TY - JOUR
AU - Kumar,, Ashutosh
AB - Abstract Zinc oxide nanoparticles (ZnO NPs) with their wide range of consumer applications in day-to-day life received great attention to evaluate their effects in humans. This study has been attempted to elucidate the DNA damage response mechanism in a dermal model exposed to ZnO NPs through Ataxia Telangiectasia Mutated (ATM)-mediated ChK1-dependent G2/M arrest. Further, viability parameters and mechanism involved in the cell death with special reference to the consequences arising due to DNA damage were explored. Our study showed that ZnO NPs at concentrations 5 and 10 µg/ml induced significant cytotoxic effect in skin cell line. Moreover, the results confirmed generation of reactive oxygen species (ROS) induces the cell death by genotoxic insult, leading to mitochondrial membrane depolarisation and cell cycle arrest. Subsequently, ZnO NPs treatment created DNA damage as confirmed via Comet assay (increase in olive tail moment), micronucleus assay (increase in micronucleus formation), double-strand breaks (increase in ATM and Ataxia Telangiectasia and Rad3 related (ATR) expression), DNA fragmentation and cell cycle (G2/M arrest) studies. Finally, marker proteins analysis concluded the mechanistic approach by demonstrating the key marker expressions HMOX1 and HSP60 (for oxidative stress), cytochrome c, APAF1, BAX, Caspase 9, Caspase 3 and decrease in BCL2 (for activating apoptotic pathway), pATM, ATR and γH2AX (for double-strand breaks), DNA-PK (involved in DNA repair) and decrease in cell cycle regulators. In together, our data revealed the mechanism of ROS generation that triggers apoptosis and DNA damage in HaCaT cell lines exposed to ZnO NPs. Introduction Zinc oxide nanoparticles (ZnO NPs) due to their versatile and distinct properties are being widely used in cosmetics, food additive, drug delivery, biosensors, bio-imaging, disease therapy and as antimicrobial agents (1–5). ZnO NPs are present in various sunscreens and facial creams as they protect the cells against the ultraviolet (UV)-induced skin damage and do not undergo any chemical decomposition when exposed to UV radiation (6). At present, many products for dermal applications use ZnO as they are more aesthetically pleasing to consumers at nanoscale. This is largely due to their physicochemical properties and functionality of ZnO NPs attributed to the unique size. Hence, a holistic understanding about the molecular effects of ZnO NPs in the living system is important, for both the design of safe functionalised particles and novel nanomaterials-based formulations for consumer products. The skin is often considered less permeable compared to other organ systems and the risk of exposure to different chemical entities by this route is generally less compared to inhalation route of exposure (7,8). Previous studies have assessed the ability of NPs to penetrate skin (9–11) and have suggested it as a major route of entry. Also, the properties of NPs and the nature of the vehicle used to suspend the NPs effects the penetration ability of different NPs (7,10). For example, TiO2 NPs have deeper penetration into human skin in an oily suspension compared to that of an aqueous dispersion (12). However, encapsulation of TiO2 NPs into liposomes leads to deep internal penetration. In addition, penetration is greater through hairy skin, suggesting surface penetration through pores or follicles of hair. It has also been reported that some NPs cross the stratum corneum and move into the dermis to reach distal organ sites (7,10). ZnO NPs have been demonstrated to cause various toxic effects such as membrane injury, inflammatory response, DNA damage and apoptosis in different models or cell lines (13–17). The particulate nature of ZnO NPs induces reactive oxygen species (ROS) generation (17–20), which further causes DNA damage ultimately culminating to apoptosis (16,21). However, there is considerable inconsistency in the reported data, which needs further investigation with special reference to DNA damage and the mechanism through which the particle alters the normal cell cycle process. ZnO NPs are known to activate the mitogen-activated protein kinase (MAPK) pathway, which is a crucial mediator of signal transduction and plays an important role in regulating different cellular processes (22,23). However, the molecular mechanisms induced by ZnO NPs in eliciting toxicity remained unclear. Therefore, this study has been attempted to elucidate the DNA damage response (DDR) mechanism for double-strand breaks in a dermal model exposed to ZnO NPs. DDR is a complex mechanism, exists in response to DNA damage and involves in both damage detection and DNA repair (24,25). Further, the study focused on determining viability parameters and mechanism involved in the cell death with special reference to the consequences aroused due to DNA damage. Materials and Methods Materials ZnO nanopowder (purity 99%; CAS No. 1314-13-2), lactate dehydrogenase (LDH) assay kit (cat no. TOX7), neutral red dye, acridine orange, 2,7-dichlorofluorescein diacetate, N-acetyl cysteine (NAC), cell lysis buffer and protease inhibitor were purchased from Sigma Chemical Co. Ltd (St Louis, MO, USA). Normal-melting agarose and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) dye were purchased from HiMedia Laboratories Pvt. Ltd (Mumbai, India). Dulbecco’s modified Eagle’s medium (DMEM), trypsin-EDTA, FBS (foetal bovine serum) and 5,5′,6,6′ tetra ethylbenzimidazolocarbocyanine iodide (JC-1) dye were purchased from Life Technologies (India) Pvt Ltd, (New Delhi, India). Annexin V-FITC Apoptosis Detection Kit was obtained from BD Pharmingen (San Jose, CA, USA). FlowCellect™ Multi-Color DNA Damage Response Kit (FCCH025104) was purchased from Merck Millipore. All other chemicals were procured locally and were of analytical reagent grade. Methods Suspension of ZnO NPs in culture media and exposure to HaCaT cells ZnO NPs (100 µg/ml) were suspended in DMEM (without FBS) and probe sonicated (Vibra Cell; Sonics & Materials Inc, New Town, CT, USA) at 30 W for 10 min (1.5 min pulse on and 1 min pulse off for four times). After sonication, the suspension was diluted in CDMEM (complete DMEM; supplemented with 10% FBS) for cell exposure. The human keratinocyte cell line (HaCaT) was obtained from National Centre for Cell Sciences, Pune, India and cultured in DMEM supplemented with 10% FBS, 2 mM l-glutamine, 0.2% sodium bicarbonate and 10 ml/L antibiotic and antimycotic solution under a humidified atmosphere at 37°C with 5% CO2. Stock suspension of ZnO NPs (100 µg/ml) in DMEM was diluted to concentrations ranging from 0.5, 1, 2.5, 5 and 10 µg/ml in CDMEM and was treated to the cell line respectively. Also, to investigate the role of ROS, the cells were treated with 1 mM NAC for 2 h before ZnO NPs treatment. Characterisation of ZnO NPs ZnO NPs were characterised in complete DMEM using dynamic light scattering (DLS) instrument (Zetasizer Nano-ZS, Model ZEN3600; Malvern Instruments Ltd, Malvern, UK). Further, size of ZnO NPs suspended in Milli-Q (50 µg/ml) was determined by transmission electron microscope (TEM; FEI Tecnai G2 Spirit TWIN, Eindhoven, the Netherlands). Cellular uptake of ZnO NPs The cellular uptake of NPs was carried out according to the method of Suzuki et al. (26) using light scattering principles by flow cytometer (BD Biosciences, San Jose, CA, USA). Moreover, ultrathin sections of HaCaT cells were analyzed to reveal the subcellular localisation of ZnO NPs in HaCaT cells by using TEM (FEI Tecnai G2 Spirit TWIN). The samples were prepared as per the protocol described by Shukla et al. (27). Cytotoxicity assays Cytotoxicity assessment of ZnO NPs was determined by MTT, neutral red uptake (NRU) and LDH assays. MTT and NRU assays were carried out according to the method of Mosmann (28), Borenfreund and Puerner (29), respectively, whereas LDH assay was conducted by kit (Sigma–Aldrich Inc, St. Louis, MO, USA) using manufacturer’s protocol. Briefly, 1 × 104 cells/well were seeded in 96-well plates and kept for 24 h. Cells were then treated to different concentrations of ZnO NPs (0.5, 1, 2.5, 5 and 10 µg/ml) for varying time intervals (3, 6 and 24 h). Nanoparticle interference with reagents was assessed using a cell-free system and absorbance was recorded at their respective wavelengths using multi-well plate reader (SYNERGY-HT, Bio-Tek (USA) using KC4 software). Reactive oxygen species Generation of ROS in HaCaT cells was quantified using 2′,7′-Dichlorofluorescin diacetate (DCFDA) dye by flow cytometer. After 3 and 6 h of exposure, the treatment of ZnO NPs (0.5, 1, 2.5, 5 and 10 µg/ml) was discarded and cells were harvested using trypsin EDTA. After washing the cells with PBS, cells were re-suspended in PBS containing 20 µM DCFDA and incubated for 30 min in dark. PBS of 300 µl was further added to the sample and fluorescent intensity was recorded using BD FACSCanto II flow cytometer (BD FACS Diva 6.2.1 software; BD Biosciences). Qualitative analysis of ROS generation was performed using fluorescence microscope (DMLB, Leica, Germany). Apoptosis markers Mitochondrial membrane potential. The effect of ZnO NPs on the mitochondrial membrane potential (MMP; Δψ) of HaCaT cells was measured using lipophilic cationic dye JC-1. JC-1 selectively enters into the mitochondria and changes its colour from red to green, in case of decreased membrane potential. Cells were treated with ZnO NPs (0.5, 1, 2.5, 5 and 10 µg/ml) for 3 and 6 h. Non-treated cells served as a control and camptothecin (1 mM) were used as a positive control. After the removal of treatment, cells were washed and incubated with 10 mM of JC-1 dye for 15 min at 37°C. Cells were acquired using BD FACSCanto II (BD FACS Diva 6.2.1 software; BD Biosciences). Further, cells were imaged for green and red fluorescence using a fluorescence microscope (DMLB). Annexin V binding assay. Apoptosis/necrotic cell population were identified by staining them with fluorescein isothiocyanate-conjugated (FITC)—annexin V and Propidium Iodide (PI) dye as per the manufacturer’s protocol (BD Biosciences). Cells were treated with different concentrations of ZnO NPs (0.5, 1, 2.5, 5 and 10 µg/ml) for 6 and 24 h. Camptothecin (1 mM) was used as a positive control. After treatment, cells were harvested and washed twice with PBS, re-suspended in 0.1-ml binding buffer containing 5 µl of FITC-annexin V, PI and incubated at room temperature in dark. After 10 min of incubation, 0.4 ml of binding buffer was further added to each sample and analysed using BD FACSCanto II (BD FACS Diva 6.2.1 software; BD Biosciences). Genotoxicity assessment The genotoxic potential of ZnO NPs was assessed by Comet assay, micronucleus (MN) assay and FlowCellect™ Multi-Color DNA Damage Response Kit. The cells were exposed to ZnO NPs (0.5, 1, 2.5, 5 and 10 µg/ml) for 3 and 6 h. Comet assay. Cells were harvested and slides were prepared by the method described by Singh et al. and modified by Bajpayee et al. (30,31). The scoring of the cells was done at a final magnification of ×400 using Komet 5.0 software provided with the image analysis system (Andor Technology, Belfast, UK) attached with fluorescent microscope (DMLB) equipped with CCD camera and appropriate filter. Olive tail moment (OTM) parameter was used to measure the DNA damage. Analysis of 50 Comets (25 from replicate slide) was carried out for all concentrations based on the protocol described by Tice et al. (32). MN assay. The frequency of MN formation in ZnO exposed cells was determined by flow cytometer. After 3 and 6 h of exposure, ZnO NPs suspension was aspirated; cells were washed with PBS and grown further for 21 and 18 h in complete medium respectively. After incubation, cells were harvested and centrifuged at 1200 rpm for 10 min. Cells were washed in 1× PBS and transferred to 1 ml of solution I containing sodium chloride (584 mg/L), sodium citrate (1000 mg/L), RNase (10 mg/L), ethidium bromide (25 mg/L), Igepal (0.3 ml/L) and incubated for 1 h at room temperature. Finally, 1 ml of solution II containing citric acid (15 mg/L), sucrose (8.55 gm/L), ethidium bromide (40 mg/L) was added in to the same tube and the samples were analysed for the presence of micronuclei in the cells using BD FACSCanto II (BD FACS Diva 6.2.1 software; BD Biosciences). DNA double-strand break detection. Millipore’s FlowCellect™ Multi-Color DNA Damage Response Kit was used to detect the DNA double-strand breaks. Cells were treated with different concentrations of ZnO NPs for 3 and 6 h. After treatment, cells were fixed and incubated with assay and permeabilisation buffer along with 10 µl of ×20 antibody solution and incubated in dark for 60 min. Cells were washed to remove the unbound antibodies and analysed using BD FACSCanto II (BD FACS Diva 6.2.1 software; BD Biosciences). Cell cycle analysis HaCaT cells (1 × 105 cells/ml) were seeded on 12-well plate and treated with different ZnO NPs concentrations (0.5–10 µg/ml) for 3, 6 and 24 h. After the treatment, medium was aspirated into individual tubes and cells were collected and fixed in 70% ethanol (at –20°C for 30 min). After fixation, cells were permeated in 1-ml PBS containing 0.2% Triton X 100 at 4°C for 30 min. Cells were pelleted down by centrifugation and re-suspended in 500 µl of PBS containing 20 µl RNase (10 mg/ml) for 30 min at 37°C. Finally, cells were stained with 10 µl propidium iodide (1 mg/ml) in 500 µl PBS for 10–15 min at 4°C and analyzed on BD FACSCanto II (BD FACS Diva 6.2.1 software; BD Biosciences). Histogram of count vs FL2 intensity was used to calculate the percentage of cells under Go/G1 (2n), S (2n+) and G2/M phase (4n). Western blot analysis HaCaT cells were treated with ZnO NPs at concentrations 2.5, 5 and 10 µg/ml for 24 h. After treatment removal, cells were harvested and lysed in lysis buffer (150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% Sodium Dodecyl Sulfate (SDS), 50 mM Tris–HCl, pH 7.5, 2 mM EDTA) containing protease inhibitor cocktail (Sigma, USA). Protein concentration was estimated using Bradford’s method (33). Protein (50 µg) from control and treated groups was separated on tricine–SDS–polyacrylamide gel (12%) and transferred to a polyvinylidene difluoride membrane by electroblotting. The membrane was incubated with primary antibodies specific for HSP60 (Anti-Heat Shock Protein 60 antibody, MAB3514), HMOX1 (Anti-Heme Oxygenase-1 antibody, 374087), p-p53 [Anti-phospho-p53 (Ser6) antibody, 04-540], BAX (Anti-BAX Antibody, MAB4601), BCL2 (Anti-BCL2 antibody, MABC573), Caspase-3 [Anti-Caspase 3 antibody, active (cleaved) form, AB3623], Caspase-9 [Anti-Caspase 9 antibody, active (cleaved) form, AB3629], APAF1 (Anti-Apaf-1 antibody, MABC1187), cytochrome c (Anti-Cytochrome c antibody, MAB1800), p-Ataxia Telangiectasia Mutated (ATM) (Anti-ATM phosphoSer1981 antibody, MAB3806), p-Ataxia Telangiectasia and Rad3 related (ATR) [Anti-phospho-ATR (Ser428) antibody, ABE389], γH2AX (Anti-Histone H2A.X antibody, ABE1960), CDC25A (Anti-Cdc25A antibody, 05-743), Cyclin D (Anti-Cyclin D antibody, 06-137), Cyclin B1 (Anti-Cyclin B1 antibody, 05-373), DNA-PK (Anti-DNA-PKcs antibody, MABC1236), CDC2 (Anti-Cdc2 antibody, AB3241-25UL), β-actin (Anti-beta-Actin antibody, MABT523; purchased from Millipore, India) and CHK1 [Anti-Chk1 phospho S345 antibody (ab58567) was purchased from Abcam, UK]. Secondary antibody incubation was done and protein bands were detected using chemiluminescence. The densitometry analysis of the bands was carried out using Quantity One Quantitation Software, version 4.3.1 (Bio-Rad, USA). Statistical analysis Mean ± SEM from three individual experiments was calculated and analysed using one-way analysis of variance for mean variation between groups. Later Dunnett’s test was used to identify the significant difference (P < 0.05) from control. Results Particle characterisation The mean hydrodynamic diameter of ZnO in PBS and DMEM media as measured by DLS was 185 and 144 nm, zeta potential values were –26 and –17.6 mV respectively. DLS measurement reflects the actual size of NPs in media, which is considered as the real size exposed to cells. However, the average size of ZnO as observed by transmission electron microscope (TEM) was 20–50 nm (Figure 1A). Figure 1. Open in new tabDownload slide Characterisation of nanoparticles and its internalization (A) photomicrograph of ZnO NPs captured by TEM. (B–F) Internalisation of ZnO NPs in HaCaT cells using (B) flow cytometer. (C–F) Transmission electron microscope. Figure 1. Open in new tabDownload slide Characterisation of nanoparticles and its internalization (A) photomicrograph of ZnO NPs captured by TEM. (B–F) Internalisation of ZnO NPs in HaCaT cells using (B) flow cytometer. (C–F) Transmission electron microscope. ZnO NPs are taken up by HaCaT cells Cellular internalisation of ZnO NPs after 6-h treatment in HaCaT was revealed by flow cytometric analysis. A significant (P < 0.05) concentration-dependent increase in NPs internalisation was observed compared to control (Figure 1B; Supplementary Figure 1, available at Mutagenesis Online), which was evident by increase in granularity (side scatter intensity). Further, subcellular localisation of ZnO NPs was confirmed using TEM. Results showed that the ZnO NPs localised both in the cytoplasm and nucleus (Figure 1E–F) compared with control (Figure 1C–D). ZnO NPs are cytotoxic to HaCaT cells To determine the cytotoxicity potential of the ZnO NPs in HaCaT cells, the MTT, LDH and NRU assays were performed. MTT results showed both concentration and time-dependent cytotoxicity after treatment to ZnO NPs. The percent cell viability (relative to control) decreased in a graded manner when HaCaT cells were exposed to the ZnO NPs at concentrations of 5 and 10 µg/ml for 3, 6 and 24 h respectively (Figure 2A). Figure 2. Open in new tabDownload slide Cytotoxic effect of ZnO NPs in HaCaT cells as assessed by (A) MTT assay (B) lactate dehydrogenase release assay (C) neutral red uptake (NRU). The viability of control cells was considered as 100%. The data are expressed as mean ± SEM from three independent experiments. (*P < 0.05 and ** P < 0.01 compared to control). Figure 2. Open in new tabDownload slide Cytotoxic effect of ZnO NPs in HaCaT cells as assessed by (A) MTT assay (B) lactate dehydrogenase release assay (C) neutral red uptake (NRU). The viability of control cells was considered as 100%. The data are expressed as mean ± SEM from three independent experiments. (*P < 0.05 and ** P < 0.01 compared to control). The cell viability percentage after treatment to 5 and 10 µg/ml of ZnO NPs compared to control was 88.6 and 67.4% at 3 h, 62.1 and 38.5% at 6 h and 50.1 and 9.2% at 24 h respectively. The IC50 value of ZnO NPs against the HaCaT cells at 6 h was 8.4 µg/ml and further decreased to 6.1 µg/ml for 24 h as calculated from the MTT assay. We also observed a significant LDH leakage from the ZnO NPs exposed cells (126–175%) after 24-h treatment compared with control at all concentrations (0.5–10 µg/ml) respectively (Figure 2B), which further corroborated the results of the MTT assay. The results of the NRU assay showed a similar concentration and time-dependent response with a loss in cell viability (5–10 µg/ml) after 6 and 24 h of treatment (Figure 2C). Treatment with ZnO NPs causes generation of ROS inside HaCaT cells To understand the molecular effects of the ZnO NPs inside HaCaT cells, we analyzed the cellular redox status after treatment to ZnO NPs by investigating the status of cellular ROS. We observed a significant concentration and time-dependent increase in the intracellular ROS levels after ZnO NP treatment. The flow cytometric analysis showed a significant increase in the intensity of FITC (9 and 14.8%) at concentration of 5 µg/ml and (11.2 and 16.7%) at concentration of 10 µg/ml compared with control (1.5 and 1.3%) after 3 and 6 h respectively (Figure 3A–F). Further, the qualitative analysis for ROS using fluorescent microscope indicated a clear increase in DCF fluorescence with increasing concentrations (Figure 3H–K) compared with control cells (Figure 3G). Figure 3. Open in new tabDownload slide ZnO NPs induces significant intracellular ROS in HaCaT cells after 3 and 6 h of treatment. (A–C) Histograms exhibiting increase in ROS after 3 h. (D–F) Histograms exhibiting increase in ROS after 6 h. (G–K) Microphotographs of HaCaT cells exhibiting ROS generation after treatment with ZnO NPs at concentrations 0, 0.5, 2.5, 5 and 10 µg/ml for 3 h. Figure 3. Open in new tabDownload slide ZnO NPs induces significant intracellular ROS in HaCaT cells after 3 and 6 h of treatment. (A–C) Histograms exhibiting increase in ROS after 3 h. (D–F) Histograms exhibiting increase in ROS after 6 h. (G–K) Microphotographs of HaCaT cells exhibiting ROS generation after treatment with ZnO NPs at concentrations 0, 0.5, 2.5, 5 and 10 µg/ml for 3 h. ZnO NPs induced generation of ROS causes mitochondrial membrane depolarisation To understand the physiological consequence of the ROS generated after treatment to ZnO NPs, we investigated the activation of cell death pathways. It is well known that mitochondrial membrane depolarisation marks the initiation of the apoptotic pathway (34,35). Thus, we investigated mitochondrial membrane depolarisation using JC-1 dye. As expected, the shift in the red (j-aggregates) to green fluorescence (JC-1 monomer) was observed in ZnO NPs-treated cells. Our data showed 2.9 and 5.2% depolarised cells at concentration 5 and 10 µg/ml compared with control 2.6% after 3-h treatment. Further, treatment for 6 h increased the depolarised cell population up to 8.1 and 11.4% at same concentrations (Figure 4A–G). In addition, a significant reduction in the mitochondrial membrane depolarisation was observed in the cells treated with NAC and 10 µg/ml of ZnO NPs showed the involvement of ROS in mitochondrial membrane depolarisation. The qualitative analysis of MMP using fluorescent microscope also showed a weak green intensity in control cells, whereas the ZnO NP-treated cells indicated a bright green fluorescence with a clear decrease in red fluorescence (Figure 4H–P). Figure 4. Open in new tabDownload slide Study on mitochondrial membrane potential changes in HaCaT cells after treated with different ZnO NPs concentrations: (A–C) dot plot analysis of JC-1 stained cells after 3-h treatment. (D–F) Dot plot analysis of JC-1 stained cells after 6-h treatment. (G) Bar graph shows the percentage of JC-1 monomer positive cells (% MMP loss). Data of % MMP loss are expressed as mean ± SEM from three independent experiments.*P < 0.05, when compared with control. (H–P) Fluorescent microphotographs of JC-1-stained control and ZnO NPs-treated cells. Figure 4. Open in new tabDownload slide Study on mitochondrial membrane potential changes in HaCaT cells after treated with different ZnO NPs concentrations: (A–C) dot plot analysis of JC-1 stained cells after 3-h treatment. (D–F) Dot plot analysis of JC-1 stained cells after 6-h treatment. (G) Bar graph shows the percentage of JC-1 monomer positive cells (% MMP loss). Data of % MMP loss are expressed as mean ± SEM from three independent experiments.*P < 0.05, when compared with control. (H–P) Fluorescent microphotographs of JC-1-stained control and ZnO NPs-treated cells. ZnO NPs treatment causes externalisation of phosphatidylserine on the surface of HaCaT cells The mitochondrial membrane depolarisation initiates a cascade of events leading to the cellular apoptosis. An important hallmark for apoptosis is externalisation of phosphatidylserine to the outer leaflet of the cell membrane, which can be experimentally quantified by the annexin V binding. Annexin V/PI double positive cells signify the apoptotic cell population. Likewise, our experimental data clearly demonstrated a significant concentration-dependent increase in the apoptotic HaCaT cell population (4.2, 7.2 and 8.9 %) after 6 h of ZnO NPs treatment (2.5, 5 and 10 µg/ml) with most of the cells in the late apoptotic stage (annexin V/PI double positive; Figure 5B–D). However, 24 h of treatment resulted in a significant increase in the necrotic population compared with the control (data not shown). Consequently, nuclear fragmentation (hallmark of apoptosis) was also observed (compared to control) by staining cells with acridine orange after treatment to different concentrations of ZnO NPs (Figure 5E–H). Figure 5. Open in new tabDownload slide Study on apoptosis/necrosis in HaCaT cells after ZnO NPs treatment: (A–D) dot plot analysis of apoptotic cells by the annexin-V/PI staining after 6-h treatment. (E–H) Photomicrographs of HaCaT cells depicting the nuclear condensation after staining with acridine orange on treatment to ZnO NPs (2.5, 5 and 10 µg/ml). Figure 5. Open in new tabDownload slide Study on apoptosis/necrosis in HaCaT cells after ZnO NPs treatment: (A–D) dot plot analysis of apoptotic cells by the annexin-V/PI staining after 6-h treatment. (E–H) Photomicrographs of HaCaT cells depicting the nuclear condensation after staining with acridine orange on treatment to ZnO NPs (2.5, 5 and 10 µg/ml). Holistically, all the aforementioned results converge to the fact that ZnO NPs induce apoptosis in HaCaT cells. ZnO NPs are genotoxic to HaCaT cells We next investigated the genotoxic potential of the ZnO NPs by different experimental approaches. Initially, we performed Comet assay to understand the DNA damage potential of ZnO NPs on HaCaT cells. A significant DNA damage (P < 0.05) was observed in the cells after 3-h treatment of ZnO NPs. The results showed a gradual increase in the OTM (2.06 ± 0.09, 2.18 ± 0.16, 2.55 ± 0.12, 3.26 ± 0.21, 4.89 ± 0.16) at concentrations 0.5, 1, 2.5, 5, 10 µg/ml respectively compared to the control value of 1.8 ± 0.13 (Figure 6A). The fact that 10 µg/ml of ZnO NPs approximately induces 2.7-fold increase in the OTM after 3 h of treatment suggests their potent genotoxic effect in the cells. Figure 6. Open in new tabDownload slide ZnO NPs treatment induces genotoxicity in HaCaT cells: (A) assessment of DNA damage potential of ZnO NPs in HaCaT cells after 3-h treatment using Comet assay. Data of Comet parameters (olive tail moment) are expressed as mean ± SD from three independent experiments.*P < 0.05, when compared with control. (B–E) Dot plot representing the phosphorylation of ATM and SMC1 in HaCaT cells after 3 h of ZnO NPs treatment (performed using FlowCellect Kit). (F–G) Dot plot signifying increase in micronucleus induction after 3-h and 6-h treatment of ZnO NPs compared to control (P2 indicate micronucleus and number above the respective box signify micronucleus percentage). Figure 6. Open in new tabDownload slide ZnO NPs treatment induces genotoxicity in HaCaT cells: (A) assessment of DNA damage potential of ZnO NPs in HaCaT cells after 3-h treatment using Comet assay. Data of Comet parameters (olive tail moment) are expressed as mean ± SD from three independent experiments.*P < 0.05, when compared with control. (B–E) Dot plot representing the phosphorylation of ATM and SMC1 in HaCaT cells after 3 h of ZnO NPs treatment (performed using FlowCellect Kit). (F–G) Dot plot signifying increase in micronucleus induction after 3-h and 6-h treatment of ZnO NPs compared to control (P2 indicate micronucleus and number above the respective box signify micronucleus percentage). To get further mechanistic insights of the DNA damage potential, the occurrence of DNA double-strand breaks in the same experimental setup was investigated using specific antibodies against the established DNA double-strand break markers. We observed a concentration-dependent rise in phosphorylation of ATM and SMC1 (used Anti-pATM tagged with phycoerythrin and Anti-pSMC1 tagged with AlexaFluor-488 were provided with Millipore’s FlowCellect™ Multi-Color DNA Damage Response Kit) protein after 3-h treatment of ZnO NPs in HaCaT cells. There was a significant rise in the intensity of dual positive (pATM and pSMC1) cells (3.6, 9.6 and 32.3% after treatment to 2.5, 5 and 10 µg/ml of ZnO NPs respectively (Figure 6C–E) compared with the control 1.9% (Figure 6B) confirming the occurrence of DNA double-strand breaks. The MN assay is routinely used for toxicological screening of potentially genotoxic compounds. Because the results of Comet assay suggested a severe genotoxic effect of the ZnO NPs, we investigated the effect of ZnO NPs using MN assay. Indeed, we observed a significant increase in the number of micronuclei after ZnO NP treatment. The flow cytometric analysis showed both concentration and time-dependent increase in the micronuclei (5.5, 8.6, 10.6) and (12, 13.4, 18.5 %) formation after 3 and 6 h of treatment respectively at concentrations 2.5, 5 and 10 µg/ml compared with controls 0.9 and 1.1% (Figure 6F–G, Supplementary Figure 2, available at Mutagenesis Online). ZnO NPs cause cell cycle arrest at G2/M phase To further understand the cellular effect of the ZnO NPs, we investigated their effect on cell cycle progression. Cell cycle arrest is a crucial component of the DDR, as it allows sufficient time for DNA repair to occur before progression into mitosis, thus protecting genome integrity. We examined the effect of ZnO NPs on the cell cycle using flow cytometry. Our data demonstrate that ZnO NPs involved in arresting the cells in G2/M phase in a time- and concentration-dependent manner. At treatment concentration of 10 µg/ml, cells were arrested 1.23-, 1.37- and 1.6-fold more than the control after 3-, 6- and 24-h treatment, respectively (Figure 7A–C, Supplementary Figure 3, available at Mutagenesis Online). However, the cell cycle arrest was not significant at concentrations 0.5, 1 and 2.5 µg/ml. The G2/M arrest was further confirmed by western blot analysis using specific antibody for pCHK1 S345, which has been reported to be a marker of G2/M arrest (36). Indeed, our data showed increase in the levels of S345 phosphorylation of Chk1 at 24 h (Figure 8). However, a significant reduction in the G2/M population was observed when cells were pre-treated with NAC, confirming the key role of ROS in eliciting DNA damage to HaCaT cells after the ZnO NPs treatment. Figure 7. Open in new tabDownload slide ZnO NPs treatment induces cell cycle arrest at G2/M phase: histogram representation of cell cycle phases after ZnO NPs treatment (A) 3 h and (B) 6 h. (C) Bar graph representing the percent of cells in G2/M phase exposed to 5 and 10 µg/ml ZnO NPs respectively. Figure 7. Open in new tabDownload slide ZnO NPs treatment induces cell cycle arrest at G2/M phase: histogram representation of cell cycle phases after ZnO NPs treatment (A) 3 h and (B) 6 h. (C) Bar graph representing the percent of cells in G2/M phase exposed to 5 and 10 µg/ml ZnO NPs respectively. Figure 8. Open in new tabDownload slide Western blot analysis of proteins involved in: (A) stress and apoptosis markers. (C) DNA double strand breaks/repair markers. (B and D) bar graphs exhibiting their densitometric analysis. β-ACTIN was used as internal control to normalize the data. Figure 8. Open in new tabDownload slide Western blot analysis of proteins involved in: (A) stress and apoptosis markers. (C) DNA double strand breaks/repair markers. (B and D) bar graphs exhibiting their densitometric analysis. β-ACTIN was used as internal control to normalize the data. Western blot analysis To get a holistic picture of the molecular changes occurring after treatment to ZnO NPs, we performed western blot analysis with antibodies against key proteins involved in stress response, apoptosis, cell cycle and DNA double-strand breaks. Oxidative stress is known to increase the levels of Heme Oxygenase I (HMOX1) (37) and HSP60 (38). Immunoblot analysis of HaCaT cells treated with ZnO NPs (2.5, 5 and 10 µg/ml) for 24 h showed a significant fold increase (P < 0.05) in the expression of HMOX1 (1.6, 2.2, 2.8) and HSP60 (1.7, 2, 2.5). DNA damage is known to activate the phosphorylation of p-p53 (39). Consequent to the genotoxic effect of the ZnO NPs, we also observed a significant increase in the levels of tumour suppressor protein p-p53 (1.67, 2.1, 2.47) after treatment to increasing concentrations of ZnO NPs (2.5, 5 and 10 µg/ml) respectively. The levels of cytochrome c (1.5, 2, 2.2), APAF1 (1.3, 2.6, 2.9), BAX (1.2, 1.6, 2.1), Caspase 9 (1.4, 1.6, 2.2) and Caspase 3 (1.6, 1.9, 2.5) increased with increasing concentrations of ZnO NPs (2.5, 5 and 10 µg/ml), whereas that of BCL2 (0.8, 0.5, 0.3) decreased confirming the activation of the apoptotic pathway in this setup (Figure 8A and B). We have observed potent genotoxic effects in HaCaT cells when exposed to ZnO NPs (Figure 6). This DNA damage (DNA strand break) will consequently activate the DDR pathway. Likewise, we observed significant increase in the levels of p-ATM (1.6, 2.1, 2.4) and ATR (1.2, 1.6, 2.1), which initiate the DDR cascade. To confirm the cell cycle arrest at G2/M phase, expression levels of pCHK1 S345 were assessed compared to control. In addition, the expression of γH2AX was increased 1.6-, 2.1-, 2.8-fold compared with control, which confirms the presence of DNA double-strand breaks when cells are exposed to the ZnO NPs. Consequently, the DNA repair pathways also get activated, which is corroborated by the increased expression of DNA-PK (1.4, 1.6, 2.2) compared with control under the same experimental setup. DNA-PK plays an important role in double-strand break repair by non-homologous end joining (40). We also observed decreased expression of CDC25A (0.8, 0.5, 0.2), CDC2 (0.8, 0.5, 0.3), Cyclin B1 (0.7, 0.5, 0.3) and cyclin D (1.2, 1.6, 2.1) compared with control, which are crucial cell cycle regulators (41–44) (Figure 8C and D). Discussion Engineered NPs show unique physicochemical properties, which make them entities of considerable interest. DNA damage and the subsequent response in cellular systems when exposed to the NPs need further evaluation as we are constantly exposed to nanoproducts in our environment. Also, the response elicited by different types and size of these nanoparticles is a subject of concern. In this context, the need of the hour is to address the biological damage induced by the NPs in relation to their specific properties. This study was carried out to gain deeper understanding of DNA damage and repair in human keratinocyte cells exposed to ZnO NPs in inducing genotoxicity. The different physicochemical properties of NPs are affected by size, shape, charge, etc. and may be responsible for different effects within biological system. Hence, the characterisation of the NPs is very important to determine the property of a particular NP (45). Different methods such as TEM and DLS have been used to determine the size of NPs both in dry and liquid state (21). Our data revealed that ZnO NPs were stable and mono-dispersed in culture media, thus confirming their suitability for toxicity studies. We observed that the zeta potential of ZnO NPs increased with time, which indicates their monodispersity in the treatment media. This could be due to the dynamic coating of protein or biomolecules over the surface of nanoparticles (known as corona), which plays an essential role in their biological fate (46–48). In this study, we demonstrate that ZnO NPs get internalised into the HaCaT cells using TEM (Figure 1) and flow cytometer. Distribution of the ZnO NPs inside cells may lead to eventual interactions with various macromolecules including proteins, lipids and nucleic acids ultimately leading to toxic effects in cellular system. Our data show that ZnO NP concentration at lower concentration 5 µg/ml can induce significant cytotoxicity after 6- and 24-h treatment (Figure 2). The cytotoxic effect was magnified with 5 and 10 µg/ml of ZnO NPs. This suggests that ZnO NPs cause cytotoxicity in HaCaT cells in both time and concentration-dependent manner. To understand the molecular mechanism of ZnO NP-induced cytotoxicity, we investigated the ROS generation in this context. Quantitative flow cytometric data showed that ROS was indeed produced in the cells after treatment with 5 and 10 µg/ml of ZnO NPs for both 3 and 6 h (Figure 3A–F). This was further confirmed by qualitative studies by fluorescence microscopy where a clear increase in the green fluorescence was observed with increasing concentrations of ZnO NPs (Figure 3G–K). This showed that ZnO NPs induced significant oxidative stress. The cellular stress was also confirmed by immunoblot analysis wherein we observed significant upregulation of HMOX1 and HSP60 (Figure 8A and B). ZnO NPs have been shown to induce the production of ROS even without photoactivation. To counteract ROS-induced cellular stress, heat shock proteins play a major role by inducing protein folding and minimising degradation. In line with this, our findings are in agreement with previous studies (21). To understand the consequences of this oxidative stress, we next investigated the effect on MMP as ROS is well known to depolarise the mitochondrial membrane (34,35). Indeed, our results show that ZnO NPs depolarise the mitochondrial membrane in time and concentration-dependent manner, the effect being more potent at 6 and 24 h (Figure 4A–G). This was also confirmed qualitatively wherein we observed a clear shift from red to green fluorescence with increasing concentrations of ZnO NPs (Figure 4H–P) confirming the depolarisation of mitochondrial membrane under these conditions. Depolarisation of the mitochondrial membrane is known to lead to apoptosis (34). Thus, we next used annexin V/PI binding assay to demonstrate apoptosis. As expected, we observed a significant increase in the annexin V/PI double positive cells (late apoptotic) by flow cytometry with ZnO NPs (2.5, 5 and 10 µg/ml) treatment after 6 h (Figure 5). Interestingly, very few cells were in the early apoptotic phase (annexin V positive, PI negative) under these conditions, which signifies the potent killing effect of ZnO NPs on HaCaT cells. In fact, increasing the time of treatment actually caused an increase in the population of necrotic cells (data not shown). We next investigated the DNA fragmentation after ZnO NP treatment and in line with the earlier results, we observed significant fragmentation of DNA compared with control after staining with acridine orange (Figure 5E–H). All these results confirm the involvement of apoptotic pathway in the context of the effect of ZnO NP on HaCaT cells. This was further corroborated by western blot analysis. Indeed, we observed significant upregulation of cytochrome c and APAF1 (Figure 8A and B) in the cytosol, which signifies the export of cytochrome c from mitochondria to the cytosol due to mitochondrial membrane depolarisation and formation of apoptosome complex. Increase in the levels of BAX (pro-apoptotic protein), Caspase-3 and 9 with concomitant decrease in the BCL2 levels (Figure 8A and B) all confirm the activation of apoptotic signalling. We surmise that increased expression of BAX further disrupts the mitochondrial transmembrane potential and released the pro-apoptotic proteins from the mitochondrial intermembrane space into the cytoplasm. Cytochrome c further activates the formation of apoptosome and activation of Caspase-9, which in turn mediates the activation of effector caspases resulting in apoptosis. Our results clearly demonstrate the activation of mitochondria-mediated pathway in ZnO NPs-induced apoptosis in HaCaT cells. Further, we investigated the effect of these ZnO NPs on the genetic material. In this line, we observed DNA damage in HaCaT cells after treatment with ZnO NPs. Comet assay revealed that increasing concentrations of ZnO NPs cause increase in the OTM and tail length signifying DNA damage. We further observed that ZnO NPs induced MN formation at increasing concentrations, which establishes the potent genotoxic effect of these NPs. The single- and double-stranded DNA breaks can be estimated by Comet assay whereas the occurrence of MN indicates DNA double-strand breaks (49). ROS production leads to DNA double-strand breaks causing activation of the ATM-Chk2 signalling pathway (50). ATM belongs to a family of kinases that play an important role in the checkpoint signalling network via either RAD3 related or DNA-dependent protein kinase (51). In our study, we also observed a concentration-dependent increase in the expression of ATM after 24 h of ZnO NPs treatment (Figure 8C and D), suggesting an increase in DNA double-strand breaks. On the contrary, the increase in expression of ATR was not upto the level of ATM. Moreover, our observations for double-strand DNA breaks were further confirmed by the increased expression of γH2AX, a sensitive biomarker for DNA double-strand break detection (52). H2AX is one of the several genes coding for histones, which is a major component for nucleosome-formation and structure of DNA (53). These findings suggest that treatment to ZnO NPs activates the ATM/Chk2 DNA damage signalling pathway in HaCaT cells. A significant concentration-dependent increase in the expression profile of tumour suppressor protein p53 was also observed in ZnO NPs-treated HaCaT cells. p53 is involved in cell cycle arrest, DNA repair and apoptosis in the context of genotoxic and oxidative stress (54) and DNA damage is known to induce phosphorylation of p53 at ser15 (55,56). Likewise, we sought to investigate p53 activation, its relationship with ROS and whether p53 is required for ZnO NPs-induced apoptosis. We observed an enhanced phosphorylation of p53 at ser15 in HaCaT cells after treatment to ZnO NPs. This activation may also be explained as a response to DNA damage, which is caused by ZnO NPs-induced ROS. To understand the effect of the earlier mentioned molecular pathways in the context of cell cycle, we also investigated the levels of certain cell cycle regulatory proteins by western blot analysis. In our experimental setup, there was a concentration-dependent increase in the expression of CdC2 as well as a significant decrease in the expression of Cyclin B1. The regulatory protein Cyclin B1 is involved in mitosis and binds with p34 (Cdk1) to form the maturation-promoting factor. Cyclin B1 is expressed during the G2/M phase of the cell cycle and promotes cell division (57,58). In our experimental setup, we observed a concentration-dependent decrease in Cyclin B1 expression, which suggested towards the onset of G2/M arrest in response to ZnO NP treatment. This was further confirmed by western blot analysis wherein we observed a dose-dependent increase in the levels of pCHK1 (S345), which is a specific marker for G2/M arrest (36). This finding confirms the occurrence of G2/M arrest in HaCaT cells when exposed to ZnO NPs. An illustration depicting the mechanisms involved in toxicity of ZnO NPs in HaCaT cells is shown in Figure 9. There are several reports in the literature about the toxic effects of ZnO NPs in different cell lines. To add novelty to our study, we tried to investigate the effect of ZnO NPs on HaCaT cells in terms of molecular effects, genotoxicity and cell death potential with an integrative approach. Taken together, our findings from this study suggest that ZnO NPs, which are blatantly used in many industries having direct relationship to human health, have significant toxic effects on HaCaT cells (human keratinocyte cell line). These NPs cause ROS generation in HaCaT cells leading to mitochondrial membrane depolarisation, which causes activation of apoptotic pathway. Consequently, these NPs also have a significant genotoxic effect, which may lead to increased frequency of somatic cell mutation. Thus, we firmly believe that due caution should be exercised while using ZnO NPs such that the advantages of its usage is not undermined by its toxic effects. Figure 9. Open in new tabDownload slide Proposed pathway illustrating DNA damage response and related protein markers involvement after ZnO NPs treatment to HaCaT cell.. Figure 9. Open in new tabDownload slide Proposed pathway illustrating DNA damage response and related protein markers involvement after ZnO NPs treatment to HaCaT cell.. Conclusion In summary, ZnO NPs internalisation in HaCaT cells elevated ROS levels leading to oxidative stress within 3 h of treatment. Results indicated significant generation of radicals at concentration 5 and 10 µg/ml compared with control. In addition, oxidative damage triggered the mitochondrial-mediated apoptotic pathway followed by DNA fragmentation. Mitochondrial membrane depolarisation was assessed from the conversion of JC-1 dye aggregates to monomers (red to green fluorescence shift) with an increase of 11.4 % than non-treated cells (2.6 %) after 6 h at 10 µg/ml. To fill the lacuna in understanding the mechanism involved in apoptosis and DNA damage, increase in apoptotic cells, cell cycle profile and MN formation was studied. Results explained double-strand breaks lead to the cell cycle arrest at G2/M phase and induction of apoptosis. Further to confirm the mechanism, key markers involved in apoptosis (BAX, Caspase 3 and 9, APAF1 and BCL2), DNA damage (ATM, ATR and pCHK1) were studied and found their expression elicited apoptosis and DNA damage (G2/M arrest) in HaCaT cells. Thus, the mechanistic approach paved a clear picture in demonstrating the genotoxic potential of ZnO NPs and pathways involved in apoptosis and DNA damage. Funding Funding received from the Department of Biotechnology, Government of India under the project “NanoToF: Toxicological evaluation and risk assessment on Nanomaterials in Food” (grant number BT/PR10414/PFN/20/961/2014) and DST SERB Project “Nanosensors for the Detection of Food Adulterants and Contaminants” (grant number EMR/2016/005286) is gratefully acknowledged. Financial assistance by The Gujarat Institute for Chemical Technology (GICT) for the Establishment of a Facility for environmental risk assessment of chemicals and nanomaterials is also acknowledged. Conflict of interest statement: Authors declare no conflict of interest. References 1. Dobrucka , R. and Długaszewska , J . ( 2016 ) Biosynthesis and antibacterial activity of ZnO nanoparticles using Trifolium pratense flower extract . Saudi J. Biol. Sci. , 23 , 517 – 523 . Google Scholar Crossref Search ADS PubMed WorldCat 2. 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TI - ZnO nanoparticles-associated mitochondrial stress-induced apoptosis and G2/M arrest in HaCaT cells: a mechanistic approach
JF - Mutagenesis
DO - 10.1093/mutage/gez017
DA - 2019-09-20
UR - https://www.deepdyve.com/lp/oxford-university-press/zno-nanoparticles-associated-mitochondrial-stress-induced-apoptosis-bEteEYBM64
SP - 265
VL - 34
IS - 3
DP - DeepDyve
ER -