Nanotoxicology, June 2011; 5(2): 254–268
Toxicity of gold-nanoparticles: Synergistic effects of shape and surface functionalization on micromotility of epithelial cells
MARCO TARANTOLA1, ANNA PIETUCH1, DAVID SCHNEIDER1, JAN ROTHER1, EVA SUNNICK1, CHRISTINA ROSMAN2, SEBASTIEN PIERRAT2, CARSTEN SÖNNICHSEN2, JOACHIM WEGENER3, & ANDREAS JANSHOFF 1 1
Institute of Physical Chemistry, University of Göttingen, Göttingen, Germany, 2Institute of Physical Chemistry, University of Mainz, Mainz, Germany, and 3Institute of Analytical Chemistry, Chemo- and Biosensors, University of Regensburg, Regensburg, Germany
(Received 28 July 2010; accepted 29 September 2010)
Abstract Nanoparticle exposure is monitored by a combination of two label-free and non-invasive biosensor devices which detect cellular shape and viscoelasticity (quartz crystal microbalance), cell motility and the dynamics of epithelial cell-cell contacts (electric cell-substrate impedance sensing). With these tools we have studied the impact of nanoparticle shape on cellular physiology. Gold (Au) nanoparticles coated with CTAB were synthesized and studied in two distinct shapes: Spheres with a diameter of (43 ± 4) nm and rods with a size of (38 ± 7) nm (17 ± 3) nm. Dose-response experiments were accompanied by conventional cytotoxicity tests as well as fluorescence and dark-field microscopy to visualize the intracellular particle distribution. We found that spherical gold nanoparticles with identical surface functionalization are generally more toxic and more efficiently ingested than rod-shaped particles. We largely attribute the higher toxicity of CTAB-coated spheres as compared to rod-shaped particles to a higher release of toxic CTAB upon intracellular aggregation.
Keywords: ECIS, micromotion, TER, D-QCM, gold nanoparticles
Introduction Although the concept of nanoparticle technology was pioneered in the 1960s, the chemical industry only discovered the benefits of this growing field in the early 90s (Oberdörster et al. 2007). Nowadays, ubiquitous applications can be found, whether it is on a big scale, i.e., in the automobile industry for friction coatings, for consumer applications or in novel medical research, ranging from targeted gene-delivery systems to imaging agents or synergistic/photothermal cancer therapy (Kumar 2006; Basu et al. 2009). However, predictions for the manifold increase in production of nanoparticles (60,000 tons by 2020) (Lewinski et al. 2008) run in parallel with the demand for strategic research on the risks upon environmental as well as human exposure, both for accidental or controlled delivery. As a consequence, high-throughput, dynamic toxicity-testing protocols
will be of great interest to support the design of safe engineering methods (Maynard et al. 2006). Gold nanoparticles (GNPs) attract considerable interest in biomedical research due to their inertness and high electron density. Only very few publications are dedicated to the shape- and size-dependent toxicity of GNPs (Chithrani et al. 2006; Chithrani and Chan 2007; Wang et al. 2008; Yen et al. 2009). With regard to surface ligands, citrate (Chithrani et al. 2006), as well as triphenylphosphin (Pan et al. 2007), and cetyl-trimethyl-ammonium bromide (CTAB) capping (Connor et al. 2005) has been analyzed among many others. Despite all these efforts to elucidate the fate of GNPs in mammalian cells, most studies lack time-resolved data from noninvasive techniques that reveal the kinetics of GNP interaction with living cells (de la Fuente et al. 2006; Takahashi et al. 2006; Khan et al. 2007; Nativo et al. 2008).
Correspondence: Prof. Andreas Janshoff, PhD, University of Goettingen, Institute of Physical Chemistry, Tammannstr. 6, Goettingen, 37077, Germany. Tel: +49 551 39 10633. Fax: +49 551 39 14411. E-mail:
[email protected] ISSN 1743-5390 print/ISSN 1743-5404 online 2011 Informa UK, Ltd. DOI: 10.3109/17435390.2010.528847
Toxicity of gold-nanoparticles In the work presented here, two label-free and noninvasive biosensor approaches based on AC impedance and quartz-crystal microbalance measurements were used to monitor specific aspects of cell physiology during GNP exposure in real time. Three of the aspects of GNPs of actual interest in life sciences as mentioned previously are addressed below: Shapedependent toxicity, uptake and intracellular effects. In particular, we have focused our work on the intracellular generation of reactive oxygen species (ROS) as a hallmark of an inflammatory response. Electric-cell substrate impedance sensing (ECIS) was introduced more than two decades ago (Giaever and Keese 1993) as a non-invasive biosensor to follow adhesion and spreading of mammalian cells by means of AC impedance measurements. In the ECIS set-up, the cells are grown on the surface of gold-film microelectrodes. The presence of the cell body increases the electrode’s impedance as the cell body behaves essentially like an insulating material forcing the current to flow around it. Accordingly, the measured impedance is dependent on the geometry of the current pathways beneath and round the cells, so that the device becomes sensitive for cell shape changes. Motility monitoring of confluent cells with the ECIS setup was first described in 1986 (Giaever and Keese 1986) using the electrical impedance read-out as an indicator for fluctuations of the individual cells within the cell layer. Thus, the signal holds information about the vitality of the cells under study (Lo et al. 1993). The second biosensor applied to study GNP toxicity is based on piezoelectric quartz disks that oscillate with only nanometer amplitudes. The parameters of the shear oscillation are sensitive to adsorption reactions at the resonator surface and hold information about the mechanical properties of the load material. The device is referred to as the quartz crystal microbalance (Wegener et al. 2001). Recording the resonance frequency and energy dissipation of the resonant oscillation when the resonators is exposed to biomolecules or cells provided numerous biophysical and medical applications due to the unprecedented sensitivity ( Δf f ≈ 10−8 ) (Cooper and Singleton 2007). In the case of cells, usually the adhesion and spreading behavior is followed. Fluctuations of these basic parameters were first described by Pax et al. (2005) as an analytical indicator to analyze cardiomyocyte contractions or to follow exocytosis and vesicle retrieval by Cans et al. (2001). Recently, we proved this technique to be sensitive enough to probe the metabolically-driven viscoelastic fluctuations of synchronized cells and fingerprint their micromechanical dynamics (Sapper et al. 2006; Tarantola et al. 2010).
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In the present work, the shape-dependent toxicity of GNPs coated with CTAB was studied by monitoring micromotion, viscoelastic fluctuations and viability of the epithelial cell line MDCK II upon cell-GNP encounter in combination with fluorescence imaging. The toxicity is sensitively mirrored in the cellular motility as recorded independently with both biosensor devices (Tarantola et al. 2009). We also applied a new method to quantify the intracellular concentration of nanoparticles using a combination of dark field microscopy and transmission electron microscopy. Moreover, we studied the impact of different GNPs on epithelial barrier function and found a shape-dependent breakdown shortly after exposure. These details of nanotoxicity are backed up by a classical, established cytological assay (MTS). Finally, generation of ROS as an indicator for inflammation was quantified and cell-cytoskeleton disruption as well as integrity of cell-cell contacts were visualized by immunochemistry.
Materials and methods Cell culture conditions and measurement procedures MDCK II cells were maintained in Earle’s minimum essential medium supplemented with 4 mM glutamine, 100 mg/mL of both, penicillin and streptomycin (purchased by Biochrom, Berlin, Germany), 10% (v/v) fetal calf serum (PAA Laboratories GmbH, Coelbe, Germany). Cells were stored in incubators (HERA cell 150, Heraeus, Germany) with a 5% CO2 atmosphere. Cells were subcultured weekly after reaching confluence by washing with PBS, followed by trypsinization and centrifugation at 110 g. Counting was carried out using a Neubauer chamber, and viability was determined using trypan blue exclusion. For the ECISbiosensor setup, 6 105 cells were seeded in the eight-well electrode arrays (Applied BioPhysics Inc., Troy, NY; electrode type 8W1E) and were treated as previously described (Tarantola et al. 2009). For QCM experiments, the resonators were cleaned thoroughly, assembled into specially designed teflon-chambers and transferred to a culture incubator with 37 C and 5% CO2 atmosphere (MMM Medcenter Einrichtungen GmbH, Gräfelfing, Germany). Cells were inoculated at a density of 6–7 105 cells/mL and handled as previously described (Tarantola et al. 2010). Exchange of medium with medium containing nanoparticles was carried out after a confluent cell layer had been established, i.e., 15–24 h after seeding. For microscopic studies, the cells were fixed by treatment with
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0.5% glutaraldehyde (GDA) (Sigma-Aldrich, Germany) solution and stained as described below.
Nanoparticle preparation Particle synthesis. Synthesis of rod-shaped particles followed the seeded growth method as described by Nikoobakht and El-Sayed (2003) and Jana et al. (2001). First, seeds were prepared by adding 0.6 mL of ice-cold 0.010 M sodium borohydride (NaBH4) to a 10 mL solution of 0.1 M cetyltrimethylammonium bromide (CTAB) containing 50 mL of 0.1 M tetrachloroauric acid (HAuCl4) under vigorous shaking. Second, rods were prepared by adding 12 mL of seed solution to a growth solution consisting of 75 mL of 0.1 M HAuCl4, 10 mL of 0.1 M CTAB, 7 mL of 0.04 M silver nitrate (AgNO3), and 105 mL of 0.08 M ascorbic acid (AA). Spherical particles were purchased from British Biocell International (Cardiff, UK). 1 mL aliquots of particles were precipitated by centrifugation (10 min, 10,000 g) and resuspended in 1 mL of 0.1 M CTAB solution in order to render surface chemistry consistent with preparation of the gold nanorods. Before usage, particles were washed by two centrifugation steps with deionized water, in order to remove excess of CTAB in solution avoiding cytotoxic side-effects, and were resuspended in HEPES buffered complete medium. Endotoxin tests of the supernatant were negative. Particle characterization. Nanoparticle size was assessed by transmission electron microscopy (TEM). In brief, samples were prepared by depositing a 10 mL drop of a solution with an approximated concentration of 6 1013 particles mL-1 on a carbon-coated copper grid (Plano GMBH, Wetzlar, Germany). After drying, images were taken with a Philips EM420 using an operating voltage of 120 kV. The mean nanoparticle size and its standard deviation resulted from the size measurement of at least 270 particles. We determined a length of 37.8 ± 6.5 nm and a width of 17.2 ± 2.9 nm for the nanorods and a diameter of 42.9 ± 4.0 nm for the nanospheres. Particle charge was determined by means of gel electrophoresis (Hanauer et al. 2007). Thermal gravimetric analysis (TGA) was carried out to measure the number of CTAB molecules attached to the surface. TGA allowed us to precisely measure very small weight variations of a sample, which are in the order of 1 mg. The instrument (TGA 7; PerkinElmer) had a noise level in the range of 0.3 mg and requires about 1 mg of starting material. Hence, 30 mL of the nanoparticle stock solution with a concentration of 3 1011 particles/mL exposed to 0.1 M CTAB
solution and subsequently washed twice with millipore water were concentrated to 50 ml by centrifugation. In order to remove water from the sample, it was heated to 105 C and kept at this temperature for 30 min. Afterwards the temperature was raised with a heating rate of 40 C per min to 800 C. For gold nanospheres, we found 32 mg CTAB per 1 mg gold, while for gold nanorods, 24 mg of CTAB per 1 mg was determined. Particle concentration. Concentrations of nanoparticle solutions were calculated from their extinction at spectra. UV-Vis spectra were recorded using a 1 cm thick precision cell (Type No. 105.202-QS, Hellma) and an OceanOptics USB2000 spectrometer set up with an OceanOptics HL-2000-FHSA halogen lamp. The extinction coefficient at 400 nm depends on the actual particle size and was calculated using a simulation software based on Mie-Gans theory in quasi-static approximation (Prescott and Mulvaney 2006, 2008). The simulation provided an extinction coefficient at 400 nm of 1.06109 L mol1 cm1 for the rods approximated as cylinder with spherical endcaps being 37.8 nm long and 17.2 nm wide as well as 6.18109 L mol-1 cm-1 for the spheres having a diameter of 42.9 nm. Stock solutions exhibited 6.21011 particles mL-1 for the nanorods and 1.11011 particles mL-1 for the nanospheres corresponding to 88.9 mg gold mL-1 for both species.
D-QCM-based viscoelastic monitoring, ECIS-based impedance studies and noise analysis Viscoelastic fluctuations of the cells were monitored with a self-made D-QCM set-up and the raw data of resonance frequency and dissipation was subsequently subjected to noise analysis algorithms as described elsewhere (Figure 1, left path) (Reiss et al. 2003; Tarantola et al. 2010). As described in detail there, short-time FFT (PSD estimation) and variance analysis of linearly detrended time series recordings were carried out. For power density spectra (PSD) analysis of the viscoelastic fluctuation data (referred to as F-QCM), the slope m was determined in the low frequency regime (Figure 1, lower part) and used as an indicator of the cells’ viability. Mean values from three independent experiments are presented. For ECIS studies, a home-made system was employed, as previously described (Tarantola et al. 2009); in our ECIS set-up, a 1 V AC signal was applied to the cell-covered electrodes through a 1 MW series resistor, and the in- and out-of phase voltages across the electrodes were recorded at 4 kHz at a sampling rate of 1 Hz (Figure 1, right path, based
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Figure 1. Data acquisition: Time-resolved recordings of the parameters f and j Z j norm, 4kHz were subjected to FFT analysis yielding PSD (power-spectral density)-diagrams. Linear slopes in the low-frequency regime of the PSD serve as measure for cell motility.
on Giaever and Keese 1986). For micromotion recordings, fluctuations of the in-phase voltages were recorded with time and Fourier transformed after subtracting a linear trend (Figure 1, lower part), as described recently. A linear fit of the low frequency part of the power density spectra provided slopes up to 2.7 for living cells. Besides micromotion recordings at one fixed frequency, we also monitored changes in the barrier function of the adherent MDCK II cell layer in a frequency range between 10 and 104 Hz. The transepithelial electrical resistance (TER), a well-accepted measure for epithelial barrier function, was extracted from these frequency-resolved impedance readings by equivalent circuit modeling (refer to Wegener et al. 1996). Micromotion and TER readings were carried out on one and the same cell layer as a function of particle type and concentration.
Dark-field- and transmission electron microscopy Confluent cell layers were incubated with a fixed concentration of 15 mg/mL of gold, corresponding to 104 gold nanorods or 2 103 gold nanospheres per cell. After 20h incubation, cells were trypsinized, fixated and embedded in an epoxy resin for preparation of cell sections for dark-field (DF) and transmission-electron-microscopy (TEM). For fixation, cells were incubated for 1 h with 3 mL of 2.5% glutaraldehyde in cacodylate buffer (0.1 M sodium cacodylate and 0.1 M sucrose at pH 7.4), washed three times with cacodylate buffer, incubated for 1 h with 1 mL of 2% osmium tetroxide (OsO4) in cacodylate buffer, and were finally washed twice with deionized water. For embedding, these cells were exposed to 2% (w/v) agar, which was dehydrated using an increasing ethanol concentration series (30, 50, 70, 80, and 96%
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absolute), and transferred into araldite epoxy resin with the help of propylene oxide. The resin was polymerized by heating to 65 C for three days. Cells in resin were cut in 1 mm or 50 nm thick sections using a Reichert Ultracut S ultramicrotome. Thin sections of 1 mm thickness were placed on freshly cleaned microscope slides for optical dark-field microscopy and ultra-thin sections of 50 nm thickness were transferred to copper grids for TEM. Contrast enhancement of biological tissue for TEM was achieved by incubation of the sample in a solution of 2% uranyl acetate in 50% ethanol for 2 min (Hayat 2000) and a lead salt solution prepared according to a modification of Sato’s method for 2 min (Hanaichi et al. 1986). Before TEM investigation, samples were rinsed thoroughly with deionized water and dried in air. Cytotoxicity assay, immuno-/fluorescence-microscopy The MTS cytotoxicity test was applied according to the manufacturer’s protocol (Promega, Mannheim, Germany). Briefly, cells were grown to a predetermined density of 12,000/well in a 96-well plate and were subsequently incubated with eight different concentrations of nanoparticles for 24 or 48 h. Washing was carried out three times with PBS and full medium before adding the MTS agent. Control experiments were performed using cell-free wells and fully vital cells. The cells were incubated with the tetrazolium educt for 45 min and absorbance was determined using a 96-well plate photometer at a wavelength of 490 nm. The color change was a direct measure of the cell’s metabolic activity due to the reduction of MTS educts to formazan by the action of mitochondrial dehydrogenases. Experiments were performed in triplicate. Immunostaining and fluorescence microscopy were applied to monitor alterations in the cell cytoskeleton and cell-cell contacts upon nanoparticle exposure as well as any inflammatory response generating reactive oxygen species. Therefore, MDCK II cells were grown to confluence on Petri dishes and were then incubated with nanoparticles for 24 h. For actin and microtubules staining, washing with PBS and fixation was carried out by immersing the cells into a 20 C cold acetone/methanol mixture (1:1 vol %) for 10 min. Afterwards, the cells were washed three times with PBS, non-specific binding sites were saturated with FCS, and incubation in staining solutions was carried out according to the manufacturer’s recommendation: Alexa Fluor 488 phalloidin (Invitrogen, Paisley, UK) was used for f-Actin staining, Alexa Fluor-conjugated IgG1 anti-b-tubulin (BD Bioscience, Heidelberg, Germany) from mouse for labeling microtubules, 4¢,6 diamidino-2-phenylindole (DAPI,
Sigma-Aldrich, Seelze) for nucleus/DNA labeling, and polyclonal IgG1 mouse antibody (Zymed Gmbh, Munich) followed by Alexa Flour-conjugated goat-antimouse IgG1 antibody (BD Bioscience, Heidelberg, Germany) were used for staining tight junctions (ZO-1 staining). Staining was carried out for 30 min at room temperature, and cells were washed several times after staining was terminated. Generation of ROS was followed by life cell staining of cells after 24 h exposure to CTAB-GNPs: 5-(and6)-chloromethyl-2¢,7¢-dichlorodihydrofluorescein-diacetate acetyl ester (CM-H2DCFDA) (Molecuar Probes, Invitrogen) was added to GNP-treated cells and the following controls were carried out: Negative control (autofluorescent untreated samples as well as autofluorescent, GNP-treated cells) and positive controls (inflammation induced by 0.03 v/v % H2O2 treatment for 30 min before staining). CMH2DCFDA appears green when oxidized by ROS; Co-staining with DAPI was applied to illustrate the topology of the cell-monolayer. The samples were imaged using an upright Olympus fluorescence microscope (Olympus BX51, 40 or 100 water immersion objectives [NA = 0.8 or 1.0] Germany), equipped with a color camera (Olympus DP71) and a dark-field condensor. Distribution of gold nanorods was visualized by dark-field microscopy, where plasmon resonance leads to green (spheres), red (rods) or orange (aggregates) color. Results Epithelial cells exposed to GNPs – influence of shape on viability and adhesion It was the primary objective of this study to elucidate whether the cytotoxicity of gold-nanoparticles (GNPs) was dependent on the shape of the particles. Thus, we studied the impact of rod-shaped (r) and spherical- (s) gold-nanoparticles (GNPs) on cell physiology using two non-invasive biosensor devices in addition to classical imaging techniques such as fluorescence, darkfield and TEM imaging. Whereas the microscopic techniques were important to visualize particle uptake and structural alterations within the epithelial cell layers, the biosensor read-outs provide time-resolved information about the impact of GNPs on more functional parameters such as cytomechanics, cell motility and tightness of epithelial junctions. We recently showed that metabolically driven micromotion of cells in a confluent monolayer grown on gold-microelectrodes – monitored via ECIS – serves as an indicator for dose-dependent nanoparticle toxicity for various nanoparticles: Semiconductor quantum-dots (Tarantola et al. 2009), synthetic
Toxicity of gold-nanoparticles HPMA-nanospheres (Barz et al. 2008) and multifunctional, spherical magnetic MnO-nanoparticles (Shukoor et al. 2009). In a first step, the impact of differently shaped GNPs on MDCK II cells was quantified by the classical MTS cytotoxicity test. Figure 2A shows exemplarily exposure to rod-like (r) and spherical (s) particles at an extracellular concentration of 9 mg/mL for an incubation time of 24 h. Corresponding to the plasmon resonance in dark-field microscopy, red color was chosen for rods (r), green for spheres (s) throughout the present work in all diagrams and plots. The detergent-coated CTABrods (r) did not reduce the mitochondrial enzyme activity significantly indicating no apparent cytotoxicity. However, the spherical CTAB-coated particles (s) showed a massive toxicity in this concentration regime with cell viabilities close to zero even though the same dose of GNPs was applied. These initial experiments demonstrated a severe difference of the biological response to particle shape. To get insight into the time scale of the observed toxicity, we studied the impact of the same dose of particles by time-resolved ECIS measurements. Figure 2B exemplarily depicts the time course of the impedance |Z| at a sampling frequency of 4 kHz when confluent cells were exposed to spherical and rod-shaped GNPs (9 mg/mL), respectively. The numerical values have been normalized to the impedance magnitude of the cell-free electrode prior to cell inoculation. The impedance magnitude as returned by ECIS provides a time-resolved analysis of GNPs impact on epithelial barrier function and cell layer integrity. The time-course for r-GNPs was not significantly different from the control population that was not exposed to NPs at all. There was only a slight drift of the signal for both conditions. S-GNPs, however, produced a sharp drop in the cells’ impedance of
60–70%, indicating a complete loss of epithelial barrier function due to particle encounter. The residual impedance compared to a cell-free electrode was due to the fact that the dead cells did not detach completely from the gold surface. Changes in cell dynamics as a function of GNP exposure – concentration-dependent toxicity and particle uptake Recently, we reported on impedimetric and acoustic motility assays based on reading cellular shape fluctuations by ECIS or recording viscoelastic fluctuations of the cell body by means of piezoelectric resonators (Tarantola et al. 2009; Tarantola et al. 2010). We could unequivocally show that ECIS micromotion was suitable to detect subtle changes in the metabolically driven fluctuations of the cell body when the cells were exposed to nanoparticles. F-QCM is appropriate to fingerprint fluctuations in the viscoelasticity of the cell body most likely related to cytoskeletal dynamics. Here, we adopted these two inherently different assays to elucidate the dose-dependent impact of GNPs on cellular dynamics as a function of particle shape. Figure 3A summarizes ECIS micromotion studies of MDCK II cells exposed to differently shaped GNPs at 9 mg/mL for 60 h. Each data point represents a snapshot of the cellular micromotility along the observation period. Analysis of the time series data to get a quantitative indicator for cellular motility is described above and illustrated in Figure 1. Green symbols denote spherical particles, while red symbols represent rodlike particles. For comparison, micromotion data of an untreated control population and a cell-free electrode are added in gray and black, respectively. Similar to the control population, MDCK cells exposed to r-GNPs showed no indication of a substantial toxicity. In
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Figure 2. (A) MTS-test for mitochondrial activity: two shapes of GNPs (9 mg/mL) were added to confluent MDCK II cells. Red color was chosen for rods (r), green for spheres (s). (B) Time courses of impedance j Z j norm, 4kHz (normalized to impedance values of the cellfree electrode prior to cell inoculation) at 10 min time resolution. Inoculum of 6 105 cells was exposed to 9 mg/mL CTAB-coated GNPs: untreated cells, empty electrode, s-GNP, r-GNP. j Z j norm, 4kHz serves as a measure for epithelial barrier function/cell layer integrity and is strongly reduced for s-GNP-treated cells. Particle addition occurred upon confluence.
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contrast, s-GNPs at the same concentration reduced viability to the same level as a cell-free electrode within 30 h of incubation. The cells were significantly affected by GNP exposure and no longer capable of performing their native cell shape fluctuations. In summary, at this rather low concentration (see extended doseresponse study below), micromotion measured by ECIS indicated a loss of cellular dynamics upon sGNP but not r-GNP exposure. We also carried out F-QCM-noise analysis in order to monitor viscoelastic fluctuations as a response to particle uptake. Figure 3B shows the time course of normalized micromotion that quantifies the extent of frequency fluctuations for comparable experimental conditions as studied in Figure 3A. Details about the PSD analysis of the frequency fluctuations are given above. It is clearly visible, that r-GNPs were less potent to reduce the viscoelastic fluctuations than their spherical counterparts, which reduced cellular fluctuations down to the level of the cell-free resonator (black lines). The data also showed that the viscoelastic noise parameter as it was recorded here is a little less reliable than the indicator for ECISrecorded cell-shape fluctuations. Figure 3C and D compare the dose-dependent toxicity of s-GNPs and r-GNPs in MDCK cells as recorded by ECIS (Figure 3C) or QCM (Figure 3D). In either case the fluctuation-based toxicity readout (full red circle, full green circle) is compared to
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the more classical MTS assay addressing mitochondrial activity (open red square, open green square). Reduction of viability was found for spherical and rod-like CTAB-GNPs, albeit at higher concentrations. In general, micromotion recorded by ECIS was more sensitive to the impact of particle exposure than the classical photometric MTS assay, while F-QCM recordings evaluated by PSD-analysis responded less sensitive to cytotoxic changes of cell-motility, at least at low concentrations Table 1. A quantitative statement on the toxicity of nanoparticles heavily relies on the exact knowledge of both the intracellular and extracellular concentration of particles. The cellular response to exposure depends on the uptake efficacy and needs to be determined independently of toxicity effects. Previously, we have developed a protocol that allows us to determine the number of nanoparticles in an individual cell based on a combination of dark-field microcopy and transmission electron microscopy (manuscript in preparation). In brief, dark-field microscopy was used to estimate the number of aggregates per cell, while TEM was used to compute the number of particles per aggregate. A combination of the two methods provided us with an accurate number of particles inside the cell. Micrometer-sized slices of cells (1 mm) were used for dark–field microscopy to determine the aggregation number of particles within the cells, while thinner (50 nm) slices were used for TEM
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Figure 3. (A) ECIS-micromotion time courses for GNP treated MDCK II cells (9 mg/mL): CTAB rods ( ), CTAB-spheres ( ) as well as references: untreated cells ( ) and noise generated from bare electrodes immersed in culture medium (.). (B) F-QCM micromotion for GNPexposed MDCK II cells (same color code as in A). Nanocytotoxicity of GNPs as determined by micromotion compared to MTS as function of concentration for ECIS (C) or F-QCM (D) after 24 h of incubation (micromotion [ , ] as compared to MTS assay [ , ]). Each data point represents the average of three independent measurements.
Toxicity of gold-nanoparticles Table I. IC50 values for the impact of gold-nanoparticles for 24 or 48 h exposure time on the viability of MDCKII cells as determined by MTS, ECIS-micromotion and F-QCM-PSD-fluctuation analysis; values are given in mg/mL. Mean SD ± 0.1 for MTS, ± 0.15 for micromotionECIS and ± 0.5 for micromotionF-QCM (n = 3). Nanocytotoxicity assay
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to assess the number of particles per aggregate (manuscript in preparation). Figure 4 shows exemplarily dark-field microscopy and TEM images of s-GNPs inclusions. We found that exposure of cells to an external concentration of 15 mg/mL GNP
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resulted in an intracellular concentration of 2000– 3000 particles per cell, regardless of their shape. Moreover, we found that particles aggregate mainly inside early endosomes forming clusters of 48 ± 18 (r) or 44 ± 12 (s) particles/aggregate. We also studied aggregation and sedimentation behavior of both types of particles in medium and distilled water by means of absorption spectroscopy (Supplementary Figure 1, available online). Particles suspended in culture medium displayed extinction spectra shifting slightly to longer wavelength, which was indicative of protein adsorption from the medium. This was in good accordance with the finding that particles sediment in culture medium within 15 h while remaining dissolved only in distilled water. Sedimentation velocity is identical for both particle types (Supplementary Figure 2, available online) essentially imposing the
Figure 4. (A) Dark-field image of 1 mm thick slices after s-GNP incubation for 20 h, showing aggregates (yellow and orange) or single spherical GNPs (green) within cell boundaries. (B) TEM image of s-GNP treated cells depicting macropinocytosis of singular particles (see inlet) and aggregation inside early endosomes. (C) TEM-slice of an MDCK II cell undergoing vacuolization upon s-GNP-uptake/particle aggregation (15 mg/mL) inside multivesicular bodies. (D) TEM-image of cells showing initial s-GNP escape from multivesicular bodies or late endosomes/ lysosomes into the cytosol and severe blebbing/vacuolization. (E) TEM-slice of an MDCK II cell after exposure to r-GNPs (15 mg/mL). (F) r-GNP clusters trapped in vesicular structure inside the cell (incubation time for C-F is 20 h).
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same dose on the cell layer irrespective of the particle type (Teeguarden et al. 2006). The sediment can be easily resuspended by gently shaking the cuvette, suggesting that the sediment consisted only of loosely connected particles and not an irreversible precipitate. From sedimentation experiments we concluded that the local particle concentration exposed to the cells was identical for both types of particles at any given time and did not explain the distinct toxicity of spherical and rod-like particles. The micrographs in Figure 4A and B show the uptake of s-GNPs by MDCK II cells. The pool of all images indicated an uptake of roughly the same number of particles per cell. Due to the individual size and geometry, the amount of gold taken up by the cells was, however, shape-dependent and should match the ratio of the particle volume: s-GNPs have approximately five times the volume of r-GNPs and display a 2.8 times larger surface area. Thus, the amount of gold per cell was five times higher for s-GNPs albeit the number of GNPs within the cells was virtually identical. Qualitative TEM analysis revealed (inlet in Figure 4B) a typical non-specific uptake pathway: Macropinocytosis. Figure 4B shows aggregated s-GNPs after uptake. In Figure 4C and D, two other cells of the same sample are shown. The cell in Figure 4C shows initial vacuolization which we assumed to occur due to CTAB desorbing from aggregating particles. This implied that GNPs serve as carriers for toxic CTAB, as speculated previously (Tarantola et al. 2009). Another process was visible in Figure 4D: Besides the aggregates in late endosomes or lysosomes, initial endosomal escape and severe vacuolization indicative of cell stress and necrosis could be observed. Aggregation of GNPs in endosomes was also found for rods, albeit without strong vacuolization (Figure 4E), but r-GNPs stay locked in lysosomes (Figure 4F). Strong vacuolization was only found for spherical GNPs. Notably, we checked that removal of excess CTAB after centrifugation was complete by perfoming MTS tests on the supernatant (data not shown). Adverse effects of GNP exposure – generation of reactive oxygen species and cytoskeletal integrity The molecular reason for changes in cell motility and/or viscoelasticity is most likely based on and reflected by changes in cytoskeletal integrity. Hence, we studied the integrity of actin or microtubule filaments under GNP influence by fluorescence microscopy. To pinpoint a possible mechanism that transmits GNP toxicity to the cytoskeleton, we first studied the induction of cell stress upon
GNP incubation, as measured by the production of reactive oxygen species (ROS). Fluorescence micrographs in Figure 5A provide an image of the proinflammatory response upon GNP exposure by probing the presence of ROS, which produce green fluorescence. All samples were counterstained with DAPI to label the nucleus, a treatment which did not provoke ROS by itself (negative control in Figure 5 A1). Figure 5A2 provides the corresponding positive control: Cells were exposed to 0.03% H2O2 for 30 min. The positive control clearly indicated the presence of ROS by a bright green fluorescence across the entire cytoplasm. Figure 5A3 shows the situation of MDCK II cells exposed to 6 mg/mL r-GNPs for 24 h. The image shows a few centers of ROS formation in merely a small fraction of cells. In contrast, MDCK cells exposed to s-GNPs (Figure 5A4) show massive ROS production either in point-like centers – which could be in membrane-enclosed compartments – or in diffuse areas of the cytoplasm, which might indicate rupture of the compartments involved in ROS formation. Figure 5B shows overlays of the combined Alexa-546-Phalloidin staining for f-actin (red), immunofluorescence staining of microtubules with alexa-488-marked mouse-anti-ß tubulin (green) and DAPI-DNA staining of the nucleus (blue). The fluorescence micrograph in Figure 5B1 represents a control population of cells 24 h after reaching confluence (48 h after seeding). The cells show a junctional actin ring, as it is typically found in barrier forming epithelial cells, long stress fibers and densely packed actin throughout the cytoplasm. Thick bundles of microtubules were found around the nucleus and homogenously distributed in the whole cell. Figures 5B2 and Figure 3 illustrate cytoskeletal integrity after exposing the cells to either r- (Figure 5B2) or s-GNPs (Figure 5B3), respectively. Images are split along the diagonal in order to display actin and microtubules simultaneously without interference from overlaying. For cells treated with r-GNPs (Figure 5B2), the junctional actin ring is partially interrupted and spacing between the cells seems extended, the microtubular degradation is significant, but still single long fibers in the cell extensions were found. In the case of s-GNPs (Figure 5B3) applied at identical concentrations, whole stress fibers were missing and the junctional actin ring was absent. The severe degradation of the dense network was accompanied by large intercellular clefts. Microtubules were only found in perinuclear arrangement, indicating complete degradation in the cytoplasm and/or storage of monomers in Golgi-associated vesicles. Therefore, it is
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Figure 5. Fluorescence microscopy images after co-staining of actin (red), microtubules (green), nucleus (blue) and/or reactive oxygen species (green). (A) Proinflammatory ROS generation upon GNP exposition (24 h, 6 mg/mL per GNP category, CM-H2DCFDA-staining): (1) Untreated cells; (2) 0.03% H2O2 treated cells, (3) r-GNPs, (4) s-GNPs. (B) Cytoskeletal integrity of MDCK II after exposition to GNPs: (1) Untreated cells; (2) 3 mg/mL r-GNPs; (3) 3 mg/mL s-GNPs (incubation time for A and B is 24 h).
safe to assume that the origin of variations in motility (ECIS and QCM) was based on the partially degraded cytoskeleton as a result of GNP exposure. GNPs influence on epithelial cell-cell contacts – barrier breakdown and recovery after exposition So far we investigated the influence of GNP exposure to cells on their adhesion (Figure 2B) and motility (Figure 3). In this section, we concentrate on changes of the barrier-forming properties of MDCK II cells as a function of particle shape. Readings of the transepithelial resistance or TER
(normalized to the electrode area) were used as a functional measure for epithelial barrier integrity, combined with fluorescence microscopy of structural modulations (Figure 6). The grey line in Figure 6A represents the control, i.e., the change in TER of cells that are not exposed to GNPs. Applying r-GNP (red curve, 9 mg/mL) resulted in a time course following the same general trend as the untreated sample. However, a more pronounced initial minimum of the TER was observed in between 20 and 30 h after particle addition, which was a slight indication of temporary cell-cell contact dissolution. Remarkably, the final value of the TER did not significantly differ from the untreated control. On
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Figure 6. (A) Time course of TER after addition of 9 mg/mL CTAB-coated GNP to confluent MDCK II cells: s-GNPs ( ); r-GNP ( ); untreated cells ( ). (B) Overlay of dark-field images and fluorescence images with e-cadherine (right) and ZO-1 (left) as well as DAPI staining of the nucleus after 24 h incubation with r-GNPs or s-GNPs. Plasmon resonance renders single spherical particles to appear green, rod-shaped particles red and aggregates orange to white.
the contrary, s-GNPs also added at a concentration of 9 mg/mL led to an irreversible loss of the barrier function within 24 h after exposure. Therefore, it can be deduced, that the TER responded extremely sensitive to the shape of nanoparticles. Data presented in the preceding paragraphs indicate that s-GNPs, but not to the same extent r-GNPs, induced the formation of ROS inside the cells which causes structural and functional alterations of the cytoskeleton. This is perfectly in line with our observation expressed in Figure 6A, which shows a severe breakdown of epithelial barrier function as a response to s-GNPs. Thus, it is most likely that the structural alterations of the cytoskeleton go hand in hand with a structural disintegration of epithelial cell-cell junctions. To address this issue, we exposed MDCK cells to r-GNP and s-GNP for 24 h, respectively, before we stained the samples for cell junctions associated proteins. Figure 6B presents fluorescence micrographs of both cell populations. Whereas incubation of MDCK II cells with r-GNPs (left) provided a staining pattern that was typically found in untreated epithelial cells (all cells in the field of view had a continuous staining around the cell periphery indicating wellestablished tight junctions) (Supplementary figure 3, available online)), incubation with s-GNP led to a complete loss of the typical cell border staining (right).
Investigating the interaction of nanoparticles with mammalian cells is an urgent task considering the worldwide increasing number and variety of available particles together with their intended or nonintended introduction into the biosphere. Classical cytotoxicity tests addressing the vitality of cells after a pre-defined time are time-consuming and might not be sufficient to capture subtle changes in the biological activity of cells. In the present work we addressed the toxicity of nanoparticles in contact to cultured epithelial cells by monitoring cellular dynamics from different viewpoints using non-invasive biosensor techniques together with optical and biochemical experiments. Changes in cellular dynamics upon exposure to gold nanoparticles of different shape were recorded by means of electric cell substrate impedance sensing (ECIS) and fluctuation mode quartz crystal microbalance (F-QCM), respectively. Previously, we showed that monitoring micromotion of cells by recording resistance fluctuations using ECIS was well suited to capture nanoparticle toxicity and even more sensitive compared to classical tests such as MTS (Tarantola et al. 2009). In recent studies, we and others (Chithrani and Chan 2007; Alkilany et al. 2009; Tarantola et al. 2009; Arnida et al. 2010; Rayavarapu et al. 2010) found that surface functionalization plays a dominant role in toxicity of gold nanoparticles. In our previous publication we showed the principle suitability of ECIS-based micromotion assays to assess the toxicity of nanoparticles in a noninvasive fashion in real time (Tarantola et al. 2009). We could show that surface chemistry is decisive for both uptake and cytotoxicity. So far, however, we did not address the impact of particle shape in conjunction with surface chemistry. Moreover, no protocol was available to determine the amount of particles taken up by the cells as a function of exposure time and concentration. Here, we address the impact of particle shape on biological activity and structural integrity of epithelial cells in addition to toxicity arising from surface chemistry employing two different micromotion assays at defined intracellular particle concentrations. Rodlike gold nanoparticles with geometric dimensions of approximately 40 20 nm2 were compared to spherical ones with a diameter of approx. 40 nm and an identical surface chemistry. A prerequisite for any quantitative comparison of particle toxicity is exact knowledge of the uptake efficiency. We developed an approach based on a combination of dark field and transmission electron microscopy to count the number of particle within a cell (manuscript in preparation). In brief, adding the same amount of gold
Toxicity of gold-nanoparticles (15 mg/mL) to cultured MDCK II cells resulted in roughly the same number of particles within the cells. For both particle shapes we found approximately 2000–3000 gold nanoparticles per cell. Noteworthy, the spherical particles were approx. 5 times larger in volume than the rod-shaped ones leading to a higher gold content inside the cell. The surface area ratio between the two particle amounted to 2.8, which might be indicative of a higher possible CTAB release from spherical particles inside the cell upon aggregation. Taken together we can safely state that based on the experiments described above, the specific toxicity per particle is significantly higher for s-GNPs relative to r-GNPs. All functional parameters that were recorded throughout this study consistently indicate this geometry-dependent specific toxicity: (1) Mitochondrial activity was drastically reduced by spheres but not by rods; (2) Metabollically driven shape fluctuations – as recorded by ECIS – were more strongly affected by spheres not by rods; (3) Active viscoelastic oscillations within the cell body – as recorded by F-QCM readings – were significantly more reduced by spheres than by rods; and (4) Epithelial barrier function was erased by spheres but left unchanged by rods. These functional observations go hand in hand with our more structural experiments that revealed an impact of GNPs on the intracellular cytoskeleton – mediated via the formation of ROS. The different aspects of the observed shape-dependent toxicity and a mechanistic model to explain the observations is discussed in more detail below. Particle uptake and cell dynamics Even though the spherical particles were applied in five times smaller particle concentration than the rodshaped, the intracellular particle concentration was found to be similar indicating a more efficient uptake of spherical particles. The reasons for this surprising finding are not yet clear. Both particle types were decorated with a CTAB bilayer which rendered them displaying the same surface chemistry. The only obvious differences between both particle populations were their geometry. But from our studies we could not provide a clear-cut explanation of how these geometric parameters affected particle uptake. We know from TEM studies that both types of GNPs are ingested by macropinocytosis. So a different membrane interaction of both particle geometries might be the key to explain the observed shape-dependent uptake. Another important feature
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that we need to address in further studies in more detail is protein adsorption from the cell culture medium to the particle surface. If both particles are coated differently with medium-borne proteins this may affect and explain their individual uptake characteristics. One might argue that the bending energy to be paid for wrapping particles with the plasma membrane is different for rod-shaped than for spherical particles explaining the higher uptake of s-GNPs. In fact, the bending energy needed for forming a bilayer sphere is independent of the radius and amounts to Esphere = 4p (2kb + kG) with k b the bending modulus (»1019 J) and k G the Gaussian curvature that is usually neglegible, while that of a cylinder with an aspect ratio of 1:2 requires Ecylinder = 2pk b. Hence, we conclude that wrapping of single particles is most likely not responsible for the different uptake efficiency, since cylinders with an aspect ratio of 1:2 require less energy to be wrapped given that the adhesion energy is identical. Since our TEM images suggest that particles are internalized via macropinocytosis, wrapping of individual particles does not play a decisive role. Once in the cytoplasm in similar concentrations both particle types show a very different toxicity profile according to all biophysical and biochemical assays performed in this study. Compared to the mitochondria-specific MTS assay, ECIS- and QCM-based experiments are more holistic approaches that report on alterations in the dynamics of the cell body as a whole – without assigning the observed changes to subcellular structures. It has been reported in several previous studies that assays reading integral parameters of cell physiology are more sensitive to detect subtle cell responses to xenobiotics compared to biochemical assays addressing one specific enzyme activity or the rate of one specific metabolic pathway (Hug 2003). Both biosensors are tuned to read the inherent dynamics of the reporter cells in real time which additionally provides a time-resolved view on the cell response. The different assays support one another as they all clearly indicated the dose-dependent toxicity of the particles. Comparing both biosensor approaches, it becomes obvious that the shape fluctuations as recorded by ECIS are more sensitive than the viscoelastic fluctuations extracted from F-QCM-readings. However, for a fair comparison it is important to recognize that the number of cells that contribute to either read-out are very different. ECIS-based experiments integrate over as many cells that cover the active electrode with an area of 510-4 cm2. In QCM experiments, the number of cells contributing to the signal is significantly bigger and corresponds to approximately 310-1 cm2 which is three orders of magnitude bigger. When fluctuation data is integrated over huge numbers of cells, the individual cellular fluctuations can easily average out
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so that the integral noise is smaller than for small populations. We think that the difference in the number of cells that contribute to the fluctuating time series account for the observed sensitivity differences between both techniques. It is, however, also important to recognize that both techniques are independent and probe different dynamic aspects of the cells: Cell shape fluctuations are traced with ECIS, whereas cytomechanical fluctuations are recorded by F-QCM. Proposed explanation for shape-dependent particle toxicity The individual particle-specific toxicity of rod-shaped and s-GNPs is surprising as the particles share the same core, the same surface chemistry and they are present in the cytoplasm in similar concentrations. The reason for the observed individual toxicity can therefore only arise from the individual particle geometry and associated secondary effects such as different degree of intracellular particle clustering or individual loss of CTAB molecules from the surface as a consequence of clustering. TEM images were qualitatively analyzed to identifiy the uptake mechanisms. We found that macropinocytosis prevailed as the major uptake pathway – rather expected for particles of this size. Uptake of particle still occurred after 20 h, thus continuous accumulation of particles took place. Additionally, we found that MDCK II cells displayed vacuolization upon GNP-aggregation inside early endosomes or multivesicular bodies within the same time interval for both types of particles. There were clear indications for particle escape from multivesicular bodies or late endosomes/lysosomes that correlated with severe blebbing/vacuolization. MTS-tests of CTAB-coated GNPs incubated with cells at 4 C (data not shown) supported our assigment of an active vesicular uptake mechanism, as almost no toxicity was detected at this temperature due to the lack of membrane engulfment. Moreover, the 4 C experiment proved that simple interactions of the GNPs with the outer leaflet of the cell membrane were not responsible for the strong toxic effects found for s-GNPs. The observed toxicity requireed access to the intracellular environment. Since dark field microscopy and TEM indicated that aggregation of particles occurred inside early endosomes and multivesicular bodies, as has been described elsewhere (Connor et al. 2005), we believe that the desorption of the protein-corona (Cedervall et al. 2007; Lynch et al. 2007; Lundqvist et al. 2008) and the intracellular release of surface-bound CTAB upon particle clustering accounts mainly for the observed toxicity. We suggest that CTAB is liberated from clustering GNPs within the endosomal
environment, destroying inner membranes and provoking the production of ROS as well as the cytoskeleton degradation. Alkilany et al. (2009) suggested that cell toxicity of CTAB-coated GNPs arises from free CTAB in solution after desorption from the particle surface. We found, however, that the addition of pure CTAB to the culture medium, in a concentration that reflected the situation as if all CTAB molecules were released from the particles, was not toxic and thus did not explain the observed response of the cells. Along these lines, we observed in TEM images and darkfield microscopy a slightly stronger propensity of s-GNPs to form intracellular clusters as compared to r-GNPs, which might explain the increased level of toxicity observed for spherical particles. Together with a 2.8 times larger surface area, spherical particles might release just enough CTAB to display the observed lower IC50 values. It is, however, also conceivable that other factors inside the cell, which sense geometry by reporting on curvature, might be responsible for the observed difference in toxicity. Notably, the use of PEGylated gold particles (data not shown) did not lead to any usable results in respect to the impact of shape on biological activity since not enough particles entered the cell to produce a cellular response. According to the suggested mechanism, nanogeometry is only indirectly responsible for the observed toxicity (Ferrari 2008; Jiang et al. 2008). This conclusion is further supported by the fact that some groups described already gold-nanoparticles of ‡15 nm to be inert like bulk gold (Shukla et al. 2005; Khan et al. 2007). Loss of actin fibers and at higher particle concentration even complete microtubule degradation was found as a result of the intracellular presence of CTAB coated GNPs. Since CTAB is not only used for seeded growth or as phase-transfer-catalyst in chemistry, but also in biology for complexation of nucleic acids or protein/polysaccharide aggregation, the latter effect could account for the detected degradation. It remains to be elucidated whether the intracellular presence of CTAB, which entered the cells as surface attached molecules decorating the GNPs, triggers the formation of ROS which then accounts for a cascade of events that eventually kills the cells. Since we detected ATP-depletion via MTS, mitochondrial damage and several morphological characteristics of apoptosis, this programmed cell death pathway is in the focus of current investigations. Nanoparticle impact on cell-cell-contacts Impedance measurements on epithelial cell monolayers such as the MDCK cells used in this study monitor changes in the epithelial specific barrier function with non-invasively and with outstanding
Toxicity of gold-nanoparticles time resolution. Nanotoxicity studies addressing epithelial barrier function have only been sparsely performed so far for nano- and microparticles. The existing studies have indicated no or only slightly increased permeability upon exposition (Chanana et al. 2005; Grenha et al. 2007; Moyes et al. 2007; Rothen-Rutishauser et al. 2009). While CTABspheres lead to a concentration dependent, immediate breakdown of the barrier integrity, CTAB-rods have basically no impact on epithelial cell junctions. ZO-1and e-cadherine immunofluorescence staining after 24 h of incubation confirm the complete loss of tight junctions after exposure to spheres but not to rods. These findings are most likely a direct consequence of the cytoskeletal changes that were induced by intracellular presence of the GNPs. It is an established fact that changes in the actin cytoskeleton as it is brought about by cytochalasin D treatment induce an immediate and severe breakdown of the epithelial integrity. Conclusion In vitro nanocytotoxicity studies are crucial steps in defining strategic safety mechanisms against the possible risks upon environmental as well as human exposure to nanomaterials. Although long-term in vivo effects such as, i.e., accumulation, failure to degrade and chronic inflammation, are thereby neglected, predictions upon the factors triggering toxicity can be made. Especially time-resolved biosensors with higher sensitivity, faster read-out, non-invasiveness are desirable for high-throughput screening systems that are required to respond to the tremendous increase in nanoparticle synthesis. Here, we have presented a double biosensor-approach making use of F-QCM and ECIS to follow cell dynamics as a measure for toxicity. With the help of fluorescence imaging of actin, microtubules, ROS and adherens/tight junctions under GNP influence, we were able to shed light on the mechanisms responsible for an apparent shape-dependent toxicity of gold nanoparticles. We found that spherical gold nanoparticles are generally more toxic than rod-like and lead to irreversible structural changes such as annihilation of cell-cell-contacts. We speculate that the higher toxicity of CTAB-coated spherical particles as compared to rod-shaped ones is either due to cluster formation and – as a consequence – subsequent release of CTAB into the cell interior or can be attributed to curvature sensing and subsequent signalling. The former pathway would describe a more secondary toxicity of the GNPs as the particle itself is not toxic but the aggregation induced desorption of the surface ligands emphasizing the synergistic effect of shape and surface functionalization.
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Acknowledgments For the fruitful collaboration in the cell laboratories of Mainz and Göttingen, we would like to thank Stefanie Klassen, Anja Herdlitschke, Angela Rübeling and Dr. Ingrid Schuberth. Declaration of interest: Financial support was granted by the German Science Foundation (DFG) through the SPP 1313 Bio-Nano-Responses program (JA 963/10-1). The authors report no conflicts of interest. The authors alone are responsible for the content and writing of the paper. References Alkilany AM, Nagaria PK, Hexel CR, Shaw TJ, Murphy CJ, Wyatt MD. 2009. Cellular uptake and cytotoxicity of gold nanorods: Molecular origin of cytotoxicity and surface effects. Small 5:701–708. Arnida, Malugin A, Ghandehari H. 2010. Cellular uptake and toxicity of gold nanoparticles in prostate cancer cells: A comparative study of rods and spheres. J Appl Toxicol 30:212–217. Barz M, Tarantola M, Fischer K, Schmidt M, Luxenhofer R, Janshoff A, Theato P, Zentel R. 2008. From defined reactive Diblock copolymers to functional HPMA-based self-assembled nanoaggregates. Biomacromolecules 9:3114–3118. Basu S, Harfouche R, Soni S, Chimote G, Mashelkar RA, Sengupta S. 2009. Nanoparticle-mediated targeting of MAPK signaling predisposes tumor to chemotherapy. PNAS 106:7957–7961. Cans AS, Hook F, Shupliakov O, Ewing AG, Eriksson PS, Brodin L, Orwar O. 2001. Measurement of the dynamics of exocytosis and vesicle retrieval at cell populations using a quartz crystal microbalance. Anal Chem 73:5805–5811. Cedervall T, Lynch I, Foy M, Berggad T, Donnelly SC, Cagney G, Linse S, Dawson KA. 2007. Detailed identification of plasma proteins adsorbed on copolymer nanoparticles. Angewandte Chemie-Int Ed 46:5754–5756. Chanana M, Gliozzi A, Diaspro A, Chodnevskaja I, Huewel S, Moskalenko V, Ulrichs K, Galla HJ, Krol S. 2005. Interaction of polyelectrolytes and their composites with living cells. Nano Lett 5:2605–2612. Chithrani BD, Chan WCW. 2007. Elucidating the mechanism of cellular uptake and removal of protein-coated gold nanoparticles of different sizes and shapes. Nano Lett 7:1542–1550. Chithrani BD, Ghazani AA, Chan WCW. 2006. Determining the size and shape dependence of gold nanoparticle uptake into mammalian cells. Nano Lett 6:662–668. Connor EE, Mwamuka J, Gole A, Murphy CJ, Wyatt MD. 2005. Gold nanoparticles are taken up by human cells but do not cause acute cytotoxicity. Small 1:325–327. Cooper MA, Singleton VT. 2007. A survey of the 2001 to 2005 quartz crystal microbalance biosensor literature: Applications of acoustic physics to the analysis of biomolecular interactions. J Mol Recognit 20:154–184. de la Fuente JM, Berry CC, Riehle MO, Curtis ASG. 2006. Nanoparticle targeting at cells. Langmuir 22:3286–3293. Ferrari M. 2008. Beyond drug delivery. Nat Nanotechnol 3:131–132. Giaever I, Keese CR. 1986. Use of electric-fields to monitor the dynamic aspect of cell behavior in tissue-culture. IEEE T Bio-med Eng 33:242–247. Giaever I, Keese CR. 1993. A morphological biosensor for mammalian-cells. Nature 366:591–592.
268
M. Tarantola et al.
Grenha A, Grainger CI, Dailey LA, Seijo B, Martin GP, Remunan-Lopez C, Forbes B. 2007. Chitosan nanoparticles are compatible with respiratory epithelial cells in vitro. Eur J Pharm Sci 31:73–84. Hanaichi T, Sato T, Iwamoto T, Malavasiyamashiro J, Hoshino M, Mizuno N. 1986. A stable lead by modification of Sato method. J Electron Microsc 35:304–306. Hanauer M, Pierrat S, Zins I, Lotz A, Soennichsen C. 2007. Separation of nanoparticles by gel electrophoresis according to shape and size. Nano Lett 7:2881–2885. Hayat MA. 2000. Principles and techniques of electron microscopy: Biological application 4th ed. Cambridge: Cambridge University Press. Hug TS. 2003. Biophysical methods for monitoring cell-substrate interactions in drug discovery. Assay Drug Dev Technol 1:479–488. Jana NR, Gearheart L, Murphy CJ. 2001. Seed-mediated growth approach for shape-controlled synthesis of spheroidal and rodlike gold nanoparticles using a surfactant template. Adv Mater 13:1389–1393. Jiang W, Kim BYS, Rutka JT, Chan WCW. 2008. Nanoparticlemediated cellular response is size-dependent. Nat Nanotechnol 3:145–150. Khan JA, Pillai B, Das TK, Singh Y, Maiti S. 2007. Molecular effects of uptake of gold nanoparticles in HeLa cells. Chembiochem 8:1237–1240. Kumar CSSR. 2006. Nanomaterials for cancer diagnosis. Weinheim, Germany: Wiley-VHC. Lewinski N, Colvin V, Drezek R. 2008. Cytotoxicity of nanoparticles. Small 4:26–49. Lo CM, Keese CR, Giaever I. 1993. Monitoring motion of confluent cells in tissue-culture. Exp Cell Res 204:102–109. Lundqvist M, Stigler J, Elia G, Lynch I, Cedervall T, Dawson KA. 2008. Nanoparticle size and surface properties determine the protein corona with possible implications for biological impacts. PNAS 105:14265–14270. Lynch I, Cedervall T, Lundqvist M, Cabaleiro-Lago C, Linse S, Dawson KA. 2007. The nanoparticle – protein complex as a biological entity; a complex fluids and surface science challenge for the 21st century. Adv Colloid Interface Sci 134–35:167–174. Maynard AD, Aitken RJ, Butz T, Colvin V, Donaldson K, Oberdörster G, Philbert MA, Ryan J, Seaton A, Stone V, Tinkle SS, Tran L, Walker NJ, Warheit DB. 2006. Safe handling of nanotechnology. Nature 444:267–269. Moyes SM, Smyth SH, Shipman A, Long S, Morris JF, Carr KE. 2007. Parameters influencing intestinal epithelial permeability and microparticle uptake in vitro. Int J Pharm 337:133–141. Nativo P, Prior IA, Brust M. 2008. Uptake and intracellular fate of surface-modified gold nanoparticles. Acs Nano 2:1639–1644. Nikoobakht B, El-Sayed MA. 2003. Preparation and growth mechanism of gold nanorods (NRs) using seed-mediated growth method. Chem.Mat 15:1957–1962. Oberdörster G, Stone V, Donaldson K. 2007. Toxicology of nanoparticles: A historical perspective. Nanotoxicology 1:2–25. Pan Y, Neuss S, Leifert A, Fischler M, Wen F, Simon U, Schmid G, Brandau W, Jahnen-Dechent W. 2007. Size-dependent cytotoxicity of gold nanoparticles. Small 3:1941–1949. Pax M, Rieger J, Eibl RH, Thielemann C, Johannsmann D. 2005. Measurements of fast fluctuations of viscoelastic properties with the quartz crystal microbalance. Analyst 130:1474–1477.
Supplementary material available online Supplementary Figures 1–3.
Prescott SW, Mulvaney P. 2006. Gold nanorod extinction spectra. J Appl Phys 99:123504. Prescott SW, Mulvaney P. 2008. Erratum: Gold nanorod extinction spectra. J App Phys 103:119901. Rayavarapu RG, Petersen W, Hartsuiker L, Chin P, Janssen H, van Leeuwen FWB, Otto C, Manohar S, van Leeuwen TG. 2010. In vitro toxicity studies of polymer-coated gold nanorods. Nanotechnology 21:145101. Reiss B, Janshoff A, Steinem C, Seebach J, Wegener J. 2003. Adhesion kinetics of functionalized vesicles and mammalian cells: A comparative study. Langmuir 19:1816–1823. Rothen-Rutishauser B, Grass RN, Blank F, Limbach LK, Muehlfeld C, Brandenberger C, Raemy DO, Gehr P, Stark WJ. 2009. Direct combination of nanoparticle fabrication and exposure to lung cell cultures in a closed setup as a method to simulate accidental nanoparticle exposure of humans. Environ Sci Technol 43:2634–2640. Sapper A, Wegener J, Janshoff A. 2006. Cell motility probed by noise analysis of thickness shear mode resonators. Anal Chem 78:5184–5191. Shukla R, Bansal V, Chaudhary M, Basu A, Bhonde RR, Sastry M. 2005. Biocompatibility of gold nanoparticles and their endocytotic fate inside the cellular compartment: A microscopic overview. Langmuir 21:10644–10654. Shukoor MI, Natalio F, Tahir MN, Wiens M, Tarantola M, Therese HA, Barz M, Weber S, Terekhov M, Schroder HC, Muller WEG, Janshoff A, Theato P, Zentel R, Schreiber LM, Tremel W. 2009. Pathogen-mimicking MnO nanoparticles for selective activation of the TLR9 pathway and imaging of cancer cells. Adv Funct Mater 19:3717–3725. Takahashi H, Niidome Y, Niidome T, Kaneko K, Kawasaki H, Yamada S. 2006. Modification of gold nanorods using phospatidylcholine to reduce cytotoxicity. Langmuir 22:2–5. Tarantola M, Marel AK, Sunnick E, Adam H, Wegener J, Janshoff A. 2010. Dynamics of human cancer cell lines monitored by electrical and acoustic fluctuation analysis. Integr Biol 2:139–150. Tarantola M, Schneider D, Sunnick E, Adam H, Pierrat S, Rosman C, Breus V, Sonnichsen C, Basche T, Wegener J, Janshoff A. 2009. Cytotoxicity of metal and semiconductor nanoparticles indicated by cellular micromotility. Acs Nano 3:213–222. Teeguarden JG, Hinderliter PM, Orr G, Thrall BD, Pounds JG. 2006. Particokinetics in vitro: Dosimetry considerations for in vitro nanoparticle toxicity assessments. Toxicol Sci 95: 300–312. Wang SG, Lu WT, Tovmachenko O, Rai US, Yu HT, Ray PC. 2008. Challenge in understanding size and shape dependent toxicity of gold nanomaterials in human skin keratinocytes. Chem Phys Lett 463:145–149. Wegener J, Janshoff A, Steinem C. 2001. The quartz crystal microbalance as a novel means to study cell-substrate interactions in situ. Cell Biochem Biophys 34:121–151. Wegener J, Sieber M, Galla HJ. 1996. Impedance analysis of epithelial and endothelial cell monolayers cultured on gold surfaces. J Biochem Biophys Methods 32: 151–170. Yen HJ, Hsu SH, Tsai CL. 2009. Cytotoxicity and immunological response of gold and silver nanoparticles of different sizes. Small 5:1553–1561.
