Article pubs.acs.org/crt

DNA Melting and Genotoxicity Induced by Silver Nanoparticles and Graphene Angela Ivask,*,† Nicolas H. Voelcker,† Shane A. Seabrook,‡ Maryam Hor,§ Jason K. Kirby,∥ Michael Fenech,§ Thomas P. Davis,⊥,# and Pu Chun Ke*,⊥ †

ARC Centre of Excellence in Convergent Bio-Nano Science and Technology, Mawson Institute, University of South Australia, GPO Box 2471, Adelaide, SA 5001, Australia ‡ CSIRO Manufacturing Flagship, 343 Royal Parade, Parkville, VIC 3052, Australia § CSIRO Health Sciences and Nutrition Flagship, Kintore Avenue, Adelaide SA 5000, Australia ∥ CSIRO Land and Water Flagship, Waite Road-Gate 4, Glen Osmond, SA 5064, Australia ⊥ ARC Centre of Excellence in Convergent Bio-Nano Science and Technology, Monash Institute of Pharmaceutical Sciences, Monash University, 381 Royal Parade, Parkville, VIC 3052, Australia # Department of Chemistry, Warwick University, Gibbet Hill, Coventry, CV4 7AL, United Kingdom S Supporting Information *

ABSTRACT: We have revealed a connection between DNA-nanoparticle (NP) binding and in vitro DNA damage induced by citrate- and branched polyethylenimine-coated silver nanoparticles (c-AgNPs and b-AgNPs) as well as graphene oxide (GO) nanosheets. All three types of nanostructures triggered an early onset of DNA melting, where the extent of the melting point shift depends upon both the type and concentration of the NPs. Specifically, at a DNA/NP weight ratio of 1.1/1, the melting temperature of lambda DNA dropped from 94 °C down to 76 °C, 60 °C, and room temperature for GO, c-AgNPs and b-AgNPs, respectively. Consistently, dynamic light scattering revealed that the largest changes in DNA hydrodynamic size were also associated with the binding of b-AgNPs. Upon introduction to cells, b-AgNPs also exhibited the highest cytotoxicity, at the half-maximal inhibitory (IC50) concentrations of 3.2, 2.9, and 5.2 mg/ L for B and T-lymphocyte cell lines and primary lymphocytes, compared to the values of 13.4, 12.2, and 12.5 mg/L for c-AgNPs and 331, 251, and 120 mg/L for GO nanosheets, respectively. At cytotoxic concentrations, all NPs elicited elevated genotoxicities via the increased number of micronuclei in the lymphocyte cells. However, b-AgNPs also induced micronuclei at subtoxic concentrations starting from 0.1 mg/L, likely due to their stronger cellular adhesion and internalization, as well as their subsequent interference with normal DNA synthesis or chromosome segregation during the cell cycle. This study facilitates our understanding of the effects of NP chemical composition, surface charge, and morphology on DNA stability and genotoxicity, with implications ranging from nanotoxicology to nanobiotechnology and nanomedicine.

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INTRODUCTION

For the biological and medicinal applications of nanomaterials, such as the design and in vivo deployment of nanodevices and nanocarriers, as well as for the general practice of safe nanotechnology, there is a crucial need to understand the fate and behavior of engineered nanomaterials in living systems.5 The biological effects associated with NPs are closely tied to the interactions between the engineered NPs and biomolecules. Among these, the interactions between NPs and proteins have been shown to render a generic entity termed “protein corona”,6 where proteins adsorb onto a NP “core” in a dynamic and competitive manner. Sorption of proteins onto NPs likely compromises the structure and function of the

The recent development of nanotechnology has inspired the design and engineering of nanomaterials of diverse chemical origin and physicochemical properties.1 Silver nanoparticles (AgNPs), for example, represent a major class of nanomaterials known for their superior antibacterial properties.2,3 The potential of AgNPs to generate surface plasmon resonance (SPR) under light excitation has further enabled their use in chemical and molecular sensing. Another major class of emerging nanomaterials, graphene and its derivatives such as graphene oxide (GO), possesses large surface area, superior mechanical strength, and high electron carrying capacity, and has shown promise for electronics, molecular assembly, chemical sensing, as well as environmental remediation.4 © 2015 American Chemical Society

Received: January 30, 2015 Published: March 17, 2015 1023

DOI: 10.1021/acs.chemrestox.5b00052 Chem. Res. Toxicol. 2015, 28, 1023−1035

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Chemical Research in Toxicology proteins7−9 and further determines the biological identity of the NPs. In contrast to the extensive literature on NP−protein interactions, the effects of NPs on other classes of biomolecules are considerably less well studied. For example, although NP− DNA interactions are of great relevance to the design and application of DNA nanotechnology and toxicology,10 research on NP-DNA binding is surprisingly scarce. Among the studies available, for example, SPR spectroscopy and surface enhanced Raman spectroscopy (SERS) revealed that DNA bases could induce differential aggregation of AgNPs.11 Functionalized with complementary oligonucleotides, AgNPs assembled to form DNA-linked NP networks, thus paving the way for plasmonic and nanoelectronic devices.12 Single-stranded DNA (ssDNA) self-assembled into two distinct crossover patterns of small spherical particles and elongated networks on the surface of graphite, driven by the competitive processes of nucleobase-tonucleobase and nucleobase-to-graphene π−π stacking.13 The adsorption and desorption of ssDNA oligonucleotides to GO nanosheets14 was mediated by electrostatic repulsion and hydrophobic interactions, further influenced by environmental factors such as cations, pH, and nature of the solvent. Furthermore, it has been reported that GO nanosheets, treated additionally with nitric acid and sulfuric acid, stabilized ssDNA from the cleavage of DNase I, thereby providing a novel DNAgraphene platform for biotechnology and biomedical applications.15 The binding of DNA to NPs may also result in the disruption of the genetic material in living cells. Such studies are, however, often inconsistent or contradictory. For example, Nam et al.16 did not detect any induction of DNA repairing enzymes by gold, silver, zinc, and titanium dioxide NPs. Yet, a number of studies revealed the disruption of genetic material as indicated by the formation of micronuclei, when human cells were exposed to NPs. Specifically, AshaRani et al.17 showed that AgNPs at 100 μg/mL induced the formation of micronuclei due to the disruption to genetic material division in U251 glioblastoma cells and IMR-90 fibroblasts. Vecchio et al.18 reported that the occurrence of micronuclei in human lymphocytes by citrate- and PVP-coated AgNPs was dependent upon the primary size of the NPs, where smaller NPs exhibited stronger genotoxic effects. Interestingly, the occurrence of genotoxicity has also been demonstrated for carbon-based nanomaterials. For example, Niwa et al.19 and Mrdanovic et al.20 observed the onset of micronuclei after the exposure of CHO (Chinese hamster ovary), HeLa (derived from cervical cancer cells), and HEK291 (human embryonic kidney) cells to very low concentrations of fullerols (C60(OH)24). Akhavan et al.21 reported that GO nanosheets exhibited genotoxic effects to human mesenchymal stem cells at 0.1 mg/mL. Similarly, carbon nanotubes have been shown to induce genotoxicity in mammalian cells starting from 0.1 mg/mL.22 While a handful studies on the genotoxic effects of nanomaterials are available, none of these studies has elucidated the possible connection between the DNA binding properties of NPs and their genotoxic potential. Accordingly, the purpose of this study is to describe the phenomena associated with NP− DNA interactions in aqueous solutions and correlate these phenomena with NP genotoxic potential in vitro. Specifically, citrate- and branched polyethylenimine (bPEI)-coated AgNPs (c-AgNPs and b-AgNPs) as well as GO nanosheets are analyzed for their capacities to bind phage lambda DNA and their abilities to induce micronuclei in human lymphocyte cells.

This design takes into consideration the effects of surface charge, coating, and geometry of NPs in impacting their physical and biological interactions with genetic material. To infer the binding effectiveness of NPs and lambda DNA, the hydrodynamic size of the NPs in the absence and presence of DNA is determined by dynamic light scattering (DLS). The thermal stability of lambda DNA in the presence of the three types of nanostructures, each of different molar ratios, is then examined using the high-throughput technique of differential scanning fluorimetry (DSF), a method we have recently first employed to investigate the NP−protein corona.23 To assess the genotoxic potential of the three nanostructures, a cytokinesis-block micronucleus assay in conjunction with a cellular viability study was carried out with human B- and Tlymphocyte cell lines as well as with human primary lymphocytes. Formation of micronuclei was selected as a measure of potential genotoxicity due to its sensitive response to various abnormalities in chromosomal segregation.24 The results from the NP−DNA physical interactions and cellular studies are compared to draw their possible connections.

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EXPERIMENTAL AND COMPUTATIONAL METHODS

Nanoparticles. BioPure citrate-coated silver nanoparticles (cAgNPs) (1 mg/mL, in water) and branched PEI (bPEI)-coated silver nanoparticles (b-AgNPs) (1 mg/mL, in water) of 20 nm in nominal diameter were purchased from NanoComposix. Both NP stock suspensions appeared homogeneous and free from precipitation when stored at 4 °C. Single-layered GO nanosheets (2 mg/mL, in water) of 20 μm in nominal size were purchased from Sigma-Aldrich. The stock GO suspension appeared dark brown and free from precipitation at room temperature. For TEM imaging, NPs suspensions were diluted to 0.1 mg/L (c-AgNPs and b-AgNPs) or 0.2 mg/L (GO nanosheets) with Milli-Q water. For zeta-potential measurements, the NPs stock suspensions were diluted down to 0.2 mg/mL with Milli-Q water. For NP-DNA studies, all three types of nanostructures were diluted to 0.2 mg/mL in 10 mM Tris-HCl (pH 7.6) before use. For toxicity and genotoxicity studies, NPs were diluted in cell culture medium (see the composition below) to 0.05 mg/mL and vortexed. The diluted NPs appeared precipitation free at room temperature. NP Size and Surface Charge. The zeta potentials of the three types of nanostructures were determined from 0.2 mg/mL NP suspensions using a Nicomp 380 Particle Sizing System. The hydrodynamic sizes of the NPs dispersed in 10 mM Tris-HCl or NPs that were incubated for 24 h in cell culture medium at 37 °C in 5% CO2 atmosphere (mimicking the conditions for cellular tests) were measured using a Zetasizer Nano S90 (Malvern Instruments) or Nicomp 380 particle sizing system. All of the measurements were performed at room temperature. XPS. The elemental composition within 1−10 nm of the surface of the GO was quantitatively determined by X-ray photoelectron spectroscopy (XPS). Droplets of GO suspension were pipetted onto a PTFE sample holder and allowed to dry before introducing the sample into the spectrometer. XPS analysis was performed using an AXIS Ultra DLD spectrometer (Kratos Analytical Inc., Manchester, UK) with a monochromated Al Kα source at a power of 96 W (12 kV × 8 mA), a hemispherical analyzer operating in the fixed analyzer transmission mode, and the standard aperture (analysis area: 0.3 mm × 0.7 mm). Survey spectra were acquired at a pass energy of 160 eV. To obtain more detailed information about chemical structure and oxidation states, high resolution spectra were recorded from individual peaks at 20 eV pass energy (yielding a typical peak width for polymers of 1.0 eV). Each specimen was analyzed at an emission angle of 0° as measured from the surface normal. Assuming typical values for the electron attenuation length of relevant photoelectrons, the XPS analysis depth 1024

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Chemical Research in Toxicology (from which 95% of the detected signal originates) ranges between 5 and 10 nm for a flat surface. Data processing was performed using CasaXPS processing software version 2.3.15 (Casa Software Ltd., Teignmouth, UK). All elements present were identified from survey spectra. The atomic concentrations of the detected elements were calculated using integral peak intensities and the sensitivity factors supplied by the manufacturer. Binding energies were referenced to the main C 1s peak at 285.0 eV. Microscopy Imaging of NPs. For TEM imaging, a 5 μL drop of 0.1 mg/mL suspension of c-AgNPs or b-AgNPs or of 0.2 mg/mL of GO nanosheets was pipetted onto carbon-coated 300-mesh copper grids. After 10 min of adsorption, excess samples were drawn off using a filter paper, and the samples were further dried overnight. The samples were examined using a JEOL JEM2100F Transmission Electron Microscope at an operating voltage of 200 kV. The sizes of primary particles were measured using Gatan Microscopy Suite Software. GO nanosheets were also imaged under a bright field microscope (Leica DMIRB) by pipetting and drying 10 μL of 0.1 mg/ L GO suspension onto a microscope slide. Dissolution of AgNPs. Dissolution of AgNPs was determined in cell culture medium (see the composition below). Eight milliliters of cAgNPs at 15 μg/mL and b-AgNPs at 10 μg/mL concentrations in cell culture medium were incubated for 24 h at 37 °C and 5% CO2. Four milliliters of the NP suspensions was filtered using Amicon Ultra-4 filter units with 3 kDa molecular weight cutoff and then acidified (2% HNO3). Four milliliters of the NP suspensions was acidified without filtration. Ag content in acidified suspensions was analyzed with ICPMS in ALS (Melbourne). To take into account the potential sorption of Ag ions onto membranes, a similar filtration procedure was carried out with AgNO3 at a concentration of 5 μg/mL. Attachment of Nanoparticles to Lambda DNA. Lambda DNA (48,502 nucleotides, 0.3 mg/mL in 10 mM Tris-HCl, and 1 mM EDTA, pH 7.6) was acquired from Thermal Scientific. This Escherichia coli bacteriophage virion DNA is linear and double-stranded with 12 bp single-stranded complementary 5′-ends. For DNA binding assays, the DNA stock was diluted to 0.1 mg/mL with 10 mM Tris-HCl (pH 7.6) and mixed with 0.2 mg/mL suspensions of NPs at different DNA/ NP ratios. The hydrodynamic sizes of lambda DNA, c-AgNPS, bAgNPs, GO, as well as the three DNA−NP mixtures each at a 1:1 DNA (0.1 mg/mL)/NP (0.2 mg/mL in 10 mM Tris-HCl buffer) volume ratio were determined using an automated, high-throughput, temperature-controlled plate reader device (DynaPro Plate Reader, Wyatt) and black 384-well plates (Thermo Fisher). The total volume of each sample well was 20 μL. Prior to the DLS measurements, plates with the samples were incubated for 2 h and spun for 1 min at 1,000 rpm/164g (Centrifuge 5804, Eppendorf). Each sample well was then topped with 10 μL of glycerol to prevent volume loss and concentration variations throughout the measurement. To ensure repeatability and statistics, each sample condition was measured in triplicate. DNA Melting Assay. White PCR plates (96-well; Thermo Fisher AB0600/W) were used for the differential scanning fluorimetry (DSF) measurement that has recently been applied to the study of protein binding with a fullerene derivative,23 to determine the melting temperature of lambda DNA in the presence of the three types of NPs of increasing concentrations. For the controls of DNA (0.1 mg/mL), c-AgNPs, b-AgNPs, and GO nanosheets (each at 0.2 mg/mL concentration), the sample volume was fixed at 19.7 μL. For the DNA−NP mixtures, the volume of the DNA was 16.7, 13.7, 10.7, and 7.7 μL, while the corresponding volume of NPs was 3, 6, 9, and 12 μL, to ensure the total volume of each sample fixed at 19.7 μL. To evaluate the effect of Ag ion release on DNA melting, an AgNO3 stock of 0.2 mg/mL was prepared in 10 mM Tris-HCl buffer. For the DNA−silver ion mixtures, the ratios of the DNA (0.1 mg/mL) and AgNO3 (0.2 mg/mL) were adjusted from 19.7 μL/0 to 16.7 μL/3 μL, 13.7 μL/6 μL, and 10.7 μL/9 μL. After 2 h of incubation, 0.3 μL of 10% of SYBR Green dye (Ex/Em: 497 nm/520 nm) was added to each sample well by an automated liquid dispenser (Phoenix, Art Robbins Instruments) to top the total volume to 20 μL. SYBR Green intercalates with double-stranded DNA to emit fluorescence. Plates with the samples

were spun for 1 min at 1,000 rpm/164g (Centrifuge 5804, Eppendorf), then loaded into a Bio-Rad CFX96 optical reaction module attached to a C1000 thermal cycler, running the Bio-Rad CFX Manager 3.0 software. The thermal cycler was first set to rise to 90 °C for 10 min to remove concatemers due to base pairing of the 12 bp single-stranded ends of the lambda DNA. The thermal cycler was then cooled down to 20 °C and held at room temperature for 1 min, before beginning a stepped heating cycle from 20 to 100 °C, of 0.5 °C every 5 s. At the end of each heating step, the optical reaction module collected data. The data were automatically processed at the end of each experiment using the CFX Manager software. To ensure repeatability and statistics, each sample was measured in triplicate. Maintenance of Human Lymphocyte Cells. Human Blymphocyte cell line WIL2-NS (ATCC CRL-8155) and human Tlymphocyte cell line JURKAT (ATCC TIB-152) were routinely maintained in RPMI 1640 medium (Sigma-Aldrich) containing 4.5 g/ L glucose and supplemented with 50 μM glutamine, 10% FBS, 1 mM Na-pyruvate, 10 mM HEPES buffer, 100 U/mL penicillin, and 100 μg/ mL of streptomycin (further designated as the cell culture medium). Cells were grown on 140 mm diameter cell culture-treated Petri dishes (NUNC) in a controlled humidified atmosphere with 5% of CO2, at 37 °C. The culture medium was changed every 3 days. Human primary lymphocytes were separated from whole blood using Ficoll-Paque PLUS (GH Health Care). Briefly, 1/4 fraction of Ficoll-Paque was pipetted onto the bottom of a 50 mL centrifugation tube, 3/4 fraction of blood collected using 9 mL Plasma Lithium Heparin collection tubes (Greiner), and diluted 1:1 with Hank’s balanced salt solution (HBSS, Sigma) carefully added to the top. The tubes were centrifuged at 410g for 30 min. The lymphocyte layer on top of the HBSS buffer layer was collected and washed twice with 10−15 mL of HBSS. The cells were centrifuged at 190g for 10 min between washes. Primary lymphocytes were suspended in RPMI 1640 medium as described above but without antibiotics. Cells were diluted until 1 × 106 cells/ mL, and 30 μg/mL of hemagglutinin was added to promote cellular proliferation. Toxicity Assay. The cell density of B and T-lymphocyte cell lines and blood lymphocytes was adjusted to 1 × 106 cells/mL. Cell culture medium was used to dilute lymphocyte cell lines and cell culture medium without antibiotics, 30 μg/mL of hemagglutinin was used to dilute blood lymphocytes. Fifty microliters of cell suspension was pipetted onto transparent 96-well microplates, and 50 μL of NPs suspension prepared in cell culture medium was added. c-AgNPs and b-AgNPs were introduced at concentrations between 0.05 and 50 μg/ mL, and GO nanosheets were added at concentrations between 6.25 and 400 μg/mL. Cell culture medium was brought to cells in nonexposed control wells, and H2O2 between concentrations 10 and 1000 μM was introduced as a positive control. The cells with test chemicals on 96-well plates were incubated at 5% of CO2 and 37 °C for 24 h (lymphocyte cell lines) or 44 h (primary lymphocytes). Then, resazurin dye dissolved in PBS buffer (pH 7.4) at a final concentration of 30 μg/mL was added, and the cells were further incubated at 5% of CO2 and 37 °C for 4 h. After that, fluorescence of resorufin, a metabolic product of resazurin forming in physiologically active cells, was measured at Ex/Em wavelengths 530/590 nm using a Synergy HTX (BioTek) plate reader. The cell viability at each NP concentration was calculated as % of the nonexposed control. Halfmaximal inhibitory concentration (IC50) values were calculated using the program GraphPad. Cytokinesis-Block Micronucleus Assay for Genotoxicity. The assay was carried out essentially as described by Fenech34 and in the OECD guidelines.43 The cell densities and exposure conditions were similar to those used in the toxicity assay (see above) except that 96well black-walled and clear bottomed Cell Carrier plates (PerkinElmer) were used. After 24 h of exposure, plates with lymphocyte cell lines were centrifuged (180g, 5 min, Heraeus centrifuge equipped with plate centrifugation units), and fresh media with 4.5 μg/mL of cytochalasin B (cytB, the chemical was first dissolved in DMSO to 600 μg/mL and then diluted in cell culture medium) were added. The primary lymphocytes were exposed to test chemicals for 44 h after which cytB at a concentration of 4.5 μg/mL was added. Both the lymphocyte cell 1025

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Figure 1. Characteristics of the tested nanostructures. (A,B) TEM images of c-AgNPs and b-AgNPs, and (C) TEM image (left) and bright field image (right) of GO sheets. Scale bars in (A,B) 50 nm, (C) 200 nm (left), and 10 μm (right). Primary sizes, surface charge (measured as zeta (z)potential), and hydrodynamic diameters of the NPs (per DLS analysis) in stock suspension, buffer for DNA melting studies, and in cell culture medium.

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lines and primary lymphocytes were then incubated for another 24−28 h at 5% of CO2 and 37 °C. Then, lymphocyte cell lines were labeled with fluorescent dyes. The cells were centrifuged and suspended in HBSS, and 5 μL/mL Vybrant DiO cell-labeling solution (Life technologies) was added. The cells were stained by incubation for 15 min at room temperature, then washed with HBSS, suspended in cell culture medium, and incubated at 5% of CO2 and 37 °C for 10 min. The cells were centrifuged again, added with 3.7% formaldehyde in phosphate buffered saline (PBS, Sigma), and incubated for 10 min. After centrifugation, the cells were permeabilized by suspension in 0.3% Triton-X 100 in PBS for 10 min. The cells were centrifuged again and resuspended in 0.12 μg/mL of Hoechst 33342 solution in PBS. After 10 min of incubation, the cells were centrifuged and suspended in PBS buffer. The cells were analyzed for mononucleated, binucleated, and multinucleated cells and for micronuclei under a fluorescent microscope using 20 and 40× objectives (Nikon fluorescent microscope; DAPI and FITC filters were used along with bright field imaging). The cells were also analyzed using the Operetta High Content Imaging System and Harmony software, version 3.1 (PerkinElmer). The primary lymphocytes were stained using Hemacolor stains (Merck). Specifically, 100 μL of cells was deposited onto a microscope glass slide using the Cytospin system (Thermo Scientific). The glass slide was then dried for 10 min at room temperature, dipped for 10 min into the Hemacolor fixing solution (Merck), quickly dried, then dipped 10 times in Hemacolor red reagent following by 6 dips in the Hemacolor blue reagent. The slides were then rinsed with deionized water and dried overnight. A coverglass was mounted using the DEPEX Mounting Medium and mononucleated, binucleated, and multinucleated cells as well as micronuclei were counted under a 100× objective (Nikon Eclipse 6300 microscope). From each treatment, 500 cells were counted (in the case of toxic concentrations, the cell count was lower, down to 100 cells per treatment). Scoring criteria were similar to those described by Fenech.34 The nuclear division index (NDI) was based on mono-, bi-, and multinuclear cells, while no apoptotic or necrotic cells were taken into account. NDI = (mononuclear cells + 2 × binuclear cells + 3 × multinuclear cells)/ (total number of cells). Also, the number of micronuclei per 500 binuclear cells was calculated. A one-tailed t test was carried out to find the significant differences (p < 0.05) between the number of micronuclei in the nonexposed control and NP exposed cells.

RESULTS AND DISCUSSION Characterization of AgNPs and GO Nanosheets. For this study, two types of AgNPs and GO nanosheets were chosen. The AgNPs employed citrate (c-AgNPs) and branched polyethylene imine (bPEI) (b-AgNPs) as surface-binding ligands. According to our recent data, these ligands are physically adsorbed on the surface of AgNPs through van der Waals and electrostatic forces.25,26 The zeta potential of the nanostructures was determined as −35.0 mV for c-AgNPs and +40.3 mV for b-AgNPs. TEM imaging revealed that both the cAgNPs and b-AgNPs were approximately 20 nm in diameter (Figures 1A and B), in agreement with the manufacturer’s specifications. The hydrodynamic diameter of the NPs determined by DLS was 18.0 nm for c-AgNPs and 27.9 nm for b-AgNPs, accordingly (Figure 1). This discrepancy between the hydrodynamic sizes of the two types of AgNPs could be due to their differences in wetting, as well as the different charges and binding affinities of the linear anionic citrate and branched cationic bPEI for the AgNP core. In Tris-HCl buffer, which was used in the DNA binding study, the hydrodynamic size of the NPs was slightly increased to 22.8 and 43.8 nm for c-AgNPs and b-AgNPs, respectively, likely due to the mediation of ions between the NPs to cause slight aggregation. In cell culture medium where the NPs were dispersed for toxicity and genotoxicity studies, the NPs appeared even larger, at 41.8 and 323 nm, respectively, for c-AgNPs and b-AgNPs, likely due to the formation of biomolecular coronas by the amino acids and proteins in the medium around the NPs27 (Figure 1). Unlike AgNPs, the surface of GO nanosheets consists of a sp2-hybridized carbon nanostructure covalently functionalized with hydroxyls and oxygens, of one to a few layers suspended in aqueous solutions. According to XPS analysis (Figure S1, Supporting Information), carbon and oxygen (atomic percentage to total carbon: 46.7%) were the main elements of the GO surfaces, but sulfur (atomic percentage to total carbon: 2%) and a trace amount of manganese (0.5%) were also present (Table S1, Supporting Information), consistent with the product specifications. The zeta potential of the GO nanosheets was −21.0 mV. The GO nanosheets appeared irregular in shape and 1026

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When DNA was mixed at a 1:1 weight ratio with b-AgNPs (0.2 mg/mL, initial hydrodynamic radius 21.9 nm; Figure 2D), the mixture displayed a broad distribution in DLS, with two peaks occurring at 28.4 and 138.5 nm (Figure 2E). This suggests that a significant fraction of the b-AgNPs was bound to DNA due to the electrostatic attractions between the negatively charged DNA phosphate backbone and the cationic bPEI coating of the b-AgNPs. In comparison, when GO nanosheets (initial hydrodynamic radius 8.3 μm, Figure 2F) were mixed with DNA at an equal volume, DNA-GO displayed a main peak at 4.2 μm in hydrodynamic radius, characterized by a wide size distribution up to tens of nanometers (Figure 2F). Since DNA and GO nanosheets repel due to their like charges, such changes in hydrodynamic size indicate an increased heterogeneity of the DNA−GO mixture as well as an improved dispersion of the GO nanosheets mediated by the DNA. Effect of NPs on DNA Melting. Prior to examining the effects of NPs on DNA melting, we determined the melting temperatures of DNA and NPs alone. As measured by DSF, a high-throughput technique which utilizes an intercalating SYBR Green dye as a reporter of DNA unzipping, the melting temperature of the lambda DNA control was at 94 °C (Figure 3). Also as shown in Figure 3, the controls of c-AgNPs, bAgNPs, and GO all displayed negligible fluorescence baselines, thereby validating the DSF method. Addition of c-AgNPs to DNA at a DNA/NP volume ratio of 16.7:3 resulted in a drop of the DNA melting temperature from 94 to 84 °C and further to 60 °C when the DNA/NP volume ratio decreased to 13.7:6 (Figure 3A). This indicates an augmented destruction to the double-stranded structure of lambda DNA by the increasing presence of the c-AgNPs. Further decreases of the DNA/NP volume ratio shifted the onset of DNA melting down to room temperature, as reflected by the disappearance of plateaus in SYBR Green fluorescence (Figure 3A). In comparison with c-AgNPs, the effect of bAgNPs on DNA melting was significantly more pronounced (Figure 3B). Specifically, at the lowest added concentration of b-AgNPs (DNA/NP volume ratio 16.7:3, or 0.08 μg/mL DNA to 0.03 μg/mL b-AgNPs) the DNA melting temperature dropped from 94 to 36 °C, and further decreases in the DNA/ NP volume ratio (or increases in the b-AgNP concentration) resulted in DNA melting occurring at room temperature (Figure 3B). Because of the possibility of AgNP dissolution,28 we also analyzed the effects of Ag ions on DNA melting temperature. When DNA/Ag ratios similar to those for AgNPs were used for AgNO3, only minimal changes in DNA melting, or ±2 °C compared with the control, was observed (Figure S2, Supporting Information). Thus, we concluded that silver ion release from c-AgNPs or b-AgNPs was an insignificant factor for elevated DNA melting in the presence of b-AgNPs and cAgNPs. Per weight basis, GO nanosheets appeared to induce comparable but slightly less drastic changes than c-AgNPs in DNA melting (Figure 3C versus B). Specifically, addition of GO sheets at a DNA/GO volume ratio of 16.7:3 caused the DNA melting temperature to drop from 94 to 88 °C. Further decrease in the DNA/NP ratio to 10.7:9 shifted the DNA melting temperature down to 47 °C, and at even lower DNA/ NP ratios, DNA melting started to occur at room temperature (Figure 3C). We propose the following reasons for decreased DNA melting temperatures in the presence of NPs. In the case of bAgNPs, specifically, DNA melting was accelerated by strong

inhomogeneous in size, ranging from a few to tens of micrometers in each dimension (Figure 1C). The hydrodynamic diameters of GO nanosheets in stock suspension, TrisHCl (used for the DNA binding study), and cell culture medium (used for the cellular toxicity studies) were 8.3, 19, and 16.8 μm, respectively (Figure 1). In contrast to AgNPs, the polydispersity of the GO nanosheets was remarkably high (see example in Figure 2F), indicating their highly heterogeneous size distribution.

Figure 2. Hydrodynamic sizes of NPs, DNA, and NP-DNA mixtures at 20 °C. Hydrodynamic radii of NPs alone (B,D,F), lambda DNA alone (A), or NP/DNA mixtures (C,E,G). For hydrodynamic size measurement, the volume ratio of all DNA-NP mixtures in 10 mM Tris-HCl (pH 7.6) was 1:1 (DNA stock, 0.1 mg/mL; NP stocks, 0.2 mg/mL).

Effect of NPs on the Hydrodynamic Size of Phage Lambda DNA. As shown in Figure 2A, the hydrodynamic radius of bacteriophage lambda DNA, here used as a model DNA molecule, was 43.9 nm at room temperature. When DNA (0.1 mg/mL) and c-AgNPs (0.2 mg/mL, initial hydrodynamic radius 10.9 nm; Figure 2B) were mixed at an equal volume ratio, the mixture assumed a single peak at 14.8 nm (Figure 2C). As there was no peak for the c-AgNPs-DNA mixture at 43.9 nm (hydrodynamic radius of DNA), we infer that no DNA-NP complexes were formed, and the DLS method detected only c-AgNPs, which scattered much more efficiently than did DNA. The slightly increased hydrodynamic radius of the c-AgNP-DNA mixture than that of c-AgNPs could be due to the hydrogen bonding between the DNA and the citrate coating of the c-AgNPs. 1027

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aromatic moieties of the GO nanosheets and the gradually opening DNA nucleobases at elevated temperatures. In addition, as double-stranded lambda DNA started melting at increasing temperatures, hydrogen bonds could be established between the opening DNA bases with the citrate, bPEI, and the hydroxyl surface groups of the c-AgNPs, b-AgNPs, and GO nanosheets, respectively. The hydrogen bonding with lambda DNA could be more pronounced for the two types of AgNPs than the GO nanosheets due to their differences in the density of surface coating (for the two types of AgNPs) or defects (for GO), one factor which could have contributed to the slower melting of DNA in the presence of the GO nanosheets. In principle, two opposing effects could be induced by hydrogen bonding: acceleration of DNA melting as more bases became available for hydrogen bonding with the nanostructure surfaces at elevated temperatures, and stabilization of DNA against melting through the formation of DNA−nanostructure complexes; the results obtained with AgNPs in Figure 3A and B suggest the net effect of the hydrogen bonding favored accelerated DNA melting. The less steep and more rugged slopes of the melting curves in the presence of the three types of nanostructures, as compared to that of the DNA control (Figure 3), suggest that the observed effects of the NPs on DNA melting could have involved multiple factors as those aforementioned, which competed in the melting process in a nonspecific and dynamic nature. Effects of NPs on Cellular Viability. Prior to the genotoxicity study, c-AgNPs, b-AgNPs, and GO nanosheets were analyzed for their general cytotoxicity both with human lymphocyte cell lines as well as with human primary lymphocytes using the resazurin assay of cellular metabolic state. The measured half-maximal inhibitory concentrations of AgNPs (2.9−15.5 μg/mL, Table 1) were similar to those concentrations obtained for AgNPs in earlier studies (based on 25 reports, the IC50 values of AgNPs for various cell lines varied between ∼0.6 and ∼120 μg/mL29). Among the three types of NPs, cationic b-AgNPs exhibited the highest toxicity (Table 1). High toxicity of cationic AgNPs has been reported by several studies, and its cause has been attributed to the adhesion of AgNPs onto the negatively charged cell membranes, their subsequent cell entry, or their released silver ions into the cells.30 The strong attachment of b-AgNPs onto the cellular surface was also observed in the current study. Figure 4 displays bright field images of NP-exposed lymphocyte cells where the accumulation of b-AgNPs in the surroundings of Tlymphocytes (indicated by red arrows) is evident. Adhesion of b-AgNPs onto lymphocyte cells was clearly visible down to the concentration of 6.25 μg/mL. In contrast, no clear cellular attachment was observed for either c-AgNPs or GO. When dealing with metal-containing NPs, dissolution is an important parameter determining their cellular effects. Also, different types of AgNPs may possess a different dissolution capability.

Figure 3. Effect of NPs on the melting temperature of lambda DNA. DNA melting was detected in relative fluorescence units. The DNA control in melting curves is shown in black (melting taking place at 94 °C). NPs were added to DNA in ratios indicated in the Figure (0.1 mg/mL DNA stock: 0.2 mg/mL NP stocks in 10 mM Tris-HCl). NP controls all induced negligible fluorescence as shown by the red (cAgNPs, A), blue (b-AgNPs, B), and gray (GO nanosheets, C) baselines, respectively. Three replicates are grouped for each NP:DNA ratio.

electrostatic attractions between the cationic b-AgNPs and the phosphate backbone of the DNA. In the case of GO nanosheets, π-stacking could be established between the

Table 1. Half-Maximal Inhibitory Concentration (IC50) Values with 95% Confidence Intervals, of the Tested NPs toward Human Lymphocytes in the Resazurin Assay c-AgNPs T-lymphoblastoma cells JURKAT B-lymphoblastoma cells WIL-2NS primary lymphocytes

b-AgNPs

GO

IC50 (μg/mL)

95% CI

IC50 (μg/mL)

95% CI

IC50 (μg/mL)

95% CI

12.2 13.4 12.5

6.3−23.4 8.64−20.9 9.5−15.2

2.9 3.2 5.25

1.9−3.7 2.4−4.1 2.7−9.3

341.7 401.9 176.7

251.0−493.8 331.1−533.5 120.2−257.5

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Figure 4. Cytokinesis block micronucleus assay results with the human T-lymphocyte cell line. Examples of NP exposed T-lymphocytes (JURKAT cell line); cells exposed to 100 μM H2O2 and nonexposed cells are shown for comparison. The tested compounds and their concentrations are indicated on each graph. Both bright field images (to visualize cellular morphology and cell-attached NPs) as well as fluorescence images (to visualize Hoechst-stained nuclei) are shown. The NPs attached to the cells or appearing in the vicinity of the cells are indicated with red arrows. Note that the cells were washed six times before imaging and thus, only NPs that were associated with cells were visualized. b-AgNPs between concentrations 25 and 6.25 μg/mL attached firmly to the cellular surface, while c-AgNPs were observed in the vicinity of cells only at the highest concentration used. Some GO nanosheets were found surrounding cells, most likely due to accidential coprecipitation of the cells and the micron-sized GO nanosheets. Apoptotic cells are identified due to their high staining intensity and fragmentated nuclear material, and necrotic cells can be identified due to their decomposed nuclei, while micronuclei can be seen as small pieces of nuclear material in the binucleated cells.

In our study, the dissolution rate of c-AgNPs and b-AgNPs in cell culture medium was determined at 7.6% and 13.9%, respectively. When we used this dissolution rate to adjust the half-maximal inhibitory concentrations of the AgNPs (Table S2, Supporting Information), the IC50 values of c-AgNPs were close to that of AgNO3, indicating that the toxicity of this type of NPs was primarily due to Ag ion dissolution. In contrast, the IC50 values of b-AgNPs were significantly lower than that of AgNO3, suggesting additional factors, such as NP cell

membrane attachment and subsequent internalization, intracellular transport, storage, and discharge, also played a significant role in the toxicity of b-AgNPs. In contrast to AgNPs, GO nanosheets did not cause any significant toxic effects up to 100 μg/mL (cellular viabilities are presented in Figures 5−7), with the half-maximal inhibitory concentrations ranging between 176 and 400 μg GO/mL (Table 1). The low toxicity of GO sheets is not surprising, as other recent reports21,31 also showed that 3.8 μm-wide GO 1029

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Figure 5. Effects of c-AgNPs, b-AgNPs, and GO sheets on cellular viability and genetic damage in the human T-lymphocyte cell line (JURKAT). Viability (left side of the Figure) was detected using the resazurin assay. For genotoxicity, the in vitro cytokinesis-block micronucleus assay was performed (right side of the Figure), and the number of micronuclei (MN) per 500 binucleated cells was calculated. At high concentrations of bAgNPs, no micronuclei were counted due to the toxicity. The solid horizontal line shows the number of MN in the nonexposed control, and the dashed line indicates the number of MN in the presence of 100 μM H2O2. At high concentrations, the number of binucleated cells was very low, and thus, MN could not be calculated (B,D). * indicates significant (p < 0.05) difference from the nonexposed control (concentration = 0).

sheets exhibited cytotoxicity only at 100 μg/mL. The cytotoxic effects of GO sheets occurring at these relatively high concentrations could be attributed to the mechanical interactions between cellular membranes and GO surface31 as well as the induction of reactive oxygen species on GO surfaces.32,33 Genotoxic Effects of NPs in the Cytokinesis-Block Micronucleus Assay. The genotoxic effects of the three types

of NPs were analyzed using the cytokinesis-block micronucleus assay (CBMN), which visualizes chemical-exposure induced formation of micronuclei which arise either from chromosome fragmentation or malsegregation of chromosomes during mitosis.24,34 The cytokinesis-block is used to restrict scoring of micronuclei in once-divided cells which are recognized by their binucleated appearance and in which micronuclei can be expressed. Examples of the CBMN assay results with T1030

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Figure 6. Effects of c-AgNPs, b-AgNPs, and GO sheets on cellular viability and genetic damage in the human B-lymphocyte cell line (WIL2-NS). Viability (left side of the Figure) was detected using the resazurin assay. For genotoxicity, the in vitro cytokinesis-block micronucleus assay was performed (right side of the Figure), and the number of micronuclei (MN) per 500 binucleated cells was calculated. At high concentrations of bAgNPs, no micronuclei were counted due to the toxicity. The solid horizontal line shows the number of MN in the nonexposed control and the dashed line indicates the number of MN in the presence of 100 μM H2O2. At high concentrations, the number of binucleated cells was very low, and thus, MN could not be calculated (D). * indicates the significant (p < 0.05) difference from the nonexposed control (concentration = 0).

lymphocytes are shown in Figure 4 and that with primary lymphocytes are shown in Figure S3 (Supporting Information), accordingly. Consistently with the discussion above, efficient colocalization of b-AgNPs with lymphocytes was observed, while no such apparent cell adhesion was found for c-AgNPs and GO nanosheets. Visualization of cellular nuclei with Hoechst 33342 staining or contrasting with Hemacolor stains after the CMBN assay is expected to show binucleated cells, if the cells have completed one division cycle. Nondivided cells show one nucleus, while cells in which two or more nuclear

divisions have occurred are expected to contain four or more nuclei.34 Indeed, the lymphocytes which had been incubated in chemical-free growth medium contained mostly two nuclei (Figures 4 and S3, Supporting Information). On the basis of the number of nuclei in cells, the nuclear division index (NDI) was calculated to quantify the efficacy of cell division, where NDI = 2 indicates that all cells have undergone one division, while NDI = 1 indicates no cell division. As expected, the toxicity and cellular division efficacy (NDI) of the lymphocytes were highly correlated (Figure S4, Supporting Information), so 1031

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Figure 7. Effects of c-AgNPs, b-AgNPs, and GO sheets on cellular viability and genetic damage in human primary lymphocytes. Viability of the primary lymphocytes (left side of the figure) was detected using the resazurin assay. For genotoxicity, the in vitro cytokinesis-block micronucleus assay was performed (right side of the Figure), and the number of micronuclei (MN) per 500 binucleated cells was calculated. At high concentrations of b-AgNPs and GO sheets, no micronuclei were counted due to the toxicity. The solid horizontal line shows the number of MN in the nonexposed control, and the dashed line indicates the number of MN in the presence of 100 μM H2O2. At high concentrations, the number of binucleated cells was very low, and thus, MN could not be calculated (D,F). * indicates the significant (p < 0.05) difference from th enonexposed control (concentration = 0).

H2O2 were counted, the number of micronuclei per 500 binucleated cells was 15.0 in the case of T-lymphocyte cell line, 13.2 in the case of B-lymphocyte cell line and 10.3 in the case of blood lymphocytes. Similar to H2O2, 500 binucleated cells of NP-exposed T- and B-lymphocytes and primary lymphocyte cells were counted, and their formed micronuclei were recorded. At high and toxic NPs concentrations, we were unable to score 500 cells as the overall numbers of these cells in the samples were exceedingly low. As seen from Figures 5−7, all three types of the tested NPs increased the number of

that at higher, toxic NPs concentrations the NDI values approached 1 indicating no nuclear division and maximal cytostatic effects among viable cells. Also, with increasing NPs concentrations, the number of necrotic and apoptotic cells increased (Figure S5, Supporting Information). Next, the induction of micronuclei by c-AgNPs, b-AgNPs, and GO sheets was evaluated. Examples of micronucleated cells induced by H2O2 treatment, which was used as a control chemical for genotoxicity, can be seen in Figures 4 and S3 (Supporting Information). When 500 cells exposed to 100 μM 1032

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pathway disrupted the normal segregation process of chromosomes in mammalian cells in in vitro conditions.

micronuclei compared to that in the nonexposed control, consistent with the increased cytotoxicity of the NPs. While several studies are available that report on the formation of micronuclei in response to NP exposure,35 there is no clear understanding on the exact mechanisms responsible. It is likely that NPs induce micronuclei due to the presence of acentric chromatide or chromosome fragments that are formed, if the level of DNA damage exceeds the cellular repair capacity and unrepaired double-stranded DNA breaks are retained.24 Indeed, several studies have shown extensive DNA damage,36 formation of DNA strand breaks37 as well as upregulation of cell cycle regulator protein p53 and DNA repair enzyme Rad5138 in AgNPs exposed cells. This suggests that AgNPs generate a remarkable level of cellular DNA damage, either as a result of NP-induced reactive oxygen species,17,39 released Ag ions, or direct attachment of AgNPs to DNA.40 The results in Figures 5−7 show clearly that the number of micronuclei in response to all three types of NPs increased at concentrations where significant decrease in cellular viability was observed. This is in accordance with the reports by Vecchio et al.,18 Li et al.,41 and AshaRani et al.,17 where relatively high (close to cytotoxic) concentrations of AgNPs caused micronuclei in various human cell lines. Only one study is available where AgNPs induced micronuclei at remarkably lower concentrations than the cytotoxic level of those NPs. Hackenberg et al.42 reported that 45 nm sized uncoated AgNPs were responsible for the formation of chromosomal aberrations (that may further induce micronuclei) in human mesenchymal stem cells at concentrations remarkably (10times) lower than those that were cytotoxic. The induction of micronuclei at concentrations lower than the cytotoxic level was observed in our case of b-AgNPs. While significant decrease in the viability of T- and B-lymphocytes was seen for 0.6 and 1.5 μg b-AgNPs/mL, elevated levels of micronuclei already occurred for 0.3 μg b-AgNPs/mL (Figures 5−7). On the basis of the literature, we propose that the reason for micronuclei was due to the high affinity of the positively charged b-AgNPs for biomembranes and biomolecules, induction of ROS, and/or release of Ag ions. In order to account for the effect of dissolved Ag ions on the observed genotoxic effect, we first calculated the concentration of Ag ions that were present at micronuclei-inducing concentrations of bAgNPs. The calculation was based on AgNPs dissolution in cell culture medium and showed that, e.g., at 0.4 μg b-AgNPs NPs/ mL that induced micronuclei formation in primary lymphocytes (Figure 7), 0.06 μg Ag ions/mL, was present. We then performed the CBMN test also with AgNO3, and apparently, increased number of micronuclei in primary lymphocytes was observed only starting from 1.7 μg Ag/mL (Figure S6, Supporting Information) which was significantly higher than the calculated concentration of Ag ions released from b-AgNPs. In contrast, when a similar calculation was done for c-Ag NPs, it showed that at 12.5 μg NPs/mL (concentration which induced micronuclei in primary lymphocytes; Figure 7), approximately 1 μg Ag ions/mL were released, and this concentration was close to the micronuclei-inducing concentration of AgNO3 (1.7 μg Ag/mL). This suggests that differently from the c-AgNPs case where the formation of micronuclei was likely dictated by released Ag ions, other factors were involved for b-AgNPs. We propose that b-AgNPs, due to their high affinity for the cellular surface and DNA, were efficiently internalized by the cells and via dissolution, physical binding to the DNA, or another

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CONCLUSIONS NP-biomolecular interactions play an important role in eliciting the biological responses to nanomaterial exposure. In this study, we have revealed a connection between NP−DNA interactions and the effects of NPs on DNA damage in vitro. It has been shown by our DSF assay that the introduction of NPs triggered an earlier onset of DNA melting, where cationic b-AgNPs elicited a significantly stronger effect than anionic c-AgNPs and planar GO nanosheets. We attribute this phenomenon to the strong electrostatic attraction between the phosphate backbone of DNA and the bPEI surface coating of b-AgNPs, as corroborated by our DLS experiment. When introduced to T- and B-lymphocytes and primary lymphocyte cells, b-AgNPs consistently displayed the most pronounced membrane adsorption as well as the most significant cytotoxicity among the three types of nanostructures. We assign the significant cytotoxicity of b-AgNPs to their high affinity for cellular membranes and DNA and consider Ag ion release in cell culture medium as the main cause of the toxic effects of c-AgNPs. GO sheets, in comparison, exhibited relatively low toxicity compared to the two types of AgNPs, and such cytotoxicity became apparent on T- and B-lymphocyte cell lines at 50−100 μg/mL resulting from the physical interactions between the cells and the GO sheets. All three types of NPs induced the formation of micronuclei in human lymphocytes. We postulate that in the case of c-AgNPs, the micronuclei were formed due to the release of toxic Ag ions into the cell culture medium, while cationic b-AgNPs, due to their high affinity for cell surfaces, were taken up by the cells to induce genotoxic effects via NP-DNA binding, intracellular release of Ag ions, or other mechanisms to interfere with normal DNA synthesis during the cell cycle. Increase in micronuclei in the case of GO sheets may have been partly due to the physical interactions between the nanostructures and cell membranes or cytoskeletal machinery within the cells, which may have disrupted the chromosome segregation process during mitosis leading to lagging or displaced chromosomes from which micronuclei can originate. In addition to contributing to the field study of safe nanotechnology, these findings may serve as a basis for the design and application of nanobiotechnology and nanomedicine.

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ASSOCIATED CONTENT

* Supporting Information S

XPS data of GO nanosheets, DSF measurement of the effect of Ag ion on DNA melting, Ag ion dissolution-corrected IC50 values of cAgNPs and bAgNPs, and the cytotoxicity and genotoxicity of cAgNPs, bAgNPs, and GO sheets. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Authors

*(A.I.) E-mail: [email protected]. Tel: 61-8-83023563. *(P.-C.K.) E-mail: [email protected]. Tel: 61-3-99039276. Funding

The South Australian Collaboration Pathway Program grant (International Research Cluster for Nanosafety) is acknowledged for providing funding for this work. P.-C.K. acknowl1033

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edges a DVS fellowship from the Office of Chief Executive at CSIRO. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank the Biomedical Program within the CSIRO Manufacturing Flagship and the TEM facility at the University of South Australia for providing the infrastructure needed to complete the experiments. We also thank Thomas Gengenbach at CSIRO for assistance with the XPS measurement, and Claudia Hager and Gianluca Brunetti for experimental assistance.

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ABBREVIATIONS AgNP(s), silver nanoparticle(s); b-AgNPs, bPEI (branched ethylene imine) coated silver nanoparticles; c-AgNPs, citrate coated silver nanoparticles; CBMN, cytokinesis-block micronucleus assay; DSF, differential scanning fluorimetry; DLS, dynamic light scattering; GO, graphene oxide; IC50, halfinhibitory concentration; NDI, nuclear division index; NP(s), nanoparticle(s); TEM, transmission electron microscopy; XPS, X-ray photoelectron spectroscopy

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DOI: 10.1021/acs.chemrestox.5b00052 Chem. Res. Toxicol. 2015, 28, 1023−1035