed well. The plates were kept on rocker shaker for 10 min at room temperature and then read at 550 nm using multiwell microplate Reader (Synergy HT, Bio-Tek, USA). Untreated sets were run under identical conditions and served as basal control. To study the effect of TiO2 NPs with color absorbance, with this assay another set with medium containing only TiO₂ NPs were also run in similar condition.

Lactate dehydrogenase (LDH) release assay

LDH release is a method to measure the membrane integrity as a function of the amount of cytoplasmic LDH released from damaged cells to the medium. LDH assay was carried out using the readymade commercially available kit for in vitro cytotoxicity evaluation (LDH Assay Kit, BioVision, California, USA). In brief, following the exposure, plates were incubated as per the experimental schedule in CO₂ incubator and centrifuged at 250×g for 4 min. Then supernatant of each well was transferred to a fresh flat bottom 96-well culture plate and further processed for enzymatic analysis as per the instructions given in the kit. Significant, though limited, cytotoxic doses of TiO₂ NPs from these studies were used in further experimentations.

Transmission electron microscopy (TEM)

The ultra structural characterization including texture, size and internalization was done by Prof. Margareta Lantow, at Institute of Biological Sciences, University of Rostock, Germany. Monolayer of A549 cells were exposed to TiO₂ NPs (1, 10 and 50 μg/ml) for 6–48 h. The cells were then fixed in 4% glutaraldehyde for minimum 10 min and processed further following the protocol of Rahman et al.²¹ Following ultrathin sectioning using LKB-ultratom III (LKB Instrument, Rockville, Maryland, USA), cells were visualized with TEM (EM 902A; Zeiss, Oberkochen, Germany).

Oxidative stress studies

ROS generation

TiO₂ NPs-induced ROS generation was assessed in A549 cells using 2′,7′-diclorodihydrofluorescein di-acetate (DCFH-DA, Sigma Aldrich, Missouri, USA) dye as fluorescence agent. ROS generation was studied by two methods as described earlier.²¹ First was the fluoremetric analysis and second was the microscopic fluorescence imaging. For fluoremetric analysis, cells (1 × 10⁴ per well) were seeded in 96-well black bottom culture plates and allowed to adhere them for 24 h in CO₂ incubator at 37°C. The medium was then aspirated and cells were exposed to TiO₂ NPs (1–50 µg/ml) for 6, 12 and 24 h. On the completion of respective exposure periods, cells were incubated with DCFH-DA (10 µM) for 30 min at 37°C. The reaction mixture was then aspirated and replaced by 200 µl of PBS in each well. The plates were kept on rocker shaker for 10 min at room temperature in the dark. Fluorescence intensity was measured using multiwell microplate reader (Synergy HT, Bio-Tek, USA) on excitation wavelength at 485 nm and emission wavelength at 528 nm. Parallel set of cells (5 × 10⁴ per well of tissue culture slide flasks) were analyzed for intracellular fluorescence using upright fluorescence microscope equipped with CCD cool camera (Nikon Eclipse 80i equipped with Nikon DS-Ri1 12.7 megapixel camera). The data were expressed as percentage of the unexposed control. Hydrogen peroxide (H₂O₂, 500 µM) was used as a positive control 1 h prior to completion of exposure time.

Glutathione (GSH) levels

GSH) levels were assessed using commercially available kit (ApoGSH Glutathione Colorimetric Assay Kit, Catalog No#K261-100, BioVision, California, USA). In brief, the deproteinated samples were transferred to 96-well plates (50 µl/well), mixed with 150 µl freshly prepared assay cocktail and read at 405 nm using Multiwell microplate Reader at 5 min intervals up to 30 min. Standard curve was plotted using the GSH standard supplied in the kit and was used to calculate the experimental values.

LPO

Following the exposure of TiO₂ NPs (1–50 µg/ml) for 6, 12 and 24 h, cells were harvested by centrifugation at 1000 r/min for 10 min and processed for the estimation of LPO using commercially available kit (Lipid Peroxidation Assay Kit, catalog no. 705002; Cayman Chemicals, USA) as per the manufacturer’s protocol.

Catalase (CAT) levels

The experimental setup for CAT activity was similar to the LPO. Activity was measured using commercially available kit for CAT activity (Catalog no. 707002; Cayman Chemicals, USA) following the protocol provided by manufacturer.

Genotoxicity studies

Cytokinesis block micronucleus assay

The micronucleus (MN) assay performed as described earlier.²¹ In brief, the cells were exposed to TiO₂ NPs (1–50 µg/ml) for 24 h. Medium was aspirated and cells were re-incubated for 22 h in medium supplemented with cytochalasin B (3 mg/ml, Sigma Aldrich, Missouri, USA) to accomplish the nuclear division (doubling time for A549 cell line is 22 h). Cells were processed further to prepare the slide for microscopic examination of MN induction. Ethyl methanesulfonate (6 mM; CasNo.62-50-0) was used as a positive control. Minimum of 1000 bi-nucleated cells with well-defined cytoplasm were scored for MN detection.

Apoptosis studies

Bis-benzimide staining

The number of apoptotic body formation and nuclear condensation was observed in A549 cells receiving exposure of TiO₂ NPs (1–50 μg/ml) for 24 and 48 h. Hochest dye No. 33342 was used for the purpose as per the standard protocol.²³

Transcriptional changes in the apoptosis markers

The unexposed control and cells exposed to TiO₂ NPs (1–50 µg/ml) for 6, 12 and 24 h were processed for total RNA isolation using Trizol reagent (Invitrogen, USA). The high capacity superscripts complementary DNA (cDNA) synthesis Kit (Applied Biosystem, USA) was used to reverse-transcribe the mRNA (2 µg) into cDNA. Random hexamer primers were used to convert total RNA into cDNA. The primer sequences used for amplification were: p⁵³: forward 5′-gaagacccaggtccagatga-3′, reverse 5′-ctccgtcatgtgctgtgact-3; p²¹: forward 5′-ggaaggaaggctggaag-3′, reverse 5′-aaccacagtgacggtacaaa-3′; Bax: forward 5′-aaagctagcgagtgtctcaagcgc-3′, reverse 5′-cccgccacaaagatggtcacg-3′; Bcl₂: forward 5′-cgacgacttctcccgccgctaccgc-3′, reverse 5′-ccgcatgctggggccgtacagttcc-3′; Caspase-3: forward 5′-tttgtttgtgtgcttctgagcc-3′; reverse 5′-attctgttgccacctttcgg-3′ and glyceraldehyde 3-phosphate dehydrogenase (GAPDH): forward 5′-catgaccacagtccatgccatcact-3′, reverse 5′-tgaggtccaccaccctgttgctgta-3′. PCR was carried out using the Bio-Rad thermal cycler machine following the thermal cycling steps: 94°C for 5 min in the first step, 94°C for 30s, 59.1°C for 30s and 72°C for 30s for 37 times in the second step and final extension at 72°C for 5 min in the third step (conditions for p⁵³ gene). These thermal cycling conditions were similar for all the genes except for annealing temperature, which was 54.4°C for p²¹, 60.7°C for Bax, 69.2°C for Bcl₂, 60°C for caspase-3 and 57°C for GAPDH. PCR products were run on 2% agarose gels. The semi-quantitative analysis for up- or down-regulation in the expression was done using Gel Documentation System (Alpha Innotech, USA) with the help of AlphaEase^TM FC StandAlone V.4.0 software.

Translational changes in apoptosis markers

Western immunoblotting was done to assess the TiO₂ NPs-induced translational changes in the expression of selected markers associated with apoptosis such as P⁵³, P²¹, Bax, BCL₂ and activated cleaved caspase-3. Following exposure of TiO₂ NPs (1–50 µg/ml) for 12, 24 and 48 h, cells were pelleted and lysed using CelLytic^TM M Cell Lysis Reagent (Cat No# C2978, Sigma Aldrich, Missouri, USA) in the presence of protein inhibitor cocktail (Cat No# P8340-5ML, Sigma Aldrich, Missouri, USA). Protein estimation was done using BCA Protein Assay Kit (Catalogue # G1002, Lamda Biotech, Inc., St. Louis, Missouri, USA). Then denatured proteins (50 µg/well) were loaded and electrophoresed using 10% Tricine-SDS gel. Proteins were transferred on Immobilon-P membrane (Millipore Cat No. IPVH00010, USA) at 180 mA current for 3 h. Nonspecific binding was blocked with 5% nonfat dry milk powder in TBST (20 mM Tris-HCl (pH 7.4), 137 mM NaCl and 0.1% Tween 20) for 2 h at 37°C. After blocking the membranes were incubated overnight at 4°C with anti-protein primary antibodies specific for P⁵³ (1:1000, Chemicon, USA), P²¹(1:1000, Chemicon, USA), BCL₂ (1:1000, CST, USA), Bax (1:500, Santa Cruz, USA), cleaved caspase-3 (1:1000, CST, USA) and GAPDH (1:1000, Chemicon, USA) in blocking buffer (pH 7.5). The membrane was then incubated for 2 h at room temperature with secondary anti-primary antibody conjugated with horseradish peroxidase (Calbiochem, USA). Then the blots were developed using (3-3′-5-5′ tetramethyl benzidine) TMB-H₂O₂ (Sigma Aldrich, Missouri, USA) and densitometry for protein specific bands was done in Gel Documentation System (Alpha Innotech, USA) with the help of AlphaEase^TM FC StandAlone V.4.0 software.

Statistical analysis

Results were expressed as mean ± standard error of mean (SEM) for the values obtained from at least three independent experiments. Statistical analysis was performed by one-way analysis of variance (ANOVA) and post hoc Dunnett test to compare the control versus treated group by SPSS Version 12. In all the cases, p < 0.01 was considered as significant

Results

Characterization studies

The mean hydrodynamic diameter and zeta potential (ζ) of the TiO₂ NPs suspension in complete medium as determined by dynamic light scattering (DLS) measurement was 434.1 nm and −7.83 mV, respectively (Figure 1(a) and (b)). The internalization of TiO₂ NPs in A549 cells through TEM analysis was found to be dose dependent. The particles adhered on the cell surface when incubated for 30 min and subsequently internalized in small vacuoles at cortical cytoplasm in extending incubations and reached to deep cell center near the mitochondria and Golgi apparatus in larger vacuoles over 48 h of exposure (Figure 1(c)).

Figure 1.

Characterization of TiO₂ NPs. (a) Size distribution and (b) zeta potential of TiO₂ NPs was determined using dynamic light scattering and PALS in a Zetasizer Nano-ZS, Model ZEN3600. (c) Representative TEM microphotographs of cell showing internalization of TiO₂ NPs. Arrow indicates the internalization of TiO₂ NPs in cytoplasm and reached to deep cell center near the mitochondria of A549 cells. PALS: phase analysis light scattering; TEM: transmission electron microscopy; TiO₂ NPs: titanium dioxide nanoparticles.

Cytotoxicity

The results of cytotoxicity assays (MTT and LDH) are summarized in Figure 2(a) and (b). In general, concentration and time-dependent decrease in percentage cell viability was observed following 6–48 h exposure of TiO₂ NPs. MTT assay demonstrated significant (p < 0.01) reduction in percentage cell viability after 24 h exposure of TiO₂ NPs at concentrations 10, 50 and 100 μg/ml (92.19 ± 1.25%, 87.07 ± 0.55% and 84.34 ± 0.44%, respectively, Figure 2(a)). The effect became more intense during the 48 h exposure period (88.72 ± 1.39%, 82.34 ± 0.44% and 78.89 ± 0.55%, respectively, for TiO₂ NPs at concentrations 10, 50 and 100 μg/ml). The results of LDH assay were similar to MTT assay (Figure 2(b)). Exposure for 6 h was found to be effective only at higher concentration in LDH assay. There was no major interaction between colors of dyes and NPs as in cell-free system (data not shown).

Figure 2.

Identification of cell viability of TiO₂ NPs in human lung epithelial cells (A549) assessed by standard endpoints such as (a) MTT assay and (b) LDH release assay. Data are represented as mean ± SE of triplicate. *p < 0.01 indicates significance. LDH: lactate dehydrogenase; MTT: 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyl tetrazolium bromide; TiO₂ NPs: titanium dioxide nanoparticles.

Oxidative stress markers

ROS generation

The results of TiO₂ NPs-induced ROS generation are summarized in Figure 3(a) and (b). Cells show significant (p < 0.01) responsiveness against TiO₂ NPs (10 and 50 μg/ml) at each time point: at 6 h (140.05 ± 3.87% and 170.61 ± 4.45%, respectively), at 12 h (140.27 ± 3.65% and 172.57 ± 1.30%, respectively) and at 24 h (142.45 ± 2.17% and 154.68 ± 3.35%, respectively). However, TiO₂ NPs at 5 μg/ml concentration was found to have significant effect only at 12 h (117.37 ± 1.77%) and 24 h (118.38 ± 1.32%) of exposure. Highest concentration (50 μg/ml) of TiO₂ NPs induced maximum ROS generation at all the time points (Figure 3(b)). Microphotographs in Figure 3(a) are representative showing the ROS generation in the cells following 12 h of TiO₂ NP_S exposure. It is well known that the free radicals are generally short-lived species due to their high reactivity so that we did observe ROS percentage at 48 h as it was already low in 24 h compared with 6 and 12 h.

Figure 3.

(a) Representative microphotographs showing TiO₂ NPs-induced ROS generation in A549 cells. ROS generation was studied using 2,7-DCFH-DA dye. Images were snapped in Nikon phase contrast cum fluorescence microscope (model 80i). (I): Control, (II): 1.0 μg/ml, (III): 5.0 μg/ml, (IV): 10.0 μg/ml, (V): 50.0 μg/ml and (VI): positive control 500 µM H₂O₂ for 1 h. (b) Percentage change in ROS generation following 6, 12 and 24 h exposure to various concentrations of TiO₂ NPs in A549 cells assessed by micro plate reader. The positive control group comprised A549 cells treated with 500 μM of H₂O₂ (1 h). *p < 0.01 considered as significant. DCFH-DA: dichlorofluorescin diacetate; H₂O₂: hydrogen peroxide; ROS: reactive oxygen species; TiO₂ NPs: titanium dioxide nanoparticles.

Reduced GSH levels

Cell exposed to TiO₂ NPs (10 and 50 μg/ml) was found to deplete the levels of GSH significantly at all the time points: 6 h (85.18 ± 2.29% and 79.27 ± 5.25%, respectively), 12 h (86.18 ± 0.81% and 78.62 ± 1.01%, respectively) and 24 h (79.38 ± 4.28% and 74.06 ± 1.89%, respectively) compared with control (Figure 4(a)). However, lower concentration of TiO₂ NPs used (1.0 μg/ml) could not pose such severe effects.

Figure 4.

(a) Change in levels of GSH, (b) lipid peroxidation and (c) change in catalase activity in A549 cells following the exposure of TiO₂ NPs for various time periods (6–24 h) assessed by micro plate reader. Data represented are mean ± SE of three identical experiments made in three replicates. *p < 0.01 considered as significant. GSH: glutathione; TiO₂ NPs: titanium dioxide nanoparticles.

LPO

TiO₂ NPs at concentrations 10 and 50 μg/ml significantly induced LPO at an exposure time of 6 h (116.79 ± 2.74% and 139.23±4.31%, respectively), 12 h (127.26 ± 2.39% and 132.65 ± 4.41%, respectively) and 24 h (138.25 ± 4.23% and 154.76 ± 5.64%, respectively) compared with control (Figure 4(b)). The induction of LPO was also concentration and time dependent.

CAT activity

Significant reduction in the activity of CAT was observed at 10 and 50 μg/ml, which was maximum at 24 h (46.83 ± 2.11% and 34.50 ± 4.15%, respectively) followed by 6 h (87.39 ± 3.29% and 84.59 ± 6.18%, respectively) and 12 h (89.88 ± 3.23% and 82.14 ± 3.12, respectively) in the TiO₂ NPs exposed cells (Figure 4(c)).

Genotoxicity study

MN assay

Result of MN assay is shown in Figure 5(a) and (b). The significant (p < 0.01) increase in MN frequencies with TiO₂ NPs at concentrations 10 and 50 μg/ml for three independent experiments were 12.66 ± 0.33 and 17.33 ± 0.33, respectively, compared with control (5.33 ± 0.33) per 1000 binucleated cells. However, lower concentration was ineffective to significantly increase MN at 24 h.

Figure 5.

(a) Representative photograph of micronucleus (I) Control and (II) 50 μg/ml treated TiO₂ NPs after 24 h. Arrow indicates the binucleated A549 cells (I, a) and binucleated cells with micronucleus (II, a) and (b) number of micronucleus/1000 binucleated cells after the exposure of various concentrations of TiO₂ NPs for 24 h. *p < 0.01 considered as significant. TiO₂ NPs: titanium dioxide nanoparticles.

Apoptosis studies

Bis-benzimide staining

Following 48 h of TiO₂ NPs exposure to cells, a concentration-dependent increase in the apoptotic bodies was observed, which was statistically significant (p < 0.01) at 10 μg/ml (13.66 ± 0.66) and 50 μg/ml (16.66 ± 0.88); however, 50 μg/ml concentration was found to be significantly effective only at 24 h (10.33 ± 0.33) of exposure (Figure 6(a) and (b)).

Figure 6.

(a) Representative microphotographs of apoptotic bodies assessed by bis-bezimide (Hochest-33342) staining in A549 cells exposed to various concentrations of TiO₂ NPs for 48 h. (I) Control, (II) 1.0 μg/ml, (III) 10.0 μg/ml and (IV) 50.0 μg/ml. Arrow indicates the apoptotic nuclei. (b) Number of apoptotic nuclei/1000 cells after the exposure of various concentrations of TiO₂ NPs for 24 and 48 h. The positive control group comprised A549 cells pretreated (2 h) with 300 μM of t-BHP. *p < 0.01 considered as significant. t-BHP: tert-butylhydroperoxide; TiO₂ NPs: titanium dioxide nanoparticles.

Transcriptional changes

TiO₂ NPs (10 and 50 μg/ml) induces significant up-regulation of mRNA expression of p⁵³ (1.12 ±0.11 and 1.39±0.26, respectively), p²¹ (4.93 ± 0.36 and 6.14 ± 0.23, respectively) and cleaved caspase-3 (1.42 ± 0.28 and 1.96 ± 0.28, respectively) and down-regulation of Bcl₂ (0.667 ± 0.04 and 0.513±0.12, respectively) compared with control at 12 h exposure (Figure 7(a) and (b)). While Bax was not affected, no such correlation could be established for the same genes at 6 and 24 h exposure (data not shown).

Figure 7.

(a) TiO₂ NPs-induced alterations in the mRNA expression of marker genes associated with apoptosis in A549 cells exposed to various concentrations of TiO₂ NPs for 12 h. Semi-quantitative PCR (RT-PCR) was performed in triplicate. GAPDH was used as internal control to normalize the data and TiO₂ NPs-induced alterations in mRNA expression are expressed in fold change. Lane 1: Control; lane 2: 1.0 μg/ml; lane 3: 10.0 μg/ml; lane 4: 50.0 μg/ml, except GAPDH. (b) Relative quantification of alterations in the expression of genes involved in the induction of apoptosis in A549 cells exposed to various concentrations of TiO₂ NPs for 12 h. GAPDH was used as internal control to normalize the data. Quantification was done in Gel Documentation System. *p < 0.01 considered as significant. GAPDH: glyceraldehyde 3-phosphate dehydrogenase; mRNA: messenger RNA; RT-PCR: real-time polymerase chain reaction; TiO₂ NPs: titanium dioxide nanoparticles.

Translational changes

Results of the Western blot analysis showed significant up-regulation of gene protein of P⁵³ (4.91 ± 0.51, 6.29 ± 0.32 and 4.86 ± 0.68), P²¹ (3.76 ± 0.20, 4.21 ± 0.41 and 3.13 ± 0.25) and cleaved caspase-3 (1.39 ± 0.62, 4.86 ± 0.37 and 3.08 ± 0.5) and down-regulation of BCL₂ (0.973 ± 0.09, 0.369 ± 0.25 and 0.215 ± 0.14) following TiO₂ NPs exposure at concentrations 1, 10 and 50 μg/ml for 24 h, respectively (Figure 8(a) and (b)). The trend of cellular response at protein level was similar to transcriptional changes. No specific correlation could be established between the exposure and effect following the exposure of 6 and 12 h (data not shown).

Figure 8.

(a) Alterations in the expression of proteins involved in the induction of apoptosis were studied in A549 cells exposed to various concentrations of TiO₂ NPs for 24 h. GAPDH was used as internal control to normalize the data. Lane (1) untreated control, lane (2) 1.0 μg/ml, lane (3) 10.0 μg/ml and lane (4) 50.0 μg/ml of TiO₂ NPs. (b) Relative quantification of alterations in the expression of proteins involved in the induction of apoptosis in A549 cells exposed to various concentrations of TiO₂ NPs for 24 h. GAPDH was used as internal control to normalize the data. Quantification was done in Gel Documentation System (Alpha Innotech, USA) with the help of AlphaEase^TM FC StandAlone V.4.0 software. *p < 0.01 considered as significant. GAPDH: glyceraldehyde 3-phosphate dehydrogenase; TiO₂ NPs: titanium dioxide nanoparticles.

Discussion

Internalization and toxic responses of NPs are suggested to be associated with their size, composition, surface charge, agglomeration roughness and surface coatings.^24,25 In the present study, the hydrodynamic size and zeta potential of TiO₂ NPs particles were measured at 434.1 nm and −7.83 mV, respectively. Size was many fold higher than that claimed (25 nm) by the manufacturer (Sigma Aldrich, Missouri, USA). This difference was not unusual, since such NPs are known to have the tendency of agglomerate in aqueous suspensions and provided clustered size rather than individual particles.²⁶ Our results show a negative zeta potential for the TiO₂ NP (−7.83 mV), which is an indirect parameter for surface charge. In cell culture medium, the surface charge on the NPs measured by zeta potential was less negative due to the presence of inorganic cations in the physiological media.²⁷ These cations are attracted to the negative charged surface of NPs. A less negative zeta potential favors the particle agglomeration more rapidly and that is why we observed enhanced agglomeration. Negative surface charge causes more cytotoxicity and can cross the blood–brain barrier.²⁸ Zeta potential gives the degree of repulsion force between particles, which can be used to predict the long-term stability of the particle in a particular medium. However, it does not give any information about the element present on the surface of the particles.²⁹ We and others have confirmed such solvent-induced agglomeration by DLS/TEM studies using other commercially available NPs.³⁰

Our cytotoxicity data reveal that the retention period and concentration of TiO₂ NPs are directly proportional to induce the detrimental responses in A549 cells. Cytotoxic effects were detectable only at higher concentrations of TiO₂ NPs. This might be due to the agglomeration of the particles in cytoplasm, which leads the physiological unrest by vacuole formation hypertrophy of other cell organelles. Since, the tendency of agglomeration of NPs was also seen in neural stem cells under similar experimental setup.³¹ The similar trend for cytotoxic responses was also observed with other cells such as human lymphoblastoid³² and mouse epidermal (JB6) cell line³³ exposed to TiO₂ NPs. Recent study conducted by Lankoff et al. examined how the agglomeration/aggregation state of silver (20 nm and 200 nm) and TiO₂ (21 nm) NPs may affect the kinetics of cellular binding/uptake and ability to induce cytotoxic responses in THP1, HepG2 and A549 cells and observed that A549 cells were most sensitive to these particles in relation to cellular binding/uptake.¹¹ NPs can hinder with various colored dyes and their intermediates used in colorimetric and fluorometric-based assays. This may result to false impression of data.³⁴ In the present study, this was obviated by a parallel set of experiments conducted to check the interaction of NPs with MTT and DCF-DA dyes. Our data reveal that TiO₂ NPs do not interrelate with these dyes used for cytotoxicity assessment as well as for ROS measurement.

ROS-mediated oxidative stress is a generalized phenomenon in most of the NP-induced damages.³⁵ Thus, the release of oxy radicals and LPO is essential to assess the role of oxidative injury. Our data demonstrate significant oxidative stress in the cells as the levels of ROS and LPO were found increased and levels of GSH and CAT were reduced. Decreased GSH and CAT activity only at higher concentration at all the exposure time might be either due to the inability of A549 cells to eliminate overwhelming quantity of H₂O₂ and its metabolites, produced by the exposure to TiO₂ NPs, or due to the enzyme inactivation caused by excess ROS production.³⁶ Increased H₂O₂ levels in the cells are known to form hydroxyl radicals, a potent activator of LPO³⁷ and we have also reported the elevated levels of LPO in A549 cells exposed to multiwalled carbon nanotubes.³⁰ The lower intracellular concentrations of H₂O₂ result in subtle signaling events by enzymatic removal of H₂O₂, while high concentrations are known to affect cellular homeostasis, which ultimately lead to apoptotic/ necrotic cell death.³⁸ H₂O₂ also mediate the apoptosis by the activation of transcription factor NF-κB-dependent signaling, which is linked to cellular glutathiolation levels.^39,40 Therefore, such oxidative stress–mediated metabolic instability in the cells possibly triggers the apoptosis cascade, as we observed the positive correlation between oxidative stress and expression change in genes involved in apoptosis. Besides, the above mechanism described, there is a direct relationship between the surface area and the ROS-generating capability in the lungs.⁴¹ TiO₂ NPs used in the present investigations were in anatase phase and were having comparatively larger surface area of about 200–220 m²/g, which might be the partial reason of higher ROS production. The data of cytotoxicity clearly indicate that the higher concentration of TiO₂ NPs (50 μg/ml) induces adverse effects in the cells when exposed for 24 h. Therefore, this might be one among the reasons being decreased response of ROS generation at this concentration at 24 h when compared with 12 h exposure. Significant MN induction in the present investigation suggests the genotoxic potential of TiO₂ NPs in A549 cells. It could be well correlated with the increased expression of P⁵³, biomarker of genetic toxicity.⁹ Our findings are parallel to the studies conducted in Syrian hamster embryo fibroblast cells using ultrafine particles of TiO₂.²² Association of MN induction and p⁵³ expression is well established with the ROS generation.⁴² Thus, the genotoxic responses of TiO₂ NPs in A549 cells may be suggested as a part of whole cascade, finally leading to the apoptosis.

We observed more number of apoptotic bodies in extended exposure (48 h) of TiO₂ NPs than shorter exposure (24 h) and no significant numbers in early hours of exposure (6 and 12 h). Such delayed phenotypic responses in the cells could be correlated with the expressional changes at both mRNA and protein level for the studied apoptosis markers. The transcriptional changes were most significant at 12 h, while this time point shifted to 24 h for translational changes. Therefore, the alterations at phenotypic expression and physiological functionality were quite anticipated few hours later, which was required to complete the post-translational processing and subsequent downstream realization.³⁰ These time- and concentration-based sequence of events also advocates the possible relationship between ROS generation and LPO-mediated apoptosis in A549 cells. Similar kind of observations has also been reported with other NPs in primary cultures.⁴³ On the event of oxidative stress and DNA damage, p⁵³ is one among the important triggering molecules responsible to regulate the intracellular redox status and induction of apoptosis by activation of downstream targets such as p²¹, Bax and caspase-3⁴⁴ and subsequent blocking of cell cycle.⁴⁵ We observed the up-regulation of P⁵³ and its downstream effectors, that is, P²¹ and caspase-3 in A549 cells. Such kind of modulation in apoptosis implicating protein P⁵³ and its downstream effectors has also been reported in human peripheral blood lymphocyte.⁴⁶ The cell cycle arrest is an important signal for apoptosis process and the probable function of p²¹ is the inhibition of cycline-dependent protein kinase and causes growth arrest in G1/S subphase of cell cycle.⁴⁵ Decrease in the expression of anti-apoptotic protein Bcl₂ might be suggested as an important finding to facilitate the formation of permeability transition pore in mitochondria and the release of cytochrome c.⁴⁴ Cytochrome c leads to the activation of the caspase cascade such as caspase-3. Caspase-3 plays an important role in the catabolic phase of cell death by cleaving the various intracellular substrates and also found to activate and amplify caspase cascade.⁴⁴ ROS-induced activation of caspase-3 in cell system is well documented.^18,44 This also supports our observation of up-regulation of caspase-3 (mRNA and protein) expression at time points of TiO2 NPs exposure, where ROS generation was most significant. Furthermore, reports show that caspase-3-mediated chromosomal condensation and DNA fragmentation induced apoptosis of cells,⁴⁷ thus supporting the findings of the present investigation. The possible schematic mechanism of TiO₂ NPs-induced cellular and molecular toxicity in human lung cancer cells (A549) under in vitro condition is illustrated in Figure 9.

Figure 9.

A schematic diagram showing the possible mechanism of nano-titanium dioxide-induced apoptosis pathway (cell death) in human lung cancer cell line (A549).

Conclusion

Experimental exposure of TiO₂ NPs (10 and 50 μg/ml) was found to induce oxidative stress, genotoxicity and apoptosis in human lung cancer cell line, A549. Expressional changes in apoptosis markers were having good correlation with endpoints of oxidative stress and phenotypic presentation of apoptotic/genotoxic events. Data reveal the toxicological consequences of A549 cells against experimental exposure of TiO₂ NPs by inducing oxidative stress, MN induction and apoptosis. The data enhance our understanding on cellular and molecular behavior of human lung cancer cells against TiO₂ NPs exposure and may be helpful to suggest the biosafety/toxicity of other nanomaterials. These results highlight the need for caution during manufacturing/use of nanomaterials as well as further investigation on signaling cascade of TiO₂ NPs-induced toxicity mechanism.

Footnotes

Acknowledgement

Authors are grateful to the Director, Indian Institute of Toxicology Research, Lucknow, India, for his keen interest in the study.

Conflict of interest

The authors declared no conflicts of interest.

Funding

Financial support by the Council of Scientific & Industrial Research, New Delhi, India (SIP-08) is acknowledged.

References

Nielsen

Roursgaard

Jensen

Poulsen

Larsen

ST.

In vivo biology and toxicology of fullerenes and their derivatives. Basic Clin Pharmacol Toxicol 2008; 103:197–208.

Scuri

Chen

Castranova

Reynolds

Johnson

Samsell

Effects of titanium dioxide nanoparticle exposure on neuroimmune responses in rat airways. J Toxicol Environ Health A 2010; 73: 1353–1369.

Liu

Zhang

Yin

Zhang

Small-sized titanium dioxide nanoparticles mediate immune toxicity in rat pulmonary alveolar macrophages in vivo. J Nanosci Nanotechnol 2010; 10: 5161–5169.

Simeonova

Erdely

. Engineered nanoparticle respiratory exposure and potential risks for cardiovascular toxicity: predictive tests and biomarkers. Inhal Toxicol 2009; 2: 68–73.

Wang

Fan

Gao

Wang

. TiO₂ nanoparticles translocation and potential toxicological effect in rats after intraarticular injection. Biomaterials 2009; 30: 4590–4600.

Kang

Kim

Lee

Hong

Chung

. Titanium dioxide nanoparticles induce apoptosis through the JNK/p38-caspase-8-Bid pathway in phytohemagglutinin-stimulated human lymphocytes. Biochem Biophys Res Commun 2009; 386: 682–687.

Foldbjerg

Dang

Autrup

. Cytotoxicity and genotoxicity of silver nanoparticles in the human lung cancer cell line A549. Arch Toxicol 2011; 85: 743–750.

De Berardis

Civitelli

Condello

Lista

Pozzi

Arancia

Exposure to ZnO nanoparticles induces oxidative stress and cytotoxicity in human colon carcinoma cells. Toxicol Appl Pharmacol. Epub ahead of print 29 April 2010. DOI:10.1016/j.taap.2010.04.012.

Ahamed

Siddiqui

Akhtar

Ahmad

Pant

Alhadlaq

. Genotoxic potential of copper oxide nanoparticles in human lung epithelial cells. Biochem Biophys Res Commun 2010; 396: 578–583.

10.

Srivastava

Rahman

Kashyap

Lohani

Pant

. Ameliorative effects of dimetylthiourea and N-acetylcysteine on nanoparticles induced cyto-genotoxicity in human lung cancer cells-A549. PLoS One 2011b; 6(9): e25767.

11.

Lankoff

Sandberg

Wegierek-Ciuk

Lisowska

Refsnes

Sartowska

The effect of agglomeration state of silver and titanium dioxide nanoparticles on cellular response of HepG2, A549 and THP-1 cells. Toxicol Lett 2012; 208: 197–213.

12.

Donaldson

Stone

Gilmour

Brown

Macnee

. Ultrafine particles: mechanisms of lung injury. Philos Trans R Soc A: Math Phys Eng Sci 2000; 358: 2741–2749.

13.

Sayes

Warheit

. Characterization of nanomaterials for toxicity assessment. Wiley Interdiscip Rev Nanomed Nanobiotechnol 2009; 1: 660–760.

14.

Hsiao

Huang

. Effects of various physicochemical characteristics on the toxicities of ZnO and TiO(2) nanoparticles toward human lung epithelial cells. Sci Total Environ 2011; 409: 1219–1228.

15.

Anselmann

. Nanoparticles and nanolayers in commercial applications. J Nanoparticle Res 2001; 3: 329–336.

16.

Chen

Dong

Zhao

Tang

In vivo acute toxicity of titanium dioxide nanoparticles to mice after intraperitioneal injection. J Appl Toxicol 2009; 29: 330–337.

17.

Duan

Hong

Zheng

Fei

Zhao

Spleen injury and apoptotic pathway in mice caused by titanium dioxide nanoparticles. Toxicol Lett 2010; 195: 161–168.

18.

Park

Chung

Ryu

Choi

Park

. Oxidative stress and apoptosis induced by titanium dioxide nanoparticles in cultured BEAS-2B cells. Toxicol Lett 2008; 180: 222–229.

19.

Liu

Zhang

Ren

Yang

. Oxidative stress and apoptosis induced by nanosized titanium dioxide in PC12 cells. Toxicology 2010; 267: 172–177.

20.

Hussain

Thomassen

Ferecatu

Borot

Andreau

Martens

Carbon black and titanium dioxide nanoparticles elicit distinct apoptotic pathways in bronchial epithelial cells. Part Fibre Toxicol 2010; 7: 10.

21.

Rahman

Lohani

Dopp

Pemsel

Jonas

Weiss

Evidence that ultrafine titanium dioxide induces micronuclei and apoptosis in Syrian hamster embryo fibroblasts. Environ Health Perspect 2002; 110: 797–800.

22.

Srivastava

Lohani

Pant

Rahman

. Cyto-genotoxicity of amphibole asbestos fibers in cultured human lung epithelial cell line: role of surface iron. Toxicol Ind Health 2010; 26: 575–582.

23.

Dini

Coppola

Ruzittu

Ghibelli

. Multiple pathways for apoptotic nuclear fragmentation. Exp Cell Res 1996; 223: 340–347.

24.

Verma

Stellacci

. Effect of surface properties on nanoparticle-cell interactions. Small 2010; 6: 12–21.

25.

Asati

Santra

Kaittanis

Perez

. Surface-charge-dependent cell localization and cytotoxicity of cerium oxide nanoparticles. ACS Nano 2010; 4: 5321–5331.

26.

Murdock

Braydich-Stolle

Schrand

Schlager

Hussain

. Characterization of nanomaterial dispersion in solution prior to in vitro exposure using dynamic light scattering technique. Toxicol Sci 2008; 101: 239–253.

27.

Thomassen

Aerts

Rabolli

Lison

Gonzalez

Kirsch-Volders

Synthesis and characterization of stable monodisperse silica nanoparticle sols for in vitro cytotoxicity testing. Langmuir 2010; 26: 328–335.

28.

Trickler

Lantz

Murdock

Schrand

Robinson

Newport

Silver nanoparticle induced blood-brain barrier inflammation and increased permeability in primary rat brain microvessel endothelial cells. Toxicol Sci 2010; 118: 160–170.

29.

Borm

PJA

Robbins

Haubold

Kuhlbusch

Fissan

Donaldson

The potential risks of nanomaterials: a review carried out for ECETOC. Part Fiber Toxicol 2006; 3: 11.

30.

Srivastava

Pant

Kashyap

Kumar

Lohani

Jonas

Multi-walled carbon nanotubes induce oxidative stress and apoptosis in human lung cancer cell line-A549. Nanotoxicology 2011a; 5: 195–207.

31.

Deng

Luan

Chen

Wang

Zhang

Nanosized zinc oxide particles induce neural stem cell apoptosis. Nanotechnology 2009; 20: 115101.

32.

Wang

Sanderson

Wang

. Cyto- and genotoxicity of ultrafine TiO₂ particles in cultured human lymphoblastoid cells. Mutat Res 2007; 628: 99–106.

33.

Zhao

Bowman

Zhang

Vallyathan

Young

Castranova

Titanium dioxide (TiO₂) nanoparticles induce JB6 cell apoptosis through activation of the caspase-8/Bid and mitochondrial pathways. J Toxicol Environ Health A 2009; 72: 1141–1149.

34.

Monteiro-Riviere

Inman

Zhang

. Limitations and relative utility of screening assays to assess engineered nanoparticle toxicity in a human cell line. Toxicol Appl Pharmacol 2009; 234: 222–235.

35.

Kim

Baek

Choi

. Comparative cytotoxicity of Al2O3, CeO2, TiO2 and ZnO nanoparticles to human lung cells. J Nanosci Nanotechnol 2010; 10: 3453–3458.

36.

Pigeolet

Corbisier

Houbion

Lambert

Michiels

Raes

Glutathione peroxidase, superoxide dismutase and catalase inactivation by peroxides and oxygen derived free radicals. Mech Ageing Dev 1990; 51: 283–290.

37.

Moller

Jacobsen

Folkmann

Danielsen

Mikkelsen

Hemmingsen

Role of oxidative damage in toxicity of particulates. Free Radic Res 2010; 44: 1–46.

38.

Winyard

Moody

Jacob

. Oxidative activation of antioxidant defence. Trends Biochem Sci 2005; 30: 453–461.

39.

Dumont

Hehner

Hofmann

Ueffing

Dröge

Schmitz

. Hydrogen peroxide-induced apoptosis is CD95-independent, requires the release of mitochondria derived reactive oxygen species and the activation of NF-kappaB. Oncogene 1999; 18: 747–757.

40.

Kim

. Oxidative stress attenuates Fas-mediated apoptosis in Jurkat T cell line through Bfl-1 induction. Oncogene 2005; 24: 1252–1261.

41.

Donaldson

Stone

Tran

Kreyling

Borm

. Nanotoxicology. Occup Environ Med 2004; 61: 727–728.

42.

Karlsson

Cronholm

Gustafsson

Möller

. Copper oxide nanoparticles are highly toxic: a comparison between metal oxide nanoparticles and carbon nanotubes. Chem Res Toxicol 2008; 21: 1726–1732.

43.

Johnston

Semmler-Behnke

Brown

Kreyling

Tran

Stone

. Evaluating the uptake and intracellular fate of polystyrene nanoparticles by primary and hepatocyte cell lines in vitro. Toxicol Appl Pharmacol 2010; 242: 66–78.

44.

Galluzzi

Blomgren

Kroemer

. Mitochondrial membrane permeabilization in neuronal injury. Nat Rev Neurosci 2009; 10: 481–494.

45.

Satyanarayana

Hilton

Kaldis

p21 Inhibits Cdk1 in the absence of Cdk2 to maintain the G1/S phase DNA damage checkpoint. Mol Biol Cell 2008; 19: 65–77.

46.

Kang

Kim

Lee

Chung

. Titanium dioxide nanoparticles trigger p53-mediated damage response in peripheral blood lymphocytes. Environ Mol Mutagen 2008; 49: 399–405.

47.

Kivinen

Kallajoki

Taimen

. Caspase-3 is required in the apoptotic disintegration of the nuclear matrix. Exp Cell Res 2005; 311: 62–73.