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Feb 18, 2013 - Biosensing Approaches for Rapid Genotoxicity and. Cytotoxicity Assays upon Nanomaterial Exposure. Xuena Zhu , Evangelia Hondroulis ...
Biosensors

Biosensing Approaches for Rapid Genotoxicity and Cytotoxicity Assays upon Nanomaterial Exposure Xuena Zhu, Evangelia Hondroulis, Wenjun Liu, and Chen-zhong Li*

The increased utilization of nanomaterials could affect human health and the environment due to increased exposure. Several mechanisms regarding the negative effects of nanomaterials have been proposed, one of the most discussed being oxidative stress. Many studies have shown that some metal oxide nanoparticles can enhance reactive oxygen species generation, inducing oxidative stress, DNA damage, and unregulated cell signaling, and eventually leading to changes in cell motility, apoptosis, and even carcinogenesis. 8-Hydroxy-2′-deoxyguanosine (8-OHdG) is one of the predominant forms of oxidative DNA damage, and has therefore been widely used as a biomarker for oxidative stress and carcinogenesis. Ther are two major objectives to this study. Firstly, the development of a novel lateral flow immunoassay (LFIA) is presented to measure the concentration of 8-OHdG in cells and thus reveal the nanotoxicity on the genomic level. The feasibility of this new method is validated by comparison with two other established methods: Alamar Blue assay and a recently developed electrical impedance sensing (EIS) system on the level of cell proliferation/ viability. Secondly, the toxicological effects of three metallic nanoparticles (CuO, CdO, and TiO2) are investigated and compared using these three methods with completely different mechanisms. The results show that there is a high variation among different nanoparticles concerning their ability to cause toxic effects. CuO nanoparticles are the most potent regarding cytotoxicity and DNA damage. CdO shows a fallen cell viability as well as DNA damage, however, to a lesser extent than CuO nanoparticles. TiO2 particles only cause very limited cytotoxicity, and there is no obvious increase in 8-OHdG levels. In conclusion, LFIA as well as the EIS system are useful methods for quantitative or qualitative nanotoxicity assessments with high sensitivity, specificity, speed of performance, and simplicity.

X. Zhu, E. Hondroulis, Prof. C.-z. Li Nanobioengineering/Bioelectronics Lab Department of Biomedical Engineering Florida International University 10555 West Flagler Street, Miami, FL 33174, USA E-mail: [email protected] Tel: (305) 348 0120 Fax: (305) 348 6954 W. Liu Department of Biochemistry and Molecular Biology University of Miami 1011 NW 15st, Miami, FL 33136, USA DOI: 10.1002/smll.201201593 small 2013, 9, No. 9–10, 1821–1830

1. Introduction Significant progress in nanotechnology has promoted the development of industrial technology greatly in recent years. While on the other hand, the increased utilization of nanomaterials could affect human health and the environment due to increased exposure.[1–3] For instance, metallic nanoparticles (NPs) are used increasingly in medicine and industry, as well as in various consumer products including personal care products, plastic paints, textiles, sunscreens, cosmetics, and food products.[4] Consequently, occupational or nonoccupational exposure to metal NPs is growing. The current

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knowledge is limited to the potential health effects caused by nanomaterials; however, it shows that they may cause adverse effects at the routes of exposure such as skin, gastrointestinal tract, and lung.[1,4,5] Furthermore, some nanomaterials made of certain metals may have genotoxic or carcinogenic effects. One of the mostly discussed mechanisms behind the health effects induced by some metal oxide NPs is their ability to enhance reactive oxygen species (ROS) generation causing oxidative stress, DNA damage,[6–15] and unregulated cell signaling, which eventually leads to change in cell motility, apoptosis, carcinogenesis etc.[16–18] Therefore, characterization and reliable toxicity screening tools are required for new and existing nanomaterials, ensuring their compatibility for medical applications and for the safety of the environment. There are a number of different approaches that can be taken to assess the toxic effects of NPs, which can be categorized into three classes based on their objects of measurement. 1) Cell proliferation/viability assay, which is also called cytotoxicity measurement in other publications, includes Trypan Blue assay,[8,19–22] Alamar Blue assay,[8,20,23] Neutral Red assay,[24,25] lactate dehydrogenase (LDH) assay,[8,19,20,26–29] Formazan-based assays (MTT, MTS, WST),[8,19,20,24–26,28,29] and clonogenic assay.[8,19,20] 2) Direct or indirect intracellular ROS measurement, such as lipid peroxidation measurement,[8,19,20,23,30] glutathione (GSH) assay,[8,20,23–25,28,31] electroparamagnetic resonance (EPR) assay,[20,21] and 2,7-dichlorofluorescin (DCFH) assay.[8,20–25,28,31] 3) Assays on the genomic level, like Comet assay,[8,19–22,25,27,29,32] and DNA damage biomarker assay.[19,21,33] It is well-known that excessive generation of ROS can oxidize cellular biomolecules.[7] Free radicals also lead to oxidative modifications in DNA, including strand breaks and base oxidation. Among oxidative DNA damage products, 8-hydroxyguanine and its nucleoside 8-hydroxy-2′deoxyguanosine (8-OHdG) are probably the most studied due to their relative ease of measurement and pre-mutagenic potential.[6,7,35,36] Evidence shows that 8-OHdG can give rise to G-to-T transversion mutations in key genes known to be involved in the development of cancer,[37] so elevated 8-OHdG levels have been noted in numerous tumors[7,35] and thus is widely used as a biomarker for oxidative stress and carcinogenesis.[38] Until now, the traditional methods for 8-OHdG quantitative analysis are high-performance liquid chromatography (HPLC) with electrochemical detection (ECD),[39,40] gas chromatography-mass spectrometry (GCMS),[41] HPLC tandem mass spectrometry,[42,43] and Enzymelinked Immunosorbent Assay (ELISA).[44] However, most of the approaches mentioned above are time-consuming, expensive, and require special techniques and equipments. Therefore, there is an urgent need to develop simple, scalable, inexpensive, and high throughput analytical tools capable of mass screening in nanotoxicological investigations. Herein, in this paper, we present two novel analytical methods developed by our lab to investigate and compare three well-studied particles (CuO, TiO2, CdO), regarding their overall cytotoxicity and ability to cause DNA damage on mouse epithelial cells (CCL-149). The toxicological effects were measured by an electrical impedance sensing (EIS) system, whose mechanism has been described by one of our

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previous studies.[34] The system measures the resistance produced by growing cell monolayers over electrodes and can detect changes in resistance that may occur with changes in the cell layer after NPs exposure, which is used for proliferation/viability assessment. Meanwhile, we demonstrate a novel lateral flow immunoassay (LFIA) to measure the concentration of DNA damage biomarker (8-OHdG) and thus reveal the nanotoxicity on the genomic level. LFIA, also known as Immunochromatographic test strip, have been widely used as in-field and point-of-care diagnostic tools for testing cancer biomarkers,[45–50] proteins,[51] drugs,[52–54] hormones,[55] and metabolites in biomedical,[51,56–58] food and environmental settings. The results show that different NPs with the similar size and concentration present different toxic effects. EIS system and LFIA strip provide useful methods for nanotoxicity assessment with high sensitivity, simplicity and speed of performance.

2. Results and Discussion 2.1. Characterization of NPs The primary sizes of the NPs were determined using Transmission electron microscopy (TEM). As shown in Figure 1, the size of nano-TiO2 was about 20 ± 5 nm (panel a). The nano-CuO had the size of about 50 nm (panel b). The size was consistent with the data specified by the manufacturer. Furthermore, dynamic light scattering (DLS) was used to characterize the behavior of the NPs, and the hydrodynamic sizes and surface charges were measured using a Zetasizer in a biological environment (i.e., DMEM cell culture medium). The NPs were suspended in deionized water or DMEM for analysis of their agglomerate sizes and zeta potential. The size and surface charge characteristics of the NPs are summarized in Table 1. Zeta potential measurements revealed that TiO2-NP had the surface charge ranging from −10.5 mV (in medium) to -15 mV (in water), and the average particle hydrodynamic diameter ranged from 574 nm (in medium) to 1510 nm (in water). CuO-NPs demonstrated mean agglomerate sizes ranging from 338 nm (in medium) to 1350 nm (in water), and the surface charge ranged from −12.5 mV (in medium) to −6 mV (in water). These differences in zeta potential of TiO2 and CuO-NPs can influence the particle uptake as well as toxicological parameters like ROS generation and genomic damage. Particle size observed by DLS did not coincide with the results obtained from the TEM. The larger size of the NPs in the hydrodynamic state compared to the size obtained by the TEM might be due to the tendency of the NPs to aggregate in aqueous state. This finding is supported by other investigators.[28,59,60] The three metal oxides used in this study are relevant NPs types and are in widespread use in a number of consumer products. CuO NPs are utilized in various applications such as industrial catalysts, semiconductor devices, antimicrobial preparations, heat transfer fluids, and cosmetics.[23,61–63] TiO2 NPs have excellent optical performance and electrical properties and are produced for applications in paints and coatings and also in cosmetics as UV-absorbers. CdO can be used

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and DNA damage. They are reasonably been choosen here as models for nanotoxicity assement. 2.2. Cytotoxicity Study of NPs by Alamar Blue Assay Although nanotoxicity of the three particles we choose have been studied individually or at certain combination in other’s publication, we need firstly to characterize their toxic effects simultaneously in our mouse epithelial cell model by a well established method, which can then serve as the reference for our newly developed detection methods. Thus, Alamar Blue assay in a time course manner was carried out with or without the presence of NPs. The result was shown in Figure 2. At time point 0 h while NPs were absent in the medium during the Alamar Blue incubation phase, four groups exhibited nearly no difference in cell metabolic activity. At the rage from 0 h to 30 h, control (no NPs) and TiO2 showed normal cell growth and increase in metabolic activity, which is consistent with many other studies stating TiO2 with a similar size as what we used has very limited cytotoxicity. Interestingly, TiO2 may have a positive effect in promoting cell growth & metabolism through an unclear mechanism. On the other hand, the other NPs CuO and CdO not only prevent the normal cell growth, but also cause severe cell viability issue. Even shorly at the point 6 hours after adding NPs, the metabolic activity falls by 40% around in both setting. Notably, every cell with metabolism including unhealthy and flow cells contribute to convertion of the redox indicator of Alamar Blue reagent, thus this assay has a lower cut off when compared to other cell proliferation/viability assay.

2.3. Cytotoxicity Study of NPs by Electrical Impedance Sensing

Figure 1. Size of NPs observed by TEM. (a) Left: the size of TiO2-NPs was about 20 ± 5 nm and bar scale is 100 nm and (b) Right: the size of CuO-NPs was about 50 nm and bar scale is 100 nm.

in batteries, electroplating baths, pigments, plastics, catalyst, ceramic glazes, synthetic products, and a variety of other materials.[64] Nevertheless, many studies[21–25,28–30,65] have demonstrated that these metal oxide NPs can induce cytotoxicity Table 1. Physico-chemical characteristics of copper oxide and titanium dioxide nanoparticles. Nanoparticles

DLS Size [nm] Water

DMEM

CuO

1350

TiO2

1510

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Zeta Potential [mV] Water

DMEM

338

−6

−12.5

574

−15

−10.5

The whole cell-based electrical impedance sensing (EIS) system, previously published by our lab,[34] is also applied to determine the cytotoxiciy of CuO, CdO, and TiO2 NPs on mouse epithelial (CCL-149) cells. The EIS system measures the resistance produced by growing cell monolayers over electrodes and can detect changes in resistance that may occur with cell density changes in the cell layer after NPs exposure, and thus provides a kinetic monitor of cell viability. In this section, three types of NPs (100 ug mL−1) were added to the wells after 24 h of cell attachment, and the resistance changes produced by the attachment of cells to the electrodes were monitored over a 40 h time period. This setup would provide the kinetics of the interactions of the NPs with the cells. Figure 3 illustrates the resistance readings for five different settings: Blank (medium only), CuO-24 hrs (CuO, 50 nm, 100 μg mL−1, added after 24 h of cell attachment), CdO-24 hrs (CdO, 100 μg mL−1, added after 24 h of cell attachment), TiO2-24 h (TiO2, 20 ± 5 nm, 100 μg mL−1, added after 24 h of cell attachment) and a control of cells only (mouse epithelial cells, CCL-149). As shown in Figure 3, resistances in all five groups increased within the first 24 h with more and more cells attaching to the surface of electrodes before adding NPs. Nevertheless, upon inoculation of the NPs after 24 h, the trends became extremely different. Once the CuO was added to the cells (Line B), we observed a rapid decrease

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No NPs (Line A) CdO (Line C)

CuO (Line B) TiO2 (Line D)

2.00

1.50

1.00

0.50

0.00 0

6

12

30

Time points when measurements are taken/hour

Figure 2. Alamar Blue assay for CuO, CdO, and TiO2 on CCL-149. Lines A represent the Cells only (cells with medium only). CuO (line B), CdO (line C), and TiO2 (line D) are added at the initial time point.

Cells only (Line A) CuO-24 hrs (Line B) CdO-24 hrs (Line C) TiO2-24 hrsl (Line D)

6000

Resistance [ohms]

5000

4000

3000

2000

1000 0

5

10

15

20 Time [hrs]

25

30

35

40

Figure 3. Resistance readings for CuO, CdO and TiO2 on CCL-149. Lines A and E represent the cells only (cells with medium only) and Blank (medium only) resistance readings. CuO (line B), CdO (line C), and TiO2 (line D) are added after 24 h of cell attachment.

in the resistance values measured, eventually returning to readings similar to the blank (Line E), indicating cell detachment and the harsh cytotoxic effect of CuO towards the CCL-149, and being consistant with the results onbtained by Alamar Blue assay. The cytotoxic measurements of TiO2 NPs towards the CCL-149 (Line D) showed slight difference in resistance values to those observed for the control (Line A). From this observation, it is evident that TiO2 NPs (25 nm) have fewer cytotoxic effects on the cells compared to the CuO NPs (50 nm) with the same concentration (100 ug mL−1) and similar size, which is also comparable with the results got from Alamar Blue assay. When cells were exposed to CdO, noticeable changes in resistance (Line C) were observed compared to the control (Line A), which is consistent with the fact that it is extremely toxic and affects the growth mechanism of the cells given by Alamar Blue assay. CdO was used here as a negative control and employed to demonstrate the ability of the EIS system. Cadmium is a toxic material that has been shown to cause lysosomal damage and DNA breakage in mammalian cells and disrupt mitochondrial function and promote apoptosis.[65] However, the value decreased slowly compared to the CuO NPs (Line B), indicating a slower detachment rate of the cells. The observed phenomena may be attributed to the fact that the smaller NPs (CuO, 50 nm) can enter the cell and damage the cells more easily.

2.4. Genotoxicity Assessment by Oxidative DNA Damage Biomarker 2.4.1. Principle of the Genotoxicity Assessment by Lateral Flow Strip

Scheme 1. Mechanism of competitive lateral flow immunoassay for 8-OHdG testing.

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The principle of the immuno strip based on the specific immunoreactions occurring between the antibodies and the DNA damage biomarker (8-OHdG) has been illustrated in Scheme 1. Generally, there are two types of formats for strip testing, named non-competitive and competitive. The competitive format is used when testing small molecules with single antigenic determinants, which cannot bind to two antibodies simultaneously. If this format is chosen, the analytes in the sample will compete with the antigen immobilized on the test line for the antibody from

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conjugation pad. So the color intensity of the test line is inversely proportional to the analyte concentration in the sample. The test strip consists of five components, each of which plays a distinct role in biomarker detection. The first part is a sample-loading zone where the sample to be analyzed is applied. The sample then moves along the strip due to capillary action and finally gets collected by the last part known as the absorption pad. Figure 4. The photographs of test strips of LFIA based on standard samples (8-OHdG was When the sample moves into the second dissolved in 1X PBS). Top line: Test-line; Bottom line: Control-line. The concentration unit for part which is called the conjugate pad, those shown on the bottom is nanogram/milliliter. the biomarkers react with the gold nanoparticles (AuNPs) labeled monoclonal antibodies (AuNPs-Abs) to form a complex. At the test line, which is the third part, BSA-8-hydroxyguanosine conjugates capture the unbound AuNPs-Abs conjugates. The accumulation of AuNPs in the test line induces a red color which is visible with the naked eye. This color change accomplishes the detection of biomarker in a sample and the color intensity is inversely proportional to the biomarker concentration. After the AuNP-Abs conjugates are captured at the test line, the unbound constituents, including the tri-complex (AuNP-Absbiomarker) formed at the conjugate pad, excess AuNP–Abs, and the fluid fraction, continue to flow along the strip. The tricomplex and unbound AuNP–Abs get captured at the control line by the polyclonal Goat anti-Mouse IgG. This induces a red color at the control line which allows us to confirm the proper functioning of the immunostrip. 2.4.2. Quantitative Analysis of the Biomarker A series of 8-OHdG standard solutions with concentrations of 0, 0.1, 1, 10, 50, 100, 500, 1000, 10 000, and 100 000 ng mL−1 in 1X PBS buffer were prepared and applied to the strips. After 15 min, photographs were taken by using digital camera, then software ImageJ[66] was used for the quantitative analysis. Figure 4 shows the typical responses of the LFIA to 8-OHdG with increasing concentrations from 0 to 100 000 ng mL−1 dissolved in 1X PBS. The color intensity of the test line deceased when the sample concentration increased in general, which was consistent with the theory of detection of competitive format. The visual detection limit is defined herein as the minimum target analyte concentration required by the T-line for showing no obvious staining effect. Following this definition, the visual detection limit achieved by the standard sample is above 1000 ng mL−1. The visual detection range is from 0 to 10 000 ng mL−1. In order to quantitatively extract the detection limit and detection range of this LFIA method, the test strips were further subjected to optical density analysis. The signals from both the T-line averaged from three parallel runs, and one from red color tape (reference) were digitized to optical density using software of ImageJ and expressed by the integral area of the cross-section of the T-line (areaT) and reference-line (areaR) within a fixed peak width. In order to eliminate the influence of artificial effects, a relative optical small 2013, 9, No. 9–10, 1821–1830

Figure 5. Optical density profiles of the T-line and C-line recorded by using software ImageJ and Sigmaplot after running a series of standard solutions with different 8-OHdG concentrations dissolved in 1X PBS.

density (ROD) defined as areaT/areaR was used in the signal analysis. The optical density profiles of both the T-line and C-line recorded under different analyte concentrations are shown in Figure 5 with the optical densities of the T-line and C-line being normalized with respect to that of baseline. The optical intensity of the T-line quite obviously increased with the decrease of the analyte concentration. The discrimination of intensities is more intuitive compared to the photograph taken by the camera. Considering the actual situation, the meaningful research range for 8-OHdG is from 0 to 500 ng mL−1. To extract the detection limit of the current LFIAs, the ROD’ defined as ROD/ABD (average blank density) of the T-line is plotted against the concentration of 8-OHdG in the logarithm scale as shown in Figure 6. Linear fitting of the dose-response curves suggests that the linear response range of the standard sample is from 0.1 to 500 ng mL−1 with a correlation coefficient of 0.9937. With the definition of the detection limit as the minimum concentration of analyte required for inducing a 10% ROD’ decrease, it was determined as 0.9 ng mL−1 for standard sample.

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1.2

Normalized ROD'

1.0

Normalized ROD'

a

y=(-2.40±0.01)x+(0.80±0.02) R2=0.9937

0.8 0.6 0.4 0.2 0.0 -1.0

1.0

y = -0.4128x + 0.9909 R² = 0.957

0.8 0.6 0.4 0.2 0.0

0.0

1.0

2.0

0.0

3.0

log10[8-OHdG] [ng mL-1]

2.4.3. Effects of NPs on 8-OHdG level as a Biomarker of Oxidative DNA Damage After confirming the feasibility of this strip for 8-OHdG testing, it can then be used for the genotoxicty assessment in cells. A series of 8-OHdG solutions with concentrations of 0, 1, 10, 20, 40 and 80 ng mL−1, both in cell lysis buffer and cell culture medium (DMEM), were prepared and applied to the strips. The cell lysate (lysed cell by cell lysis buffer) and cell culture medium (supernant) after NPs (CuO, CdO and TiO2) exposure were also collected and applied to the strips. After 15 min, photographs were taken by using photo scanner (Figure 7), and then software ImageJ was used for the quantitative analysis as illustrated above. Figure 7(a) and (c) show that the optical intensity decreased obviously when the concentration of 8-OHdG increased, both in the cell lysis buffer and in the cell culture medium, which is still consistent with the mechanism of the competitive format and can be used as

1.0

1.5

2.0

-1

Log10 [8-OHdG] [ng mL ]

b Normalized ROD'

Figure 6. Dose-response curves for 8-OHdG based on optical density analysis using standard samples. Values are mean SD from three independent experiments.

0.5

1.0

y = -0.0058x + 0.9457 R² = 0.9632

0.8 0.6 0.4 0.2 0.0 0

20

40 60 [8-OHdG] [ng mL-1]

80

Figure 8. Calibration curves for 8-OHdG both in cell lysis buffer (a) and cell culture media (b). Values are mean SD from three independent experiments.

a calibration standard. Figure 7(b) and (d) demonstrate the results by using different NPs, in which the color intensities of all the three experimental groups (NPs) are slightly lighter compared to the control indicating that these metallic NPs can lead to more 8-OHdG generation, in other words, are genotoxic. Among the three experimental groups, CuO NPs seemed to be the most genotoxic with the highest 8-OHdG generation in both the cell lysate and the cell culture medium, which was consistant with the results given by both Alamar Blue assay and EIS system. Figure 8 illustrates the linear fitting curves for 8-OHdG, both in the cell lysis buffer (panel a) and the cell culture media (panel b) with correlation coefficients of 0.957 and 0.9632, respectively. 8-OHdG concentrations in different experimental groups can be estimated by these two curves presented in Table 2 and Table 3. The CuO groups, both in the cell lysate and the cell culture medium, have the highest 8-OHdG concentrations (23.24 and 13.78 ng mL−1, respectively), followed by the CdO groups (12.35 and 8.43 ng mL−1, respectively). While TiO2 shows no special genotoxicity with similar 8-OHdG concenFigure 7. (a) The photographs of test strips based on 8-OHdG samples dissolved in cell lysis trations compared to the control. By combuffer. (b) The photographs of test strips based on cell lysate after cells were treated with paring the two tables, we can see that the NPs solutions. (c) The photographs of test strips based on 8-OHdG samples dissolved in cell culture medium (DMEM). (d) The photographs of test strips based on culture medium DNA oxidative biomarker is prone to stay after cells were treated with NPs solutions. Top line: Test-line; Bottom line: Control-line. The inside the cells instead of being released out of the cells (i.e. for CuO, there is concentration unit for those shown in the middle is nanogram/milliliter.

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Table 2. Estimated 8-OHdG concentrations in cell lysate after cell were exposed to different NP solutions. Sample Typea)

ROD’b)

8-OHdG Concentration [ng mL−1]

Control

0.6535

2.17

CuO

0.4269

23.34

CdO

0.5402

12.35

TiO2

0.7449

1.76

a)

Different cell lysate samples after cells were exposed to different NPs solutions; b)ROD’:ROD

(relative optical density)/ABD (average blank density)

Table 3. Estimated 8-OHdG concentrations in cell culture media after cells were exposed to different NP solutions. Sample Typea)

ROD’b)

8-OHdG Concentration [ng mL−1]

Control

0.9308

2.56

CuO

0.8658

13.78

CdO

0.8968

8.43

TiO2

0.9291

2.86

a)Different

cell culture media samples after cells were exposed to different NPs solutions;

b)ROD’:ROD

(relative optical density)/ABD (average blank density)

and same concentration by using an EIS system. In addition, we successfully designed and presented a novel paper based immunoassay to measure the concentration of 8-OHdG in cells. This simple and rapid assay could provide a high throughput analysis capable in mass screening in nanotoxicological investigations. Combined together with Alamar Blue assay, these three methods provide a wide angle scope of nanotoxicity with readout from physical cell attachment, metabolic activity and genomic lesion. The data indicated that there was a high variation among different NPs regarding their ability to cause cytotoxicity and DNA oxidative lesions. CuO NPs were the most toxic, both in cell viability level and DNA damage level. CdO showed some to severe cytotoxic effects as well as increased 8-OHdG levels, however, less than those caused by CuO NPs. TiO2 particles caused very limited cytotoxicity, while, there was no obvious 8-OHdG increase. There is a correlation between DNA damage and cytotoxicity, however, the mechanism of how NPs with similar size convey different genotoxicity and cytoxicity on the molecular and whole cell level respectively is not well understood and could be a subject for further investigation. In conclusion, the EIS system and LFIA strip can provide useful methods for quantitative or qualitative nanotoxicity assessment with high sensitivity, specificity, speed of performance, and simplicity based on the comparison with the Alamar Blue assay.

23.24 ng mL−1 8-OHdG in cell lysate, while 13.78 ng mL−1 in cell culture medium). Genomic instability, under certain thresholds, could be partially fixed by the mechanism of DNA replication coupled repair and become tolerated by cells. However, cells undergoing proliferation, like germ cells and stem cells, may be redirected to cell cycle arrest and programmed cell death/ apoptosis given severe genomic damage. The mechanism of how NPs with similar size convey different genotoxicity and cytotoxicity on the molecular and whole cell level respectively is not well understood. However, in this study we do observe a correlation between DNA damage and cytotoxicity. Comparing the above sections, we notice that the high toxicity of CuO NPs leads to almost 100% cell death within 2 h after NPs addition (Figure 3, line B), which could be a response to the DNA damage (illustrated in Table 1 and 2, CuO NPs induce the highest 8-OhdG generation in cell) as a way to prevent a mutagenic outcome. In the meantime, it is also possible that DNA damage to some extent is a consequence of cytotoxicity. However, from the present data, it is hard to clarify which process is the driving force. On the other hand, the TiO2 NPs cause cytotoxic effects (Figure 3, line D), however, without apparent increase in 8-OHdG levels, indicating different mechanisms underlying these effects.

3. Conclusion In this work, we use two novel methods developed by our lab to assess the cytotoxicity and oxidative DNA damage in mouse epithelial cells (CCL-149) after exposure to three different metal oxides (CuO, TiO2, CdO) NPs. We established a resistance-time relationship for three NPs with similar size small 2013, 9, No. 9–10, 1821–1830

4. Experimental Section Materials: PBS (1X, PH = 7.4), Triton X-100 and trisodium citrate dihydrate were purchased from VWR (West Chester, PA). Gold chloride trihydrate (HAuCl4 •3H2O), bovine serum albumin (powder), sodium borohydride (NaBH4), sodium periodate (NaIO4), ethylene glycol, potassium carbonate (K2CO3), sodium phosphate (Na3PO4), sodium chloride (NaCl), copper oxide (CuO), cadmium oxide (CdO), and titanium dioxide (TiO2) were purchased from Sigma-Aldrich (St. Louis, MO). Tween 20 (polyoxyethylene-20-sorbitan monolaurate), sucrose, and tris–HCl (1 M) were obtained from Fisher Scientific (Fairlawn, NJ). Plastic backing, nitrocellulose membrane, absorbing pad, and cellulose paper were acquired from Millipore (Billerica, MA). 8-Hydroxy-2-deoxyguanosine and 8-hydroxyguanosine were purchased from Cayman chemical (Ann Arbor, MI). Mouse monoclonal antibodies to 8-hydroxyguanosine and polyclonal Goat anti Mouse IgG were purchased from Abcam (Cambridge, MA). Alamar Blue, DMEM, fetal bovine serum, and penicillin–streptomycin were obtained from Invitrogen (Merelbeke, Belgium). Mouse epithelial cells, CCL-149 were bought from ATCC (Manassas, VA). Equipment: HP Scanjet G3110 Photo Scanner was bought from Hewlett-Packard, Palo Alto, CA. The Zetasizer is from Malvern Instruments, Woodstock, GA. Drying Oven was purchased from VWR (West Chester, PA). Dispenser Linomat 5 was purchased from CAMAG (Wilmington, NC). ImageJ and SigmaPlot software were downloaded from the internet. Synthesis of Antibody-Conjugated Gold NPs (AuNPs–Abs): The AuNPs were synthesized by a modified citrate reduction method.[67] Briefly, HAuCl4 solution (50 mL, 0.01% in superpurified water) was heated to a boil, and then trisodium citrate solution (1 mL, 1%) was added rapidly under constant stirring. Gradually, the color changed from pale yellow to bright red. After the color change,

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the solution was boiled for another 10 min and stirred without heating for another 10 min to complete the reduction of the gold chloride. After the solution reached the room temperature, the size of the AuNPs were characterized by the Zetasizer and found to be ∼20 nm. The AuNPs solution was concentrated 5X and the pH of the AuNPs solution for antibody labeling was adjusted to pH 8.5∼9.0 with K2CO3 (0.1 M). The method for optimal antibody labeling concentration determination was followed from Y Zhao et al.[68] Purified anti-8-OHdG mAb (60 μL, 0.54 mg mL−1) was added to the AuNPs solution (750 μL, 5X) and stirred gently at room temperature for 1 h. The conjugate was stabilized by adding BSA (90 μL, 10%) in sodium borate (20 mM) for a final concentration of 1% and incubate for another 20 min. Then the mixture was centrifuged for 15 min at 7000 rcf. Two phases can be obtained: a clear to pink supernatant of unbound antibodies and a dark red, loosely packed sediment of the AuNPs-Abs conjugates. The supernatant was discarded and the pellet was resuspended in BSA/PBS (900 μL, 1%). Following the same centrifugation step, the supernatant was removed and the soft sediment of conjugates was resuspended in buffer (900 μL), containing sodium phosphate (20 mM), Tween 20 (0.25%), sucrose (10%), and BSA (5%) by the end. The conjugate was stored at 4 °C until required for use. Preparation of BSA-8 Hydroxyguanosine Conjugates (for Test Line Capture): 8-Hydroxyguanosine (5 mg) was dissolved in NaIO4 (1 mL, 50 mM) and the mixture was incubated for 1 h in the dark. The reaction was stopped by adding ethylene glycol (2.5 μL) for 5 min. Then the mixture was mixed with BSA (2 mL, 25 g L−1, pH = 9.5, adjusted by K2CO3 (50 g L−1)) under constant stirring dropwise and incubated for another 1 h. After that, NaBH4 (2 mL, 24 g L−1) was added and the mixture was incubated in the dark at 4 °C overnight (12–16 h). Finally, the conjugates were dialyzed against 1X PBS and stored at −20 °C. Assembly of the Lateral Flow Immuno Strip: BSA-8 hydroxyguanosine conjugates were used as the test line (T) capture reagent, while goat anti-mouse IgG (1 mg mL−1) was used as the control line (C) capture reagent. These capture reagents were dispensed by the Linomat 5 dispenser onto a nitrocellulose membrane as the test and control lines. The sample pad (5 mm × 19 mm) was treated with buffer Triton X-100 (containing 0.25%), Tris-HCl (0.05 M) and NaCl (0.15 mM), then AuNp-Abs conjugates (30 uL) was dispensed by pipette onto a glass fiber membrane which was called the conjugate pad (5 mm × 9 mm). After drying these membranes, the sample pad, conjugate pad, nitrocellulose membrane, and absorbent pad were pasted onto a plastic backing plate which was already cut into 5 mm-wide strips (Scheme 1) using a strip cutter. Then the strips were stored in a self-sealing plastic bag until use. Preparation of Standard Solution and Samples: Stock solutions of 8-OHdG (500 μg mL−1) were prepared before use by dissolving the 8-OHdG powder in purified PBS (1X) solution, DMEM and cell lysis buffer respectively. Working standards ((0.1–100000) ng mL−1, (1–80) ng mL−1 and (1–80) ng mL−1) were prepared further in the corresponding media before use and kept at 4 °C. Particle Source and Characterization: TiO2-NPs (ϕ < 25 nm) and CuO-NPs (ϕ < 50 m) were obtained in powder form from SigmaAldrich (Cat No. 637254 and 544868). The dry powder of NPs was suspended in deionized water (1 mg mL−1) and cell culture medium (1 mg mL−1) respectively, and then sonicated using a sonicator bath at room temperature for 20 min (120 V/50–60 HZ) to form a

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homogeneous suspension. For size measurement, sonicated NPs stock solutions (1 mg mL−1) were then diluted to working solutions (100 μg mL−1). TEM was used to characterize the size and shape of the NPs. A drop of aqueous NPs suspension was placed onto a carbon-coated copper grid, air-dried and observed with TEM (2000FX, JEOL). DLS was used to determine the hydrodynamic size and zeta potential of the NPs suspension both in the DI water and culture medium. Cell Culture and Exposure to NPs: Mouse epithelial cells (CCL149) were obtained from ATCC and cultured in DMEM/F-12 medium supplemented with FBS (10%) and penicillin-streptomycin (5%) at 5% CO2 and 37 °C. At confluence, cells were harvested using trypsin (0.25%) and sub-cultured into EIS chips (6 × 104 cells per well), 24-well plates (5 × 104 cells per well) or 12-well plates (106 cells per well) according to the selection of experiments. Cells were allowed to attach the surface for 24 h prior to treatment. CuO, CdO and TiO2 NPs were suspended in cell culture medium and diluted to the same concentration (100 μg mL−1). The appropriate dilutions of NPs were then sonicated using a sonicator bath at room temperature for 20 min (120 V/50–60 HZ) to avoid NPs agglomeration prior to administration to the cells. Following treatment, the cells were harvested to determine cytotoxicity, oxidative DNA lesion parameters. Cells not exposed to NPs served as controls in each experiment. Alamar Blue Assay: Cells are seeded in 24-well plates at confluence of 5 × 104 cells per well. One day later and at time point 0, medium or medium containing NPs (100 μg mL−1, final) are added on top of cells. Since incubation of Alamar Blue takes 6 h according to our prerunning test, the reagent of 10% sample volume was added at time point –6, 0, 6, and 24 h. Thus measurement of net absorbance at 570 nm was carried at time point 0, 6, 12, 30 h. Every data points are subtracted by reference number which comes from the reading of the mixture of medium and Alamar Blue only, and then normalized to the average of all the reading at time point 0. Cytotoxicity Assays: As mentioned above, cell suspension (0.6 mL) was applied into each well in the EIS chip for the experiments. After 24 h of cell attachment, prepared NPs solutions (100 μL) were added to corresponding wells. The resistance changes produced by the attachment of cells to the electrodes were monitored over a 40 h time period (both before and after NPs addition). The EIS chip design was previously reported.[34] In short, as cells are placed in each well of the chip, they settle down onto the electrode surface creating a barrier for the flowing current increasing the resistance measurements. Thus, it is possible to monitor the cell attachment and proliferation from the change in resistance measurements. Genotoxicity Biomarker Assays: For investigation of NP-induced oxidative DNA-damage, the cell culture medium were collected firstly at the end of the exposure period. After centrifugation, the supernants were recollected and stored at −20 °C until tested. The cells left in the wells and pellets in the tubes were resuspended and lysed by cell lysis buffer containing Tris (20 mM, pH = 8.0), NaCl (137 mM), Triton X-100 (1%), Glycerol (10%) and EDTA (5 mM). The cell lysates were obtained by collecting the supernant of the mixture after centrifuging. Paper strip assay: standard solution or sample (100 μL) was added onto the sample pad, and the solution migrated toward the absorbent pad; a result could be seen after 10 min.

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Biosensing for Rapid Genotoxicity and Cytotoxicity Assays

Statistical analysis: Results are expressed as means ± SEM of three experiments and data were analyzed quantitatively by using ImageJ and SigmaPlot software.

Acknowledgements This research and development project was supported by This research and development project was supported by the grant NIH R15 ES021079-01 and the grant W81XWH-10-1-0732 by US Army Medical Research & Materiel Command (USAMRMC) and the Telemedicine & Advanced Technology Research Center (TATRC).

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