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"content": "Extract sample context and measurement list. <doc> Contents lists available at ScienceDirect jou rn al h om epage: www.elsevier.com/locate/toxlet a College of Pharmacy, Dongguk University, Goyang, Gyeonggi-do 410-820, Republic of Korea b College of Pharmacy, Dongduk Woman's University, Seoul 136-714, Republic of Korea c Department of Chemical Engineering, Kwangwoon University, Seoul 139-701, Republic of Korea d Department of Bionano Technology, Gachon Bionano Research Institute, Gachon University, Sungnam, Gyeonggi-do 461-701, Republic of Korea Article history: Received 22 April 2013 Received in revised form 7 January 2014 Accepted 10 January 2014 Available online 22 January 2014 Keywords: Silver nanoparticles Cytotoxicity Lactate dehydrogenase leakage assay Protein adsorption Reactive oxygen species A growing number of studies report that conventional cytotoxicity assays are incompatible with certain nanoparticles (NPs) due to artifacts caused by the distinctive characteristics of NPs. Lactate dehydrogenase (LDH) leakage assays have inadequately detected cytotoxicity of silver nanoparticles (AgNPs), leading to research into the underlying mechanism. When ECV304 endothelial-like umbilical cells were treated with citrate-capped AgNPs (cAgNPs) or bare AgNPs (bAgNPs), the plasma membrane was disrupted, but the LDH leakage assay failed to detect cytotoxicity, indicating interference with the assay by AgNPs. Both cAgNPs and bAgNPs inactivated LDH directly when treated to cell lysate as expected. AgNPs adsorbed LDH and thus LDH, together with AgNPs, was removed from assay reactants during sample preparation, with a resultant underestimation of LDH leakage from cells. cAgNPs, but not bAg-NPs, generated reactive oxygen species (ROS), which were successfully scavenged by N-acetylcysteine or ascorbic acid. LDH inhibition by cAgNPs could be restored partially by simultaneous treatment with those antioxidants, suggesting the contribution of ROS to LDH inactivation. Additionally, the composition of the protein corona surrounding AgNPs was identified employing liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis. In sum, the LDH leakage assay, a conventional cell viability test method, should be employed with caution when assessing cytotoxicity of AgNPs. 2014 Elsevier Ireland Ltd. All rights reserved. Tel.: +82 31 961 5222; fax: +82 31 961 5206. Nanoparticles (NPs) are drawing attention with the rapid development of nanotechnology. A growing number of engineered NPs are being developed and put to practical use. For industrial and biomedical applications, safety is a primary consideration. As the basic and simplest way to assess safety, cytotoxicity is a standard measure, and cellular viability assays are widely employed for this purpose. Most common cytotoxicity tests include assessments of membrane integrity such as lactate dehydrogenase (LDH) release assay, and measurement of metabolic activity such as the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. Although these methods have been successful in the toxicity screening or evaluation of NPs, recent studies noted particular problems centered around artifacts attributable to the specific characteristics of certain NPs (Love et al., 2012; Monteiro-Riviere et al., 2009; Stone et al., 2009). Cell viability assays rely on chemical reactions involving diverse substance and optical characteristics such as light absorption or fluorescence. Accordingly they may be distorted if NPs affect any aspects of the assay. Possible causes of artifacts ascribable to NPs are broad and include optical properties of NPs that interfere with light absorption or fluorescence, chemical reactions between NPs and reactants involved in assay procedure, and protein adsorption to the NP surface (Holder et al., 2012; Kroll et al., 2011; Monteiro-Riviere et al., 2009). Indeed, most NPs absorb light and interfere with the measurement of optical signals, which is dependent on the particle composition and concentration (Kroll et al., 2011; Monteiro-Riviere et al., 2009). Carbon-based NPs such as single-well carbon nanotubes react with dyes in viability tests and interfere with the chemical reactions required for the assay (Belyanskaya et al., 2007;Casey et al., 2007). Due to the large surface area or surface properties, NPs possess a high adsorptive capacity allowing trapping of molecules that are critical in assay reactions (Worle-Knirsch et al., 2006; Zaqout et al., 2012). A wide range of NPs adsorb proteins and form NP complexes with proteins, the socalled protein corona (Casals et al., 2010; Lundqvist et al., 2008). Other mechanisms may be associated with artifacts in NPs toxicity assays in addition to those described. Accordingly, cytotoxicity tests for NPs need to be performed and interpreted with caution (Monteiro-Riviere et al., 2009; Stone et al., 2009). Silver nanoparticles (AgNPs) are the single most commonly used NPs in consumer products (Stensberg et al., 2011). AgNPs are considered relatively harmless and exhibit useful characteristics such as antibiotic activity. They are becoming increasingly popular for commercial purposes. However, potential toxicity has been described in in vivo as well as in vitro studies, demanding reconsideration of their putative safety (Stensberg et al., 2011; Sung et al., 2009). We recently observedstrikingdiscrepancies betweenthe results of LDH release assay and MTT assay during the study testing the AgNPs cytotoxicity. LDH is a cytosolic enzyme and leaks out of cells when the membrane is damaged. AgNPs obviously deteriorated the membrane integrity in cultured cells, but the LDH leakage assay failed to detect cytotoxicity. This observation led us to postulate that AgNPs may distort the LDH leakage assay. This interpretation agrees with recent findings that AgNPs inhibit LDH activity, although the mechanism remains elusive (Han et al., 2011). Based on this hypothesis, we explored the mechanism underlying interference with LDH assay by AgNPs. Two different AgNPs, bare AgNPs (bAgNPs) and citrate-capped AgNPs (cAgNPs), were used for our study. Protein adsorption and production of reactive oxygen species (ROS) by AgNPs were examined as possible mechanisms related with the interference of LDH assay. In addition, the composition of protein corona formed on AgNPs was analyzed and identified. cAgNPs were kindly provided by ABC Nanotech (Daejeon, Korea). Water soluble tetrazolium salt (WST)-1 was provided by Takara Bio (Shiga, Japan) and calcein/AM was from Invitrogen (Carlsbad, CA). Protease inhibitor cocktail and sequencing grade modified trypsin were purchased from Roche Diagnostics (Indianapolis, IN) and Promega (Madison, WI), respectively. Anti-LDH monoclonal antibody (clone EP1566Y) and horseradish peroxidase-conjugated anti-mouse IgG were supplied by Abcam (Cambridge, MA) and Thermo Scientific (Rockford, IL), respectively. Recombinant human LDH A was purchased from ProSpec (Ness-Ziona, Israel) and carbon black (Corax N330) was from Evonic Carbon Black Korea (Incheon, Korea). The following reagents were purchased from Sigma-Aldrich (St. Louis, MO): bAgNPs, 2',7 -dichlorodihydrofluorescein diacetate (H2DCF-DA), 4',6-diamidino-2 phenylindole (DAPI), sodium pyruvate, NADH, l-ascorbic acid, N-acetyl-l-cysteine (NAC), formic acid, dithiothreitol (DTT), xanthine, and xanthine oxidase. All other chemicals used here were of the highest purity available and purchased from standard suppliers. cAgNPs were nano-sized colloidal silver coated with citrate, which was provided as a 20% (w/v) aqueous solution. According to the manufacturer's information, the amount of capping agent was less than 1% by weight. bAgNPs were silver nanopowder with a particle size of <100 nm. Both cAgNPs and bAgNPs were dispersed in either cell culture medium (Dulbecco's modified Eagle's medium, DMEM) or ECV304 cell lysate. The surface charge was measured using a zeta potential analyzer (ZetaPlus; Brookhaven Instruments; Holtsville, NY) in 100 g/mL dispersions. Size and morphology of AgNPs were analyzed in images of the dispersed particles acquired by a JEM1010 transmission electron microscope (TEM; JEOL, Tokyo, Japan). In addition to TEM analysis, particle sizes and their changes over time were also analyzed by measuring electrophoretic light scattering with an ELS-Z particle size analyzer (Otsuka Electronics, Osaka, Japan). Silver ion concentration in AgNP dispersions was measured with an Elan 6100 inductively coupled plasma–mass spectrometry (ICP–MS; PerkinElmer, Waltham, MA) in the supernatant after centrifugation at 120,000g for 6 h using an Optima L-100XP ultracentrifuge with TY90Ti rotor (BeckmanCoulter,Indianapolis,IN) as previously described (Hagendorfer et al., 2012). ECV304 human endothelial-like umbilical cell line and HeLa human epithelial cell line were obtained from the American Type Culture Collection (Manassas, VA). Both cells were grown in DMEM supplemented with 10% fetal bovine serum, 100 U/ml penicillin, and 100 g/ml streptomycin (Invitrogen). Cells were maintained at 37 ◦C and 5% CO2 in a humidified incubator and were subcultured when they reached 80–90% confluence. Cytotoxicity of AgNPs was examined by LDH leakage and formazan formation assay as previously described (Oh et al., 2013). ECV304 cells were seeded in 96 well plates at a density of 1 × 104 cells/well and were grown for 24 h. Cells were treated with AgNPs by substituting culture media with 200 -L DMEM containing dispersed AgNPs and incubated for 24 h. For the LDH leakage assay, cell culture media were transferred to 1.5 mL microtubes and centrifuged at 16,000g for 20 min to remove cell debris and AgNPs. Twenty microlitres of supernatant was added to a 200--L aliquot of working reagent containing 0.2 mM NADH and 2.5 mM sodium pyruvate in wells of 96-well plates. The decrease in absorbance at 340 nm was measured for 3 min with a SpectraMax M3 microplate reader (Molecular Devices, Sunnyvale, CA). Relative LDH activity was calculated from the slope of decreasing absorbance. LDH activities measured with the media from the untreated cells and the cells treated with 1% Triton X-100 were regarded as 100% and 0% viability, respectively. Assays were conducted in duplicate for each sample. The viability of cells treated with AgNPs was expressed as a percentage of that of untreated cells. In the formazan formation assay, cell viability was determined using the Premix WST-1 Cell Proliferation Assay System (Takara Bio). Cells were prepared and treated with AgNPs as described in LDH leakage assay except that phenol red-free DMEM was used instead of normal, phenol red-containing medium.Atthe end oftreatment, Premix WST-1 was added to each well and incubated for additional 4 h. Media were collected and centrifuged at 16,000g for 20 min. Cell viability was determined by measuring the absorbance at 440 nm with a reference wavelength of 630 nm using the SpectraMax M3 microplate reader. Alternatively, cell membrane integrity was examined with digital imaging employing a membrane-permeable cytosolic dye, calcein-acetoxymethyl ester and a membrane-impermeable nucleus stain, DAPI. After treatment of AgNPs for 24 h, cells were stained with 1 -M calcein and 1 -M DAPI for 20 min. Stained cells were imaged with an Eclipse Ti-U inverted microscope equipped with a S Fluor 20X objective lens (Nikon, Tokyo, Japan) and an Evolve 512 electron-multiplying chargecoupled device camera (Photometrics, Tucson, AZ). Illumination was provided by a Sutter DG-4 filter changer (Sutter Instruments, Novato, CA). Excitation and emission wavelengths used for calcein were 480 and 530 nm and those for DAPI were 380 and 520 nm, respectively. Images were acquired and analyzed with a Meta Imaging System (Molecular Devices). Cells were grown in 100-mm-diameter dishes to approximately 90% confluence. Cells were lysed by incubation in 5 mL of phenol red-free DMEM containing 1% Triton X-100 and protease inhibitor cocktail for 4 h on ice. Lysed cells were transferred to 1.5-mL microtubes and centrifuged at 16,000g for 20 min. The supernatant was collected and pooled as a cell lysate. Protein content was quantified with a bicinchoninic acid protein assay kit (Thermo Scientific). Cell lysate was treated with NPs at 37 ◦C for the indicated time with mild agitation. The lysate was centrifuged at 16,000g for 20 min to sediment AgNPs. The supernatant was used for LDH activity assay and immunoblotting. For the analysis of absorbed proteins on AgNPs, the sediment was washed twice with phosphate-buffered saline by repeating resuspension and centrifugation. The resultant sediment was suspended in sample buffer solution (5% -mercaptoethanol, 5% sodium dodecyl sulfate (SDS), 25% glycerol, 0.01% bromophenol blue, 0.32 M Tris-HCl; pH 6.8) and was boiled for 10 min. Following centrifugation, the supernatant was obtained and subjected to protein analysis as described below. LDH activity was examined with the same methods described in Section 2.4. In recombinant LDH experiments, 0.01% bovine serum albumin was added to reaction mixture to disperse AgNPs and to improve LDH stability. The supernatant obtained as described above was analyzed by conventional SDS–polyacrylamide gel electrophoresis (SDS–PAGE) (Lee et al., 2009). Concentrations of polyacrylamide were 4% and 15% for stacking and resolving gels, respectively. The gel-separated proteins were transferred to a polyvinylidene difluoride membrane by applying 100V for 90 min. LDH was probed with an anti-LDH primary antibody (1:2000) and a horseradish peroxidase-conjugated anti-mouse IgG secondary antibody (1:5000). Following the application of Immobilon Western detection reagents (Millipore, Billerica, MA), chemiluminescence images were obtained and analyzed with a Molecular Imager ChemiDoc XRS+ imaging system (Bio-Rad Laboratories, Hercules, CA). Cell lysates were treated with 1 -M H2DCF-DA and incubated for 2 h in the dark to cleave the diacetate group. The lysates were treated with AgNPs with or without antioxidants such as NAC and ascorbic acid, and incubated for 2 h. AgNPs were removed by centrifugation and the fluorescence intensity of the supernatant was measured at 480/535 nm excitation/emission wavelengths with a SpectraMax M3 microplate reader. The 0.5 mM xanthine/10 mU/ml xanthine oxidase free radical generation system was used as the positive control. Proteins were separated by SDS–PAGE and were excised from the gel. Gel pieces containing proteins were dehydrated in acetonitrile (ACN) and vacuum-dried for 20 min with a SpeedVac concentrator (Thermo Scientific). Proteins in gel pieces were reduced by incubating in 50 mM NH4HCO3 solution containing 10 mM DTT for 45 min at 56 ◦C. Cysteine residues were alkylated in 55 mM iodoacetamidecontaining 50 mM NH4HCO3 solution for 30 min. Each gel piece was digested with 12.5 ng/-L sequencing grade modified trypsin (Promega)in 50 mM NH4HCO3 buffer solution (pH 7.8) at 37 ◦C overnight. Tryptic peptides were extracted with 5% formic acid in 50% ACN solution for 20 min at room temperature. Supernatants were collected and dried with a SpeedVac concentrator. Following resuspension in 0.1% formic acid, proteins were concentrated using C18 ZipTips (Millipore). Tryptic peptides were loaded onto a fused silica microcapillary column (12 cm × 75 m) packed with C18 reversed phase resin (5 m, 200A). ̊ The LC eluents were: A, distilled water (DW) containing 0.1% formic acid and B, ACN containing 0.1% formic acid. Starting from an A:B composition of 70:30, the linear gradient reached the 100% of B concentration in 60 min at a flow rate of 0.25 -L/min. The column was directly connected to Finnigan LTQ linear ion-trap mass spectrometer (Thermo Scientific) equipped with a nano-electrospray ion source. The electrospray voltage was set at 1.95 kV and the threshold for switching from MS to MS/MS was 500. The normalized collision energy for MS/MS was 35% of main radio frequency amplitude and the duration of activation was 30 msec. All spectra were acquired in data-dependent scan mode. Each full MS scan was followed by five MS/MS scan corresponding from the most intense to the fifth intense peaks of full MS scan. Repeat count of peak for dynamic exclusion was 1 and repeat duration was 30 sec. The dynamic exclusion duration was set for 180 sec and width of exclusion mass was ± 1.5 Da. The list size of dynamic exclusion was 50. Acquired LC-ESI-MS/MS fragment spectra were searched in the BioWorks Browser (version Rev. 3.3.1 SP1, Thermo Scientific) with the SEQUEST search engines against National Center for Biotechnology Information (www.ncbi.nlm.nih.gov) non-redundant human database. The searching conditions were trypsin enzyme specificity, a permissible level for two missed cleavages, peptide tolerance; ±2 amu, a mass error of ±1 amu on fragment ions and fixed modifications of carbamidomethylation of cysteine (+57 Da) and oxidation of methionine (+16 Da) residues. ClueGO, a Cytoscape plugin, was used for data visualization from the analysis of protein ontology and the biological processes. Gene ontology categories were used to capture biological information and kappa statistics was applied to the creation of networks for protein interactions. The networks of proteins adsorbed to AgNPs were visualized using the GOlorize. Each biological process was represented with the nodes connected with edges to indicate interactions. The mean and standard error (SE) of the mean were calculated for all experimental groups. The data were analyzed using one-way analysis of variance followed by Dunn's test to determine significant differences from the control. Statistical analysis was performed using SigmaStat software version 3.5 (Systat Software, San Jose, CA). Null hypotheses of no difference were rejected if P values were < 0.05. Information regarding NPs used in this study is summarized in Table 1. AgNPs were dispersed in either DMEM or ECV304 cell lysate, and the morphology and size distribution were examined by TEM. Most of the particles appeared globular and exhibited a typical particle size distribution (Fig. 1A and B). Primary sizes of AgNPs were 43 ± 1 and 50 ± 1 nm in DMEM and cell lysate, respectively. Those of bAgNPs were 82 ± 4 and 72 ± 3 nm in the same respective media. The hydrodynamic sizes measured by DLS were larger than primary sizes, indicating that both AgNPs form aggregates or agglomerates in DMEM and cell lysates (Table 1). Indeed, agglomerates were observed in TEM analysis as shown in Fig. 1A, and were more frequent in bAgNPs (right panel). AgNPs maintained their initial size without significant change for at least 24 h in DMEM and cell lysate (Fig. 1C). Silver ion concentration in AgNPs dispersion measured by ICP–MS in 100 g/mL AgNPs dispersion was <0.7 g/mL regardless of the dispersion medium and did not increase significantly up to 24 h (Supplemental Fig. 1A). ECV304 cells were incubated with the indicated concentrations of AgNPs for 24 h and the resultant cytotoxicity was assessed with LDH leakage assay and formazan formation assay using WST-1. The latter assay revealed that the treatment of cAgNPs resulted in decreased viability that was concentration-dependent in the range of 0.1–10 g/mL (Fig. 2A left panel). However, the LDH assay failed to detect any decrease in viability. Comparable results were obtained with bAgNPs. Cytotoxicity of bAgNPs was marginal in the LDH assay, while viability was significantly decreased in the WST-1 assay (Fig. 2A right panel). Discrepancies between the LDH assay and WST-1 assay may occur if AgNPs deteriorate cellular metabolic systems such thatformazan is notformed butthe plasma membrane remains undamaged. Therefore, membrane integrity was investigated to confirm whether AgNPs injured cellular membranes. Cells were incubated with 3 g/mL cAgNPs or 50 g/mL bAg-NPs for 24 h to induce intermediate cytotoxicity. Treated cells were All values are expressed as the mean ± SE. a Hydrodynamic size was measured using electrophoretic light scattering. b Data were cited from previous reports (Kim et al., 2012a, b). stained with 1 -M calcein and 1 -M DAPI for 20 min. Acetoxyester form of calcein can penetrate plasma membrane and is cleaved to release calcein, which is retained in cytosol. This stains cells with intact membranes. DAPI is a nucleic acid stain that is membrane permeable in principle. However, it passes through the membrane less efficiently in live cells and thus a low concentration of DAPI hardly stains the nucleus (Zink et al., 2003).Accordingly, a relatively low concentration of DAPI(1 -M) stains only the nuclei of cells with damaged membrane. As expected, all cells were stained with either calcein or DAPI, and the distributions of calcein- or DAPI-stained cells were mutually exclusive (Fig. 2B). DAPI-stained but calceinnegative cells were observed in both cAgNPs- and bAgNPs-treated cell populations, indicating plasma membrane rupture by AgNPs. The percentage of calcein-positive cells was 75 ± 13% (n = 6) and 71 ± 14% (n = 4) of all cells (DAPI-positive + calcein-positive cells) for cAgNPs and bAgNPs, respectively. These values appeared to be slightly higher than viabilities assessed by theWST-1 assay (41 ± 8% for cAgNPs and 59 ± 7% for bAgNPs), suggesting the presence of cells with intact membranes and damaged metabolic activity. Only calcein stained untreated control cells. These results clearly indicated that AgNPs induced membrane disruption and presumably the leakage of cytosolic components including LDH. Summarizing, the conventional LDH assay underestimated or failed to detect the activity of LDH released from cells to culture media and thus could not properly reflect cytotoxicity related to AgNPs. The effect of AgNPs treatment on LDH activity was examined assuming a potential inhibition of LDH or an interference with the LDH assay. Cell lysate rather than purified LDH was used as a source for LDH, because it may better mimic the assay sample of culture medium containing material released from ruptured cell than purified LDH. Lysate was prepared from ECV304 cells. Protein concentration of lysate was 0.2607 ± 0.0278 mg/mL (n = 14). Lysate was treated with 100 g/mL AgNPs, carbon black or their vehicles. After 0.17 (10 min), 1, 2, 4, 8, 12, or 24 h, NPs were removed by centrifugation and LDH activity was measured in the supernatant. cAgNPs inhibited LDH nearly completely from 0.17 h after treatment (Fig. 3A). bAgNPs also inhibited LDH significantly, but the potency was weaker than those of cAgNPs. Carbon black, a pure carbon NPs used as a control, did not change LDH activity. Triton-X100, a detergent used for lysate preparation, was confirmed notto affect LDH activity at 1% concentration (data not shown). Untreated cell lysate maintained its LDH activity during test, at least up to 24 h. DW, a vehicle for AgNPs and suspension buffer solution (134 mM NaCl, 12.0 mM NaHCO3, 2.9 mM KCl, 0.34 mM Na2HPO4, 1 mM MgCl2, 10 mM HEPES, 5 mM glucose, 1 mM CaCl2, 0.3% bovine serum albumin; pH 7.4), a vehicle for carbon black did not affect LDH activity (Fig. 3A; Kim et al., 2012). Lower concentrations of AgNPs were tested to figure out the concentration and treatment time dependency of LDH inhibition (Fig. 3B). cAgNPs inhibited LDH in a concentration-dependent manner and such inhibition was statistically significant over 1 g/mL (Fig. 3B left panel). Similar inhibition was observed with 50 and 100 g/mL bAgNPs, but the potency was lower than that of cAgNPs (Fig. 3B right panel). Comparable results could be obtained with the lysate of primary cultured vascular smooth muscle cells in addition to ECV304 (data not shown). LDH adsorption by AgNPs was investigated as a possible mechanism for the decrease in LDH activity in the assay (Wigginton et al., 2010). Cell lysate was incubated with indicated concentrations of AgNPs for 24 h, or with 10 g/mL cAgNPs or 100 g/mL bAgNPs for indicated times. Following centrifugation, both supernatants and AgNP sediments were subjected to Western blot analysis. Supernatant from lysate treated with cAgNPs contained less LDH and such reduction was well correlated with the concentration and the treatment time of cAgNPs (Fig. 4A upper panels and 4B). Instead, LDH was detected in cAgNPs sediment, which was also proportional to the concentration and treatment time. These results suggest the adsorption of LDH by cAgNPs. In case of bAgNPs, a concentration- or treatmenttime-dependentincrease was observed in NPs sediment, but decrease in supernatant was clear only in 100 g/mL, the highest concentration tested (Fig. 4A lower panels and 4C). In addition, -actin was tested for comparison. Similar to LDH,the density of actin was correlated with the quantity of AgNPs in AgNPs sediment, indicating that -actin as well as LDH adsorbed onto AgNPs. ROS generation by AgNPs was examined in cell lysates to assess the involvement of ROS in LDH inhibition by AgNPs. When cAgNPs were used to treat lysates containing DCF, a substantial amount of ROS was produced, which was also concentration-dependent and statistically significant over 1 g/mL. However, bAgNPs failed to generate ROS up to 100 g/mL (Fig. 5A upper panel). ROS generated by cAgNPs were successfully scavenged by simultaneous treatment with antioxidants including NAC and ascorbic acid, and the scavenging effect was correlated with antioxidant concentration (Fig. 5A lower panel). NAC or ascorbic acid alone did not induce ROS production, and the positive control, xanthine/xanthine oxidase system regenerated considerable amounts of ROS. The effect of antioxidants on LDH inhibition was tested to confirm the contribution of ROS to LDH inhibition by cAgNPs. Concomitant treatment of 1 mM NAC or ascorbic acid blocked LDH inhibition by cAgNPs significantly, although it was a partial and not complete restoration (Fig. 5B). NAC or ascorbic acid alone did not affect LDH activity. This result suggested that a reduction in LDH activities by cAgNPs could be attributed to ROS production in addition to the removal of LDH by adsorption. Cytotoxicity, LDH activity inhibition and ROS generation by AgNO3 were examined to test whether silver ions released from AgNPs might contribute to the effect of AgNPs that had been observed. ECV304 cells were treated with AgNO3 for 24 h and cytotoxicity was tested by WST-1 assay. AgNO3 was toxic to cell at concentration exceeding 100 -M but did not induce significant cytotoxicity at concentrations up to 30 -M (Supplemental Fig. 1B). Cell lysate was treated with AgNO3 for 24 h and LDH activity was assayed to test the effect on LDH activity. AgNO3 failed to inhibit LDH up to 30 -M although it was effective at 100 -M (Supplemental Fig. 1C). ROS generation by AgNO3 was tested under the same experimental condition as described in Section 3.4. However, it failed to produce ROS up to 100 -M (Supplemental Fig. 1D). Considering the silver ion concentration in AgNP dispersion was <0.7 g/mL (- 6.5 -M), these results indicated that although silver ions were eluted from AgNPs,they did not contribute to LDH inhibition and were not related with the cytotoxicity and ROS generation by AgNPs (Supplemental Fig. 1). To confirm direct effect of AgNPs on LDH, experiments were performed with recombinant LDH. Recombinant LDH was incubated with AgNPs for 24 h and the activity was assayed. Similar to the previous results (Fig. 3), both cAgNPs and bAgNPs inhibited recombinant LDH in a concentration-dependent manner (Fig. 6A). Furthermore, such inhibition by cAgNPs was restored by simultaneous treatment of 1 mM NAC or 1 mM AA, indicating the involvement of ROS generation in LDH inhibition (Fig. 6B). These results are consistent with the results obtained from cell lysate experiments (Figs. 3 and 5) and suggest the presence of direct interaction between AgNPs and LDH. The amount of protein adsorbed by AgNPs was estimated by analyzing the decrease in total protein content in cell lysate after incubation with AgNPs. Following 24 h incubation in cell lysate, AgNPs were removed by centrifugation, and protein was quantified in supernatant. Protein concentration was significantly reduced by incubation with cAgNPs and the decrease was concentrationdependent within the range of 0.1–10 g/mL (Fig. 7 left panel). Similar results were seen with bAgNPs at concentrations of 10–100 g/mL, but the decrease in protein concentration was less than that by cAgNPs (Fig. 7 right panel). Protein concentration in 10 g/mL cAgNPs-treated lysate was 53 ± 5% of control and that in 100 g/mL bAgNPs-treated lysate was 64 ± 5% of control. This clearly demonstrates protein adsorption and removal of proteins from cell lysate by AgNPs. Proteins adsorbed by AgNPs were analyzed by LC-MS/MS to identify the composition of the protein corona. AgNP containing sediments were obtained after incubating in lysate, and proteins were digested and subjected to analysis. The identified proteins are listed in Supplemental Table 1. Subunits of LDH were identified in protein coronas of both cAgNPs and bAgNPs, and were ranked 20th, 32nd, and 36th(cAgNPs) and 61st and 159th(bAgNPs)(Supplemental Table 1). More different kinds of proteins could be detected in the cAgNP corona. ClueGo analysis indicated that major proteins adsorbed to cAgNPs included proteins involved in intracellular transport,mitotic cell cycle,RNAsplicing, glycolysis, ribonucleotide metabolic process, and ubiquitin-protein ligase regulation (Supplemental Fig. 2). The protein corona surrounding bAgNPs included proteins related with nucleocytoplasmic transport, intracellular transport, mRNA processing, protein complex assembly, posttranscriptional regulation, and proteasomal ubiquitin-dependent protein catabolic process. Various NPs interfere with conventional cell viability assays leading to erroneous interpretation (Love et al., 2012; Monteiro-Riviere et al., 2009; Stone et al., 2009). In this study, LDH leakage assay failed to reflect the cytotoxicity of AgNPs, so the underlying mechanism was investigated. There are three main findings of our study. (1) Both cAgNPs and bAgNPs interfere with LDH leakage assay. (2) AgNPs adsorb LDH and LDH, together with AgNPs, can be removed from an assay sample during sample preparation, with resultant underestimation of LDH leakage from cells. cAgNPs, not bAgNPs, produce ROS, which leads to LDH inactivation. (3) In addition to LDH, wide ranges of cellular proteins can adhere to AgNPs and make the protein corona. Therefore, the LDH leakage assay, a conventional cell viability test, should be employed withcautionfor the assessment of NPs, especially ofAgNPs. Since the LDH assay utilizes light absorption, experimental processes to remove NPs from assay sample are necessary because the presence of NPs physically interferes with the optical signal. Therefore, NPs with a potential to adsorb proteins are likely to cause artifacts by eliminating LDH from the assay sample. Since NPs tend to adsorb protein, attention needs to be paid to AgNPs and, indeed, most NPs in LDH assay. Metal NPs are likely to release ions in dispersion and AgNPs also elute silver ions in DMEM and cell lysate (Supplemental Fig. 1A). Silver ions are cytotoxic (de Lima et al., 2012; Foldbjerg et al., 2009) and have a potential to inhibit LDH (Menon and Wright, 1989; Schmitt and Ritter, 1975). They also showed cytotoxicity and inhibited LDH activity in our experimental condition, but the concentration of silver ions eluted from AgNPs was not high enough (Supplemental Fig. 1A–C). Silver ions are capable of producing ROS, which mainly contributes to cytotoxicity (de Lima et al., 2012; Foldbjerg et al., 2009). However, they currently failed to generate ROS when present at a substantially high concentration, 100 -M, in cell lysates (Supplemental Fig. 1D), suggesting that they may not induce ROS in a cell-free system. This result agrees with the ineffectiveness of bAgNPs in ROS generation and its inhibition of LDH (Fig. 5). Both cAgNPs and bAgNPs release silver ions, and therefore bAgNPs must have produced ROS and inhibited LDH as did cAgNPs if silver ions are responsible for ROS generation. All the results consistently indicate that silver ions originating from AgNPs are not related with cytotoxicity, LDH inhibition, and ROS generation mediated by AgNPs. In addition to silver ions, capping material and its stability may be other factors affecting biological activity of NPs. Although little information was available regarding capping stability in this study, the citrate content was less than 1% of AgNPs according to manufacturer. Assuming that 100% of citrate is released from cAgNPs,the maximal concentration of citrate reaches 1 g/mL in a 100-g/mL cAgNP dispersion, which is equal to 5.3 -M (molecular weight of citrate = 189). Citrate did not induce cytotoxicity or inhibit LDH at concentrations up to 10 -M in the experiment performed with citric acid (data not shown). Therefore, at least the capping agent citrate is not involved in cytotoxicity or LDH inhibition caused by cAgNPs, although capping stability was not tested. To avoid interference with optic signal, AgNPs were removed from samples before being assayed by centrifugation at 16,000g for 20 min. This experimental condition cannot remove AgNPs completely. After centrifugation, cAgNPs still remained in supernatant; 3.993 ± 0.119 and 4.388 ± 0.284 g/mL when cell lysate and DMEM containing 100 g/mL cAgNPs were tested, respectively. The concentration of cAgNP was measured by ICP–MS by subtracting silver ion concentration from silver concentration of 16,000g supernatant. In case of bAgNPs, relatively less AgNPs were detected in 16,000g supernatant; 0.211 ± 0.094 and 0.355 ± 0.153 g/mL in cell lysate and DMEM, respectively. If the concentration in supernatant is proportional to the concentration of AgNPs dispersion, it must be less in the lower concentration of dispersed AgNPs. These results indicate that small particles remained in the supernatant even after centrifugation at 16,000g. However, these concentrations minimally affect LDH and MTT assay because light absorption is not significant at these levels. They may also cause artifacts in LDH analysis by Western blot, which may underestimate both LDH disappearance in the supernatant and LDH increase in the AgNP containing sediment. However, these amounts of AgNPs are unlikely to change the general conclusions of our study, although they may not be ignorable in certain experiments. As with other NPs, AgNPs adsorb proteins to form a protein corona in biological fluids, although its composition has not been described extensively (Ashkarran et al., 2012; Wigginton et al., 2010). Currently, LDH adsorbed onto AgNPs, evidenced by Western blot and LC-MS/MS analysis (Fig. 4 and Supplemental Table 1). Han et al. (2011) observed LDH inhibition by AgNPs, although all the types of AgNPs tested did not inactivate LDH and the mechanism was not studied (Han et al., 2011). Indeed, their observation is consistent with our finding and may be attributed to LDH removal from reactants through adsorption by AgNPs. Similar to our finding, interference with LDH assay by LDH adsorption has been reported in the study of titanium dioxide NPs (Zaqout et al., 2012). A number of NPs are capable of adsorbing proteins to forma corona (Lundqvist et al., 2008; Tenzer et al., 2011). LDH may well be presentin the protein corona surrounding NPs, although the composition of protein corona is dependent on physicochemical properties of NPs including surface, size, and concentration, and thus is hard to predict or analyze (Tenzer et al., 2011). The result of the LDH leakage assay requires careful interpretation, especially if NPs have high protein affinity. It is hard to estimate the relative contribution although both LDH adsorption and LDH inactivation by ROS are related to LDH assay interference by cAgNPs. LDH was not removed completely from cell lysate and a substantial amount of LDH remained even after incubation and removal of cAgNPs, although LDH inhibition was nearly complete in the same condition (Figs. 3 and 4). This implies the involvement of mechanisms other than LDH removal through adsorption. Together with adsorption, protein inactivation is the most common type of NPs–protein interaction. Indeed, cAgNPs generated ROS in cell lysate, and its scavenging by an antioxidant restored LDH inhibition (Fig. 5). Additionally, such restoration of LDH inhibition by antioxidants was also confirmed with recombinant LDH (Fig. 6). It is hard to explain the mechanism underlying ROS-mediated inactivation of LDH with current experimental results. However, the inhibition of LDH by ROS has already been described in previous studies (Andersson et al., 2000; Kendig and Tarloff, 2007; Olson and Massey, 1980; Pamp et al., 2005). A monomer of LDH contains six cysteine residues out of a total of 571 amino acids. Thus, there are multiple sites within LDH capable of accepting a reactive oxygen intermediate, which appears to be the reason why LDH are susceptible to ROS. ROS generation is also a common characteristic of a wide range of NPs (Xia et al., 2009). Hence, ROS may be another factor giving false results in an LDH leakage assay for NPs. LDH disappearance from lysates was evident only in cell lysates treated with 100 g/mL bAgNPs and was not evident in lower concentrations (Fig. 4AandC). Our interpretationis thatthe pool of LDH is large in cell lysates and a relatively small portion of LDH adheres to bAgNPs. Therefore, the increment of LDH in bAgNP sediment could be easily detected whereas a small change in a large pool, the supernatant, would barely be evident in Western blot analysis. bAgNPs was 0.0149 (=0.2493–0.2344) g/-L (Fig. 7), equal to 11.53 g protein/g cAgNPs and 1.49 g protein/g bAgNPs. Since the specific gravity of silver is 10.490 g/cm3, the protein adsorption was 120.95 g protein/cm3 cAgNPs and 15.63 g protein/cm3 bAg-NPs, which in turn correspond to 1.008 g protein/m2 (=120.95 g protein/120 m2) and 0.188 g protein/m2 (=15.63 g protein/83 m2), respectively. Therefore, cAgNPs adsorbed approximately 5.4 times more protein than bAgNPs. Overall, the protein binding of bAg-NPs appears to be weaker than that of cAgNPs (Fig. 7), which correlated with the potency of LDH inhibition (Fig. 3). LC-MS/MS analysis also revealed the presence of LDH subunits in protein corona, and LDH was ranked in higher order in cAgNPs list than in bAgNPs list, which was dependent on the quantity of protein detected (Supplemental Table 1). It is unclear why the extent of protein binding differed between cAgNPs and bAg-NPs. Protein binding capacity is generally dependent on surface characteristics such as surface area, charge, and coating material (Soenen et al., 2011). Therefore, the difference of surface area and the surface properties characterized by a coating agent, citrate, may be responsible for the difference between cAgNPs and bAgNPs. Whether mechanisms other than protein adsorption are involved in the interference of LDH assay by bAgNPs is unclear. It is widely accepted that interaction with proteins is a critical factor determining the biological activity of NPs. Understanding the dynamics of NPs–protein corona is essential for predicting the fate, transport, and toxicity of NPs in living systems (Albanese et al., 2012; Tenzer et al., 2011). AgNPs are capable of penetrating biological membrane (Farkas et al., 2011). Currently, LC-MS/MS analysis revealed that diverse proteins formed a protein corona (Supplemental Table 1). Given that AgNPs enter cells, they may affect intracellular protein functions by adsorbing proteins. AgNPs may inactivate proteins or, at least, change protein localization and translocation physically, ultimately altering protein functions. This issue is being studied. Indeed, bAgNPs adsorb heat shock protein 90 (HSP90), a chaperon of endothelial nitric oxide synthase (eNOS) (Supplemental Table 1) and the changes in eNOS activity were observed in bAgNP-treated ECV304 cells (Oh and Lee, unpublished data). Further study will be needed to clarify the effect of protein adsorption by AgNPs on protein and cellular functions. The authors declare that they have no conflicts of interest. This work was supported by the Dongguk University Research Fund of 2011. Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/. 01.015. Worle-Knirsch, J.M., Pulskamp, K., Krug, H.F., 2006. Oops they did it again! Carbon nanotubes hoax scientists in viability assays. Nano. Lett. 6, 1261–1268. </doc> ###",
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"content": "Extract sample context and measurement list. <doc> Linda Böhmert, Matthias Girod, Ulf Hansen, Ronald Maul, Patrick Knappe, Birgit Niemann, Steffen Michael Weidner, Andreas F Thünemann, Alfonso Lampen doi: Orally ingested nanoparticles may overcome the gastrointestinal barrier, reach the circulatory system, be distributed in the organism, and cause adverse health effects. However, ingested nanoparticles have to pass through different physicochemical environments, which may alter their properties before they reach the intestinal cells. In this study, silver nanoparticles are characterized physicochemically during the course of artificial digestion to simulate the biochemical processes occurring during digestion. Their cytotoxicity on intestinal cells was investigated using the Caco-2 cell model. Using field-flow fractionation (A4F) combined with dynamic light scattering (DLS) and small-angle X-ray scattering (SAXS), we found that particles only partially aggregate as a result of the digestive process. Cell viabilities were determined by means of CellTiter-Blue® assay, DAPI-staining and real-time impedance. These measurements reveal small differences between digested and undigested particles (1 to 100 μg/mL or 1 to 69 particles per cell). Our findings suggest that silver nanoparticles may indeed overcome the gastrointestinal juices in their particulate form without forming large quantities of aggregates. Consequently, we presume that the particles can reach the intestinal epithelial cells after ingestion with only a slight reduction in their cytotoxic potential. Our study indicates that it is important to determine the impact of body fluids on the nanoparticles of interest to provide a reliable interpretation of their nanospecific cytotoxicity testing in vivo and in vitro. Informa UK, Ltd. This provisional PDF corresponds to the article as it appeared upon acceptance. Fully formatted PDF and full text (HTML) versions will be made available soon. DISCLAIMER: The ideas and opinions expressed in the journal's Just Accepted articles do not necessarily reflect those of Informa Healthcare (the Publisher), the Editors or the journal. The Publisher does not assume any responsibility for any injury and/or damage to persons or property arising from or related to any use of the material contained in these articles. The reader is advised to check the appropriate medical literature and the product information currently provided by the manufacturer of each drug to be administered to verify the dosages, the method and duration of administration, and contraindications. It is the responsibility of the treating physician or other health care professional, relying on his or her independent experience and knowledge of the patient, to determine drug dosages and the best treatment for the patient. Just Accepted articles have undergone full scientific review but none of the additional editorial preparation, such as copyediting, typesetting, and proofreading, as have articles published in the traditional manner. There may, therefore, be errors in Just Accepted articles that will be corrected in the final print and final online version of the article. Any use of the Just Accepted articles is subject to the express understanding that the papers have not yet gone through the full quality control process prior to publication. Cells Linda Böhmert1,*, Matthias Girod2,*, Ulf Hansen1,2, Ronald Maul2 , Patrick Knappe1,2, Birgit Niemann1 , Steffen M. Weidner2 , Andreas F. Thünemann2 , Alfonso Lampen1 1BfR Federal Institute for Risk Assessment, Max-Dohrn-Str. 8-10, 10589 Berlin, Germany 2BAM Federal Institute for Materials Research and Testing, Unter den Eichen 87, 12205 Berlin, Germany E-mail: ; *Corresponding author: Linda Boehmert, Federal Institute for Risk Assessment, Max-Dohrn-Str. 8-10, 10589 Berlin, Germany. Phone: +49 1888 4123718, FAX: +49 18884123715 Matthias Girod, Federal Institute for Materials Research and Testing, Richard-Willstätter-Str.11, 12489 Berlin, Germany. Phone: +49 30 81045586, Fax: +49 30 81041137 KEYWORDS: Silver nanoparticles, in vitro digestion, field-flow fractionation (FFF), small-angle Xray scattering (SAXS), dynamic light scattering (DLS), Caco-2 cells Orally ingested nanoparticles may overcome the gastrointestinal barrier, reach the circulatory system, be distributed in the organism, and cause adverse health effects. However, ingested nanoparticles have to pass through different physicochemical environments, which may alter their properties before they reach the intestinal cells. In this study, silver nanoparticles are characterized physicochemically during the course of artificial digestion to simulate the biochemical processes occurring during digestion. Their cytotoxicity on intestinal cells was investigated using the Caco-2 cell model. Using field-flow fractionation (A4F) combined with dynamic light scattering (DLS) and small-angle X-ray scattering (SAXS), we found that particles only partially aggregate as a result of the digestive process. Cell viabilities were determined by means of CellTiter-Blue® assay, DAPI-staining and real-time impedance. These measurements reveal small differences between digested and undigested particles (1 to 100 μg/mL or 1 to 69 particles per cell). Our findings suggest that silver nanoparticles may indeed overcome the gastrointestinal juices in their particulate form without forming large quantities of aggregates. Consequently, we presume that the particles can reach the intestinal epithelial cells after ingestion with only a slight reduction in their cytotoxic potential. Our study indicates that it is important to determine the impact of body fluids on the nanoparticles of interest to provide a reliable interpretation of their nanospecific cytotoxicity testing in vivo and in vitro. Due to their anti-microbial properties, silver nanoparticles are one of the most widely used nanoparticles in consumer products, especially in products that are to be used in close contact to the human body, such as textiles, bandages, cosmetics, water purification devices and household items that are in contact with food (Marambio-Jones and Hoek, 2010). Moreover, an application for the use of a nanoscaled silver hydrosol for nutritional purposes has been submitted to the European Union (EFSA 2008). Silver nanoparticles are known to be toxic to many different types of cells (Ahamed et al., 2010; Foldbjerg et al., 3 2009; Gopinath et al., 2010). Furthermore, subtoxic silver concentrations may also have a long-term effect on cells (Kawata et al., 2009; Nowrouzi et al., 2010). In addition, chronic exposure to silver is known to cause argyria in humans (Drake and Hazelwood, 2005; Kim et al., 2009b). Furthermore, silver supplements and silver nanoparticles from food contact materials or contaminated drinking water are frequently ingested orally. Several studies have been performed to investigate specific aspects of the intestinal uptake of silver nanoparticles. For example, the detection of silver in organs after ingesting food with silver nanoparticles (Kim et al., 2009a; Loeschner et al., 2011), the cellular uptake by the intestinal M-cells (Bouwmeester et al., 2011), and the effect of mucus on nanoparticle uptake (Behrens et al., 2002). Chen et al. have demonstrated the significantly size-dependent biodistribution between ionic-, micron- and nanoparticulate metal in in vivo experiments with mice. Nanoparticles ingested in vivo have to pass through different physicochemical environments before they reach the intestinal cells. They are exposed to saliva as well as gastric and intestinal juices before they reach the intestine. During the digestive process, physiological parameters such as pH, ionic strength, as well as protein content and composition may change several times. It is probable that the size, shape, stability, or aggregation state of the nanoparticles is affected and could therefore also substantially influence their cytotoxic potential. To monitor changes in the nanoparticle system, Small-Angle X-ray Scattering (SAXS) is applied to characterize nanoparticle ensembles with respect to their core sizes, shapes and size distributions. SAXS has the advantage of being applicable without extensive sample preparation or even in situ to a particle ensemble; it thus allows measurement times of minutes down to seconds. Complementarily, Dynamic Light Scattering (DLS) measures the hydrodynamic radius; this includes the size of I.) the core, II.) the shell, which consists of a particle stabilizer if present, and III.) a potential protein corona and the hydration shell. Both SAXS and DLS are very sensitive to particle aggregation and they are non-destructive techniques (Thunemann et al., 2009). Furthermore, both are integral methods and simultaneously measure the scattering signal of the entire particle ensemble. Therefore, SAXS and DLS provide good particle statistics by averaging more than 106 particles (Bienert et al., 2009). A major drawback is that interpretation of the data becomes increasingly ambiguous if the sample of interest is highly polydispersed in size, or if the samples contain various types of aggregates (Glatter, 1982). In order to circumvent this problem, asymmetrical flow field-flow fractionation (A4F) can be applied as a particle size separation method before investigation of the fractionated particles with SAXS and DLS (Thünemann et al., 2008). The separation of the A4F is based on the different hydrodynamic diameters and thus on the different diffusion coefficients of the analytes. After injection into a separating channel, particles are affected by a parabolic flow profile of a chosen solvent, i.e. the highest flow rate in the middle of the channel decreasing to the sides. The A4F additionally provides a cross-flow, orthogonal to the solvent flow, which draws the particles to a permeable membrane at the bottom of the separation-channel. The higher their diffusion coefficients are, the more easily they overcome the cross-flow. Thus smaller particles stay in regions of faster solvent flow and elute before larger particles. A4F is an appropriate separation method because it applies much lower shear forces to the samples than, e.g., gel permeation chromatography/size exclusion chromatography (GPC/SEC), and is therefore a suitable method for the investigation of agglomeration/aggregation processes (Thünemann et al., 2008; Knappe et al., 2011). The combination of A4F with SAXS and DLS as analytical tools allow a detailed characterization of the particles during the entire digestion process. The first objective of this study was the detailed characterization of a commercially widespread silver nanoparticle material before and after each step of an in vitro digestion process. To the best of our knowledge, most in vitro studies dealing with the toxicological effects of nanomaterials administered orally use nanoparticles suspended in cell culture media without previous contact to gastrointestinal juices. Therefore, our second objective was a comprehensive cytotoxicological investigation and comparison of digested and undigested particles. Undigested and digested nanoparticles were used for toxicity testing on proliferating and differentiated Caco-2 cells. Caco-2 cells are a standard in vitro model for the intestinal barrier, representing a test system that is widely accepted by pharmacological industries and regulatory authorities because the permeability determined for Caco-2 cells has been shown to correlate well with in vivo absorption of orally administered drugs in humans (Artursson and Karlsson, 1991). The cytotoxicity of the particles was determined using a CTB viability assay, DNA staining with 4',6-diamidino-2-phenylindole (DAPI) and impedance measurements performed with xCELLigence. These are three assays that are typically used for cytotoxicity testing; they measure the metabolic activity, the cell number and the morphological changes of the cells. They are suitable for silver nanoparticles, showing a definite concentration dependence. Nanoparticles. The silver nanoparticles AgPURETM (Lot# A1009016) were purchased from Rent a Scientist® GmbH (Germany) and contain 10% (w/w) silver stabilized with 4% (w/w) polyoxyethylene glycerol trioleate (trade name Tagat® TO), and 4% (w/w) polyoxyethylene (20) sorbitan monolaurate (Tween 20). They are being assessed for the use as a BAM reference material for particle size distribution of silver nanoparticles, which is provided as BAM N001, they are similar to the reference material reference material NM-300 available from JRC (Menzel et al., 2003; de van C.L. Klein, 2011). These particles were 7.02 ± 0.68 nm in silver metal core radius as determined by means of TEM measurements. They had a mean hydrodynamic radius in aqueous suspension of 14.7 ± 0.2 nm. In vitro digestion. The in vitro digestion was closely modeled in accordance with Versantvoortet et al. (Versantvoort et al., 2005). Initially, 15 mL of synthetic saliva was heated to 37°C in a water bath. Then 1 mL of silver nanoparticle dispersion was added to the saliva and stirred for 5 minutes. Subsequently, 35 mL of the artificial gastric juice was added to the solution and the pH value was set to 2 using hydrochloric acid. The solution was stirred again for 2 hours at 37 °C, and the pH value was monitored every half hour. After 2 hours the pH value was set to 7.5 by adding sodium bicarbonate powder to the reaction solution. Thereafter, 50 mL of artificial intestinal juice was added, and the solution stirred for two more hours. When the artificial digestion was completed, the samples were used without further processing for the analytical and toxicological testing. Table 1 shows the components of the artificial digestion solutions. The 6 integrity of the digestion enzymes is verified prior to every set of experiments using distinct control substrates for each step of the digestion process. Amylase activity is confirmed using amylopectin azure, pepsin activity is confirmed using an albumin/bromophenol blue complex; tryptic activity is confirmed using azocasein, and lipase activity is confirmed using 4-methylumbelliferyl oleate as substrates respectively. All resulting cleavage products are monitored photometrically. Asymmetrical flow field-flow fractionation (A4F). The A4F unit is manufactured by Postnova Analytics GmbH (Germany) and consists of an AF 2000 Focus system containing a PN 5200 sample injector, a PN 7505 inline degasser, and PN 1120 tip and focus pumps. The device was equipped with a custom-built slot outlet function for increased sensitivity. The channel thickness was 500 μm and a polyethersulfone (PES) ultra filtration membrane with a molecular weight cutoff of 4 x 103 g mol-1 was used. For detection a Spectrograph 3000 (LDC/Milton Roy [U.S.A]) UV detector was used at a wavelength of 430 nm. A solution of Millipore grade water with 200 mg/L of sodium azide and 500 mg/L of sodium dodecylsulfate (SDS) filtered through a 0.1 μm filter made of regenerated cellulose was used as the carrier solution. The samples were used without further dilution. A total of 200 μL of the sample was injected into the AF4, and the fractionated samples were collected with a Gilson Abimed 221XL automated sample collector for further analysis. The cross-flow was controlled by the AF2000 control software (Postnova Analytics) and set to an exponential decrease of 0.30 ml min-1 as exponent over 35 min from 1.5 ml min-1 to 0 ml min-1 . Small angle X-ray scattering (SAXS). The SAXS measurement was performed at the BAMline at BESSY (Berlin, Germany) on a Kratky type camera SAXSess (Anton Paar, Austria) at 25 °C (± 1 °C). The measuring time was 60 seconds. The measured intensity was corrected by subtracting the intensity of a capillary filled with pure eluent solution. After background correction the scattering data was deconvoluted (slit length de-smearing). All data processing was performed with the SAXSquant 3.5 software (Anton Paar, Austria). The scattering vector q is expressed in terms of the scattering angle Θ and the wavelength λ = 0.154 nm. Transmission electron microscopy (TEM). One drop of the A4F-fraction was dried dust free on a carbon-coated copper-grid and TEM images were taken with the TEM JEM 2200FS (JEOL, Japan) at 200 kV. Viability and toxicity assays. After exposure of the cells, the cell viability and proliferation of the Caco-2 cells was assessed using the commonly used Promegas Cell Titer Blue® (CTB) Assay, DAPI staining and impedance measurement with the xCELLigence from Roche. The CTB Assay is based on the ability of living cells to convert resazurin into resorufin and measures the oxidative metabolism. Non-viable cells rapidly lose their metabolic capacity and thus do not generate a fluorescent signal. DAPI is a DNA fluorescent stain, and the quantity of DNA is related to the number of cells. However, DAPI staining could be influenced by apoptosis, because apoptotic cells concentrate DAPI and thus could incorrectly indicate a higher cell number. To prevent misinterpretation, we tested undigested silver nanoparticles for apoptosis induction in Caco-2 cells, but could not detect an increase in apoptosis for up to 48 hours of exposure (data not shown). The impedance measurement uses the effects of the cells attached to the xCELLigence electrodes on the local ionic environment at the electrode solution interface. The impedance depends on the number of attached cells, and the quality of the cell interaction with the electrodes. An increase in the cell number, cell adhesion or spreading, will result in a change in electrode impedance, which is displayed as cell index (CI) values. 9 the metabolic activity and cell number. Finally, CTB was added to each well, incubated for an additional 2 hours for proliferating Caco-2 cells and 30 minutes for differentiated Caco-2 cells, and measured on a micro plate reader with 540 nm excitation and 590 nm emissions. After the CTB cell viability assay, the cells were fixed and lyced with methanol. The DNA was stained with 100 μL of 20 L of 20 μM M DAPI for at least 30 minutes. The resulting fluorescence was measured using a micro plate reader with 380 nm excitation and 460 nm emissions. The medium control was set to 100%. Means and standard deviations were calculated on the basis of at least three independent experiments. Statistical analysis was done with Excel Student's t-test. For the xCELLigence measurements the Caco-2 cells were transferred to E-plates at a density of 6125 cells per well in 200 μL culture medium. Then the proliferating Caco L culture medium. Then the proliferating Caco-2 cells were allowed to attach for 24 hours, and the differentiated Caco-2 cells were differentiated for 20 days before treatment. During this time the medium was replaced every 2 days. Subsequently, the culture medium was replaced by 200 μL nanoparticle suspensions with medium control, an L nanoparticle suspensions with medium control, andd different concentrations of digested and different concentrations of digested and undigested silver nanoparticles and corresponding control, consisting of the complete digestion fluid mixture after the artificial digestion process without silver nanoparticles. Cells were exposed for 48 hours. During this period the impedance was measured at least every minute. The incubation start was set to CI =1. The means and standard deviations were calculated transferred to at least two independent experiments. Microscopic images were taken in the 96 well plates with Zeiss Axio Observer and Zeiss AxioVison. To estimate the particle number per cell, we assumed the following: a mean radius 7.9 nm for the silver nanoparticles, a perfectly round shape, no agglomeration, a silver density of 10.49 g/cm3 , a cell number of 20 000 proliferating and 200 000 differentiated Caco-2 cells in the 96-Well plates and 12 250 proliferating and 112 000 differentiated cells in the E-plates. However, the values are only based on estimates and cannot duplicate the actual size distribution, shape and irregularities in the particle surface; this approximation is just provided together with the particle concentration to make the different assays more comparable. Prior to further treatment and application of the particles, their characteristics (in terms of core radius, hydrodynamic radius, size distribution and shape of the primary silver nanoparticles) were determined using A4F, DLS and SAXS measurements (see Figure 1). Detailed results are given below. Digestion of the silver nanoparticles was performed in accordance with the German DIN 19738 standard procedure and the similar approach of Versantvoort et al., (Versantvoort et al., 2005) with minor modifications as described in Figure 2. Samples were taken after each defined step of digestion, i.e. mouth, stomach and intestine, indicated as steps 2, 3 and 4, respectively. The samples taken from each step were consecutively fractionated with A4F without further preparation; subsequently, SAXS and DLS data were determined to obtain information about changes of shape and the size distribution. Fractions were collected at the maxima of the A4F UV traces after 22, 23, 46 and 50 minutes and analyzed offline by DLS in order to measure changes in the hydrodynamic radii, RH . The primary results 11 The hydrodynamic radius characterizes the overall size of the particle, including particle core, stabilizing agents, and other molecules, e.g. proteins attached to the particle's surface. Consequently, RH cannot be used to distinguish between aggregates of particles and particles capped with an additional protein corona. To compensate for this limitation, we also performed SAXS measurements on the fractionated samples. X-rays are scattered by electrons, and the scattering contrast is proportional to the square of electron density differences between the sample and its surroundings. Due to the large difference in electron density between the silver core and its surfactant-containing shell, SAXS only provides The SAXS results are in agreement with the A4F results, and also with the TEM measurements that are shown in Figure 4. The left-hand picture in Figure 4 shows well-dispersed particles prior to digestion without significant agglomeration and a mean radius of approximately 7.5 nm, whereas the right-hand picture shows the presence of agglomerated particles with radii between approximately 15 nm and 20 nm. In order to determine changes in the particle toxicity induced by digestion, different cell viability assays were employed on proliferating and differentiated Caco-2 cells. These cells were exposed to primary and digested particles as well as a digestion fluids mixture without nanoparticles for control. In addition, the impact of the nanoparticle stabilizer itself on the Caco-2 intestinal model was analyzed and had no significant effect (see supplementary material). The cell viabilities were measured using the CellTiter Blue® assay followed by a DNA cell staining with DAPI in the same 96-well plate (Ilavsky and Jemian, 2009). Additionally, impedance measurements were made with the xCELLigence System (Roche). The CTB® cell assay estimates the oxidative metabolism of viable cells by converting a redox dye (resazurin) into a fluorescent end product (resorufin). The data is corroborated by DNA staining with DAPI. We have chosen incubation times of 24 hours to allow pronounced cytotoxic effects. Proliferating and differentiated Caco-2 cells were exposed to primary and digested silver nanoparticles, as well as digestion fluids mixture without nanoparticles for control. The results of both assays for silver concentrations up to 25 μg/mL for proliferating and 100 μg/mL for differentiated cells are presented in Figure 5. The proliferating Caco-2 cells do not respond to the digestion fluid mixture without nanoparticles up to a concentration of 2.5% (this corresponds to the digestion fluid mixture content in 25 μg/mL digested nanoparticles), whereas the differentiated Caco-2 cells already start to lose metabolic activity at this concentration. Interestingly, the corresponding cell number detected by DAPI staining starts to decrease, but not until incubation with 5% digestion fluid mixture without nanoparticles (corresponds to the digestion fluid mixture content in 50 μg/mL digested nanoparticles). The digested and undigested silver nanoparticles decreased the cell viability of both growth stages of Caco-2 cells in a concentrationdependent manner. Although there were almost no apparent differences between digested and undigested nanoparticles. Initially, under the microscope, a detachment of proliferating Caco-2 cells became apparent at low concentrations • 5 μg/mL of silver nanoparticles as a first indication of the silver particles' cytotoxic effects, and before the results of CTB assays indicated distinct effects. The cells lost adherence and became spherical (see Figure 6). The xCELLigence system measures the electric impedance of cells grown on electrodes in real time. The electric impedance is displayed as the cell index (∆CI) curves (Figure 7), which provides information on the cell numbers and on inherent morphological and adhesive characteristics of the cells (Solly et al., 2004). The incubation of the cells with the digestion fluid mixture without nanoparticles showed no visible influence on the cell index up to 5% for proliferating Caco-2 cells (5% digestion fluid mixture corresponds to the content in 50 μg/mL digested nanoparticles), and up to 10% for differentiated Caco-2 cells (corresponds to 100 μg/mL digested nanoparticles). Within these concentration ranges, considerable negative/detrimental effects of digested silver nanoparticles were observed. These effects were particularly pronounced for proliferating cells compared to the undigested particles (Figure 7). The results of impedance measurements on the differentiated cell monolayer show clear differences between both digested and undigested particles. No significant effects of the digestion fluid mixture without nanoparticles on both proliferating Caco-2 cells up to 5% (corresponding content in 50 μg/mL digested nanoparticles) and differentiated Caco-2 cells up to 10% (equals 100 μg/mL digested nanoparticles) were observed. The A4F, DLS, and SAXS data indicate only small changes during the first step of the digestion process with regard to the size and shape of the silver nanoparticles, although the saliva solution contains a high content of ions as well as mucin, urea and amylase (see Table 1 for details). Due to the particles' steric stabilization with non-ionic surfactants, the ionic strength of the physicochemical environment seems to have only minor influence on the aggregation state. Furthermore, the mucins present in the solution, which have very high molecular weights of 120 kDa (Dekker et al., 2002), obviously do not bind to the particles. The small increase of the hydrodynamic radius could instead be due to the adsorption of small proteins like alpha-amylase with a molar mass of 42 kDa (Khoo et al., 1994). In contrast, the strong acidic shift from pH 6.4 in saliva to pH 2.0 in the gastric juice, which contains hydrochloric acid as well as salts, mucin and pepsin (see Table 1 for details), results in a major increase of the hydrodynamic radius, which suggests significant changes in the binding of the stabilizing agents followed by aggregation and/or formation of a protein corona. This is supported by A4F elugrams, in which the main peak becomes broader and shifts towards later elution times, and by the DLS data, which shows mean hydrodynamic radius changes from 14 nm to 99 nm. Furthermore, the SAXS measurements also show an increase in mean radius from 8.7 to 12.5 nm, and suggest the additional formation of larger particles or aggregates beyond the SAXS resolution range of 100 nm. The formation of those particle aggregates can be expected from the drop of pH, which causes protonation and diminished function of the stabilizer, and the presence of proteins, which may adsorb and act as junctions between the particles under these circumstances (Zook et al., 2011). The acidic environment of the gastric juice may lead to an increased ion release which was not monitored in this study. Liu et al. measured a weight loss of 30 % for a silver nanoparticle of 20 nm in diameter after 2 h at a pH 1.5. (Liu et al., 20012) However, a weight loss of 30 % would only result in a decrease of 2-3 nm in diameter for our particles, which hardly can be observed due to strong aggregation effects. The measured zeta potential of -20 mV measured for the untreated nanoparticles at pH 5.4 indicates at least partial ionic stabilization. At pH 2.0 the zeta potential of approximately +10 mV is slightly positive, and the contribution of electrostatic repulsion is negligible. The partial de-aggregation after the final pH increase may indicate a combination of the two effects. In the intestinal digestion step, the hydrodynamic radius decreased from 99 nm to 74 nm, whereas the core radius determined by SAXS remained the same. At pH 7.5 some of the more weakly bound proteins on the surface of the particles or aggregates may become detached again in the less acidic environment; this results in the reduction of hydrodynamic radius. In addition to salts, urea and enzymes, bile was also added during this step; the latter contains bile acids that act as surfactants and therefore can also induce deaggregation, at least in some caces. For a more comprehensive explanation of these findings detailed studies of the formation of a protein corona, as well as conformational changes or detachment rate measurements of the stabilizer under these conditions, are necessary. Such studies are beyond the scope of this study. The human dietary uptake of silver is estimated by 70 to 90μg/day, whereas in vivo studies on silver nanoparticles use up to 1000 mg/kg (Wijnhoven et al., 2009; Kim et al., 2008). In this context the used concentrations between 1 and 100 μg/mL are in a realistic concentration range. The cytotoxicity measurements using CellTiter Blue® assay and DNA staining indicate only a minor influence of the artificial digestion process to the impact of the tested material on Caco-2 cells despite the observed changes in aggregation status. Changes in cell morphology were visible under the microscope before the results of CTB assays indicated distinct effects. This behavior could be observed distinctively on proliferating cells, which exhibit a different metabolism and had not yet formed a tight monolayer with strong cell-cell contacts as the differentiated Caco-2 cells did. Furthermore, these effects appear earlier and at lower concentrations than the loss of metabolic activity compared to the associated DNA quantities (relative cell number, determined by DAPI staining). Although only differentiated Caco-2 cells represent the epithelial cell layer of the small intestines, immature intestinal cells are also present in the small intestines due to cell renewal there. Proliferating Caco-2 cells therefore proved to indicate external disturbances more sensitively when exposed to the silver nanoparticles, whereas the control experiments, with only the digestion fluid in the corresponding amounts, showed no visible or significant effects on the proliferating cells. In summary the Caco-2 cells first lost cell-cell contacts and became spherical prior to further damage and cell death. To verify this observation additional impedance measurement experiments were performed; they detect morphological changes in the cells with even greater sensitivity. The real time impedance measurement the silver nanoparticles showed a concentration and timedependent increase in the cell index compared to the medium controls followed by a decrease in both cell stages. Compared to medium controls, particle-treated cells showed a significant decrease in cell index curves for low concentrations during the next 10 to 20 hours; this resulted in a slope almost parallel to the abscissa. Differentiated Caco-2 cells are less susceptible to these effects. Except for untreated particles and elevated particle concentrations, a difference between the control groups and proliferating cells is rarely observed, and the cellular response to higher particle concentrations equilibrates after 6 hours. The digestion mixture itself has only a negligible direct adverse effect. This is also supported corroborated by a study from Frontela-Saseta et al. (Frontela-Saseta et al., 2011), who used a simplified in vitro digestion model, consisting of pepsin/HCl for 1 hour and salts/pancreatin for 2 hours in combination with proliferating Caco-2 cells and MTT assays. Their cells tolerated a 4% digestion control solution; a result that is comparable to the impedance measurements performed in the present work, which showed a resistance up to a concentration of 5%. This confirms our observation related to the wider diagnostic window of the impedance measurement compared with the metabolic activity, which is suitable for detecting gradual effect differences in digested and undigested nanoparticles. Taken altogether, the digested particles show a delayed and reduced response compared to primary silver nanoparticles at lower silver concentrations, but retain their toxicity in general. Especially for proliferating Caco-2 cells, effects can be detected using the xCELLigence impedance measurements at an earlier growth stage due to the sensitivity of the cells. The observed effects are in accordance with the fact that analytically observed changes in particle core size and hydrodynamic radius occur during artificial digestion. As a consequence of the aggregation of the particles and the adsorption of proteins, digested silver nanoparticles appear to have a slightly lowered bioavailability and silver ion release, and exhibit a change in toxicokinetic behavior compared to the undigested nanoparticles. The objective of this study was to obtain information on the impact of an artificial digestion on the physico-chemical properties and the cytotoxicity of well characterized silver nanoparticles. For this purpose, a combination of analytical particle separation and characterization techniques with common cytotoxicity testing methods was used. The nanoparticles were successfully separated from the complex matrix of the digestive fluids with asymmetrical flow field-flow fractionation and analyzed by small-angle X-ray scattering and dynamic light scattering. The resulting data show that the artificial saliva has almost no effect on the size and aggregation state of the nanoparticles. However, in the strongly acidic environment of the stomach and intestinal fluid, the DLS show a greatly increasing hydrodynamic particle radius. On the other hand, the SAXS data show that the majority of the particles are still in a monomeric state only a minor fraction is forming dimers or larger aggregates. This allows for the conclusion that an increase in the overall size of the particles is due to conformational changes of the stabilizing agents and/or to the adsorption of proteins. On basis of this data, different viability assays were performed on Caco-2 cells using undigested and digested particles to analyze the impact of the observed physicochemical changes on the particles' cytotoxic effects. The direct combination of in vitro digestion with Caco-2 cells has a clear limitation due to the effect of the high concentration of digestion mixture on the cells. Additionally, the classical end point methods, CellTiter Blue® (CTB) assay and the DAPI staining, did not appear to be sensitive enough to detect the more subtle differences at low particle concentrations which are visible in microscopic images. With the latter methods, the particles' toxicity remains the same whether digested or used from stock. In contrast, the more sensitive real time impedance measurement was appropriate to detect differences in cytotoxicity. In accordance with the analytically observed changes, the digested silver nanoparticles showed a delayed decrease in the impedance-time curves compared to the undigested nanoparticles. The results of the cytotoxicity tests in combination with the size and size distribution analysis may lead to the conclusion that the particles ́ surroundings, including stabilizers and a possible protein corona, do not have much impact on the cytotoxic effect of silver nanoparticles on the Caco-2 cells. The cytotoxicity is almost the same before and after digestion. Due to the fact that the cytotoxicity of silver nanoparticles is strongly size and ion release dependent, the lowered cytotoxicity visible in the more sensitive real-time impedance measurements is probably caused by the formation of small quantities of dimers and higher aggregates and hence a reduced silver ion release (Kim et al., 2012). However, the results confirms the interference of the digestive process with the physicochemical, toxic and toxicokinetic characteristics of nanoparticles, and this effect should be considered when testing adverse effects in in vitro models for oral adsorption studies. We would like to thank Olga Koshkina from the University of Mainz (Germany) and Ilona Dörfel from BAM for the TEM measurements. Furthermore we thank Heinrich Riesemeier and Ralf Britzke from BAM for their help at the BAMline. I declare no conflict of interest. Loeschner K, Hadrup N, Qvortrup K, Larsen A, Gao X, Vogel U, Mortensen A, Rye Lam H, Larsen E. 2011. Distribution of silver in rats following 28 days of repeated oral exposure to silver nanoparticles or silver acetate. Particle and Fibre Toxicology 8:1–14. certificates_media/rm_cert_particle_size/bam_n001repe.pdf Accessed on 5. June 2013 Table 1. Contents of the artificial digestion juices. hydrodynamic radii Rh were derived from . aNo second particle population could be detected aNo oligomers detected Figure 1. (a) A4F fractograms (UV/v is signal at 430 nm) with cross-flow profiles, (b) DLS data and (c) SAXS data size distribution with corresponding fits of the primary particles prior to the artificial digestion process and cytotoxicity testing. Figure 2. In vitro digestion process in three steps: the addition of saliva to the primary particles (1) simulates the conditions in the mouth (2); it follows the addition of gastric juice (3) and finally, intestinal juice (4). For composition of the various juices see Table 3. Each digestive step is analytically monitored. Figure 3. Analytics of the digested silver nanoparticles after the step with artificial saliva (2), gastric juice (3) and after the complete digestive process (4). (a) A4F fractograms with UV/vis detector signals at 430 nm, same cross-flow profile as in Figure 1(a) was used throughout. (b) DLS normalized field correlation functions G1 (t) data with stretched exponential fits, and (c) SAXS volume weighted size distributions of silver nanoparticles (circles), curve fits (solid lines) and contributions of monomers, dimers and trimers (dashed, dotted and dash-dotted lines). Insets: SAXS curves of silver nanoparticles and curve fits from Monte Carlo simulations (solid and dotted lines, respectively). Figure 4. Transmission electron microscopy images of silver nanoparticles prior to digestion (step 1) (left), and after artificial digestion (step 4) (right). Figure 5. Results of the CTB assays to assess cell viabilities (left) and DAPI staining to assess the cell number (right) of proliferating and differentiated Caco-2 cells after 24-hour incubation. Results using primary and digested silver nanoparticles, as well as digestion fluid mixture without nanoparticles for control (top) in DMEM/ITS medium buffer are shown. Figure 6. Microscopic images (200-fold magnification) of proliferating Caco-2 cells after 24 hours of exposure with digestion fluid mixture without nanoparticles, digested and undigested silver nanoparticles with concentrations of 5 and 15 μg/mL or 12•105 and 35•105 particles per cell are shown. Figure 7. The xCELLigence impedance measurements are shown for proliferating and differentiated Caco-2 cells. ∆ Cell index is plotted as a measure of impedance-based time-dependent cell response (TCRPs) of proliferating and differentiated Caco-2 cells. The cells were incubated in a nutrient solution with (top row) the addition of digestion fluid, with volumes corresponding to the addition of nanoparticles, with (middle row) digested and (bottom row) untreated silver nanoparticles, respectively. The nanoparticles concentration for proliferating cells are 5, 10 and 15 μg/mL or 19 105 , 38 105 and 57 105 particles per cell and for differentiated Caco-2 cells 15, 25 and 50 μg/mL or 12 105 , 21 105 and 41 105 particles per cell. jpg jpg A </doc> ###",
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"content": "Extract sample context and measurement list. <doc> S. Kittler,a C. Greulich,b J. S. Gebauer,c J. Diendorf,a L. Treuel,c L. Ruiz,d J. M. Gonzalez-Calbet,d M. Vallet-Regi,d R. Zellner,*c M. K€oller*b and M. Epple*a Received 22nd July 2009, Accepted 19th October 2009 First published as an Advance Article on the web 11th November 2009 DOI: Spherical silver nanoparticles with a diameter of 50 - 20 nm and stabilized with either poly(N-vinylpyrrolidone) (PVP) or citrate were dispersed in different cell culture media: (i) pure RPMI, (ii) RPMI containing up to 10% of bovine serum albumin (BSA), and (iii) RPMI containing up to 10% of fetal calf serum (FCS). The agglomeration behavior of the nanoparticles was studied with dynamic light scattering and optical microscopy of individually tracked single particles. Whereas strong agglomeration was observed in pure RPMI and in the RPMI–BSA mixture within a few hours, the particles remained well dispersed in RPMI–FCS. In addition, the biological effect of PVP-stabilized silver nanoparticles and of silver ions on human mesenchymal stem cells (hMSCs) was studied in pure RPMI and also in RPMI–BSA and RPMI–FCS mixtures, respectively. Both proteins considerably increased the cell viability in the presence of silver ions and as well as silver nanoparticles, indicating a binding of silver by these proteins. The biological activity of nanoparticles depends on various parameters, e.g. particle number, size, charge, and agglomeration. For the biological response, their size1–4 and surface chemistry5–7 are of utmost importance. As the strong antimicrobial activity of silver has stimulated the large-scale application of silver nanoparticles in medical devices and also in consumer appliances like refrigerators, mobile phones, and textiles, it is of considerable interest to investigate their behavior in a biological environment. Despite a number of biological studies on the toxicity of silver nanoparticles towards cells and bacteria,4,8–20 strict physicochemical data on the primary steps of the interaction with biological molecules are still missing. In particular, the interaction with proteins will influence the surface chemistry of nanoparticles and lead to changes in their charge and agglomeration state which may also be time-dependent. These, in turn, will influence the biological activity, e.g. the release of silver ions from the nanoparticles. Silver ions are most likely the active (1⁄4toxic) species in biological systems10,21 as also indicated by our earlier comparative studies with silver nanoparticles and silver ions.19,20 It was reported that silver nanoparticles aggregate in media with a high electrolyte content, and that the presence of proteins within the nanoparticle solution can stabilize the silver nanoparticles against aggregation.5,20 After cellular uptake, silver and gold nanoparticles were found in vesicles in the cytoplasm of macrophages.4 In contact to soft tissue after uptake into the body by any route the silver nanoparticles will be found within a protein-containing biological fluid where proteins will adsorb onto the particle surface. The nature and the concentration of these proteins will not only determine the nanoparticle behavior within the fluid, e.g. the agglomeration capacity, but also cellular uptake, intra-cellular distribution and possible toxic effects. This will also be of decisive interest for in vitro cell-biological studies where the agglomeration state of the applied nanoparticles is often neglected. Therefore, it was the purpose of this study to analyze the dispersability of silver nanoparticles under conditions close to body fluids. We used RPMI cell culture medium as a prominent example which was supplemented with different amounts of two proteins, i.e. blood serum (fetal calf serum, FCS) and a common serum protein (bovine serum albumin, BSA), and analyzed the cell-biological responses to dispersed silver nanoparticles using human mesenchymal stem cells as reporter cells. PVP-coated silver nanoparticles were synthesized by reduction with glucose in the presence of PVP according to Wang et al.22 Briefly, 2 g glucose and 1 g PVP were dissolved in 40 g water and heated to 90 C. Then 0.5 g AgNO3 dissolved in 1 mL water was quickly added. The dispersion was kept at 90 C for 1 h and then left to cool to room temperature. The particles were collected by ultracentrifugation (30 000 rpm, 30 min), redispersed in pure water and collected again by ultracentrifugation. Thereby NO3 , excess glucose and its oxidation products, excess PVP, and excess Ag+ were removed. The silver nanoparticles were then redispersed in water. The yield with respect to Ag was about 5%. PVP-coated silver nanoparticles for TEM analysis were synthesized by the polyol process from ethylene glycol following the method of Xia19,23 as described in detail in ref. 19. Citratestabilized silver nanoparticles for TEM analysis were prepared by dissolving 9 mg silver nitrate in 50 mL water and bringing it to boiling. A solution of 1% sodium citrate (1 mL) was added under vigorous stirring. The solution was kept boiling for 1 h, and then allowed to cool to room temperature. The silver nanoparticles were purified by ultracentrifugation (30 min at 30 000 rpm), followed by redispersion in water. The typical yield of citratestabilized silver nanoparticles was around 65% (with respect to silver). The final silver concentration in all dispersions was determined by atomic absorption spectroscopy (AAS). All dispersion experiments were carried out with a silver concentration of 50 mg mL1 . Polyvinylpyrrolidone (PVP K30, Povidon 30, Fluka, molecular weight 40 000 g mol1 ), tri-sodium citrate dihydrate (Fluka, p.a.), silver nitrate (Fluka, p.a.), and D-(+)-glucose (Baker) were used. Ultrapure water was prepared with an ELGA Purelab ultra instrument. The cell culture medium RPMI1640 was obtained from GIBCO, Invitrogen GmbH, Karlsruhe, Germany. It consists of amino acids (e.g., L-cystine, L-histidine), vitamins (e.g., biotine, folic acid), inorganic salts (e.g., calcium nitrate, magnesium sulfate) and other components such as D-glucose, glutathione and phenol red. Human mesenchymal stem cells (hMSCs, 3rd to 7th passage, Cambrex Bio Science, Walkersville Inc., MD, USA) were cultured in cell culture medium (RPMI–FCS) consisting of RPMI1640 supplemented with 10% fetal calf serum (FCS, GIBCO, Invitrogen GmbH) and L-glutamine (0.3 g L1 , GIBCO, Invitrogen GmbH) using 24 well cell culture plates (Falcon, Becton-Dickinson GmbH, Heidelberg, Germany). Cells were maintained at 37 C in a humidified atmosphere of 5% CO2. hMSCs were subcultured every 7–14 days depending on the cell proliferation. For that the adherent cells were washed with phosphate buffered saline solution (PBS, GIBCO, Invitrogen GmbH) and detached from the culture flasks by the addition of 0.2 mL cm2 0.25% trypsin–0.1% ethylenediamine tetraacetic acid (EDTA, Sigma-Aldrich, Taufkirchen, Germany) for 5 min at 37 C. Subsequently, the hMSCs were collected and washed twice with RPMI–FCS. Subconfluently growing hMSCs were cultured in RPMI1640 containing different concentrations (0.001% to 10%) of either FCS or bovine serum albumin (BSA, Serva Electrophoresis GmbH, Heidelberg, Germany) for 24 h. To the culture media, we added either 20 mg mL1 PVP-coated silver nanoparticles or 250 ng mL1 Ag+ (as silver acetate). All silver concentrations corresponded to the silver content in the solutions as determined beforehand by AAS in each case. The cell viability and the morphology of the incubated cells were analyzed after 24 h by calcein-acetoxymethyl ester (calcein-AM, Calbiochem, Schwalbach, Germany) fluorescence staining. After incubation, the cell cultures were washed twice with RPMI1640 and then incubated with calcein-AM (1 mM) for 30 min under cell culture conditions. Subsequently, the adherent cells were washed again with RPMI1640 and analyzed by fluorescence microscopy (Olympus MVX10, Olympus, Hamburg, Germany). Fluorescence microphotographs were taken with an F-view II camera (Olympus) and the Cell P software (Olympus) and digitally processed using Adobe Photoshop- 7.0. Phasecontrast microscopy was performed using a BX61 microscope (Olympus) and the F-view II camera. The quantification of cell viability was performed by phase analysis of calcein-positive fluorescence signals.24 Differences in the cell viability in the presence of the cell culture medium RPMI1640 containing different proteins and different protein concentrations were determined by the paired t-test. P values of less than 0.05 were considered to be statistically significant. Scanning electron microscopy (SEM) was performed with an FEI Quanta 400 ESEM instrument in high vacuum without sputtering. Transmission electron microscopy (TEM) was carried out both with JEOL 400EX and with JEOL 3000FEG electron microscopes, equipped with an Oxford LINK EDS analyzer. The samples were ultrasonically dispersed in ethanol and then transferred to holey carbon-coated copper grids. The hydrodynamic diameter and the zeta-potential were measured by dynamic light scattering (DLS) using a Malvern Zetasizer Nano ZS. In all cases, the z-average value was used as average particle diameter. The polydispersity index (PDI) was below 0.3 in all cases. The total amount of silver was determined by atomic absorption spectroscopy (AAS, Thermo Electron Corporation, M-Series). Dynamical viscosities of all solutions before the addition of nanoparticles were measured with an Ubbelohde capillary viscosimeter at 25 C. These data were used for the evaluation of the dynamic light scattering and optical microscopy data. The hydrodynamic diameter of our nanoparticles was also determined by the computerized evaluation of the Brownian motion of individual nanoparticles with a self-constructed setup. The setup essentially consisted of a modified reflected-light microscope with two lasers (632 nm HeNe and 488 nm Ar-ion) coupled into the system as additional illumination sources. This illumination technique provided enough scattered light to detect and track the motion of single nanoparticles down to about 10 nm in hydrodynamic diameter, depending on their refractive index. Individual particles were visible as scattering sources. The optical resolution of this approach obeys the Abbe-limit. A microscope was fitted with a calibrated CCD camera recording the motion of the individual nanoparticles with a rate of 30 frames per second. A LabView program was developed to track the coordinates of individual nanoparticles across the frames. The path length travelled by the individual particles per unit time was derived from these data and used for the determination of the hydrodynamic diameter of the nanoparticles by the Einstein–Smoluchowski and the Stokes–Einstein equations, respectively.25 The individual sizes of about 1000 to 3000 individual nanoparticles were then represented as a histogram, and the maximum was determined by fitting a suitable distribution function. All experiments were carried out at 20 C. The silver nanoparticles were analyzed by scanning electron microscopy as reported earlier.19 Both PVP- and citratestabilized nanoparticles were approximately spherical with typical diameters of 50 - 20 nm in both cases. Note that PVP gives a steric stabilization whereas citrate gives an electrostatic stabilization. The zeta-potential of PVP-stabilized nanoparticles was about 20 mV, and that of citrate-stabilized nanoparticles was about 30 mV. Further information on the nanoparticles' ultrastructure was obtained by high-resolution transmission electron microscopy (HRTEM) as shown in Fig. 1. All particles were clearly twinned. All dispersion experiments and cell-biological studies described in the following were carried out with PVP-stabilized silver nanoparticles. The hydrodynamic diameter of the nanoparticles was measured by dynamic light scattering (DLS). A representative curve is shown in Fig. 2. Note that the hydrodynamic diameter includes the polymer layer and the hydration shell. It is therefore always larger than the diameter of the silver core as determined by electron microscopy under high vacuum where the outer layers have collapsed. Size determination by Brownian motion analysis complemented the dynamic light scattering (DLS) data. Whilst both methods rely equally on light scattering, the DLS tends to overestimate the fraction of large particles due to their much higher scattering efficiencies. This source of uncertainty is absent in the Brownian motion analysis in which Brownian trajectories of individual particles are monitored (Fig. 3). As typical model proteins, we added bovine serum albumin (BSA) and fetal calf serum (FCS). Their concentrations in the RPMI solution were varied from 0.001 to 10 wt%. To exclude the time-dependent effect (see below), all data were recorded 20 min after the dispersion of the nanoparticles in the media, i.e. as fast as experimentally possible. We observed that the interaction of proteins with nanoparticles led to time-dependent agglomeration of the particles in biological media. This is in contrast to dispersion experiments in pure water where the PVP-coated nanoparticles were stable for at least 2 months and did not show sedimentation.19,20 The agglomeration was observed by both DLS and Brownian motion analysis. It is interesting to note that the particles remained stable in RPMI for about 5 h and then rapidly agglomerated and sedimented. The extent of sedimentation was clearly visible from the decrease of the total scattering intensity, i.e. the disappearance of particles from the scattering volume (Fig. 5). It was not possible to follow the particles by Brownian motion analysis because the particle number was significantly reduced in these processes. With the resulting particle concentrations no results with sufficient statistical reliability could be achieved. We ascribe the agglomeration effect in pure RPMI to the increased electrolyte content which reduced the electrostatic repulsion (note the negative zeta-potential). In RPMI containing 10% BSA, the agglomeration started soon after dispersion and also led to sedimentation (Fig. 6). In RPMI containing 10% FCS, the particles did not show any sign of agglomeration for at least one week (Fig. 7). These observations show that BSA initially stabilized the silver nanoparticle dispersion and prevented agglomeration. However, this stabilization was not permanent. After a couple of hours agglomeration, followed by sedimentation, occurred. In contrast, FCS seems to coat the particles and to prevent their agglomeration more permanently, possibly due to steric stabilization. As FCS consists mainly of albumin lipoproteins, glycoproteins, and globulins, this combination of biomolecules may be responsible for this stabilizing effect. In addition, a ligand exchange of PVP by proteins cannot be excluded. We note that electron microscopy of the particles after dispersion in these media would not give clear insight into the agglomeration phenomena because it is impossible to distinguish between those agglomerates that were present in solutions and those which were formed during the drying process. In addition, the presence of further solid components (proteins, salts) would strongly complicate the interpretation of the images. In addition to the size analysis experiments, cell-biological experiments were performed to correlate the actual agglomeration status of the nanoparticles with their biological impact on cells. Human mesenchymal stem cells (hMSCs) were used as reporter cells. The primary cell culture medium of subconfluently grown hMSCs was aspirated, and washed cells were then supplied with RPMI1640 containing different concentrations (0.001–10%) Representative cell culture results are shown in Fig. 8. In the presence of silver nanoparticles, cell-associated agglomerates were visible (see arrows in Fig. 8B and C) which were not observed in the presence of silver ions (Fig. 8E and F). As no agglomeration was observed in vitro in the presence of 10% FCS (Fig. 7), these agglomerates are probably due to cellular uptake of the nanoparticles and agglomeration within the cell, e.g. within a vesicle, as it was also reported by Yen et al.4 The addition of BSA to the cell culture medium led to a clearly enhanced cell viability in the presence of silver nanoparticles, except for very high BSA concentrations (Fig. 9). This protecting effect was even more pronounced for silver ions. In the presence of FCS, the viability of hMSCs in the presence of silver depended more strongly on the FCS content in the cell culture media (Fig. 10). Moreover, the viability of the cells in the presence of silver nanoparticles was correlated with the protein content in the media (r 1⁄4 0.932). A significant increase in the cell viability was observed at protein concentrations above 3% (see also Fig. 8C). However, the cell viability in the presence of silver ions was already increased above an FCS concentration of 0.1%. We found that PVP-stabilized silver nanoparticles rapidly agglomerate and precipitate in protein-free cell culture medium RPMI. The addition of BSA prevented this agglomeration only for a few hours. This was in contrast to FCS which seems to coat the particles and to prevent their agglomeration. An (even partial) ligand exchange of PVP by protein is also a possible explanation. However, in cell culture in the presence of FCS, cellular uptake may lead to intra-cellular aggregation in vesicles.4 These results are important to judge the fate of silver nanoparticles both during in vitro cell culture studies and also for toxicological testing because the uptake of nanoparticles by cells strongly depends on their size. mm-Sized agglomerates may undergo a different biological pathway than dispersed nanoparticles.2,4,7,26–28 Even at low concentrations of BSA or FCS, the toxicity against human mesenchymal stem cells is decreased with a more pronounced effect for FCS. It is believed that the silver-induced cell toxicity is mainly due to the release of silver ions.16 Thus, BSA and FCS obviously bind free silver ions, thereby reducing direct cell toxicity. It is quite conceivable that the toxic silver ions are bound by the proteins because they are present in large excess, except at the lowest concentrations that we used. However, several mechanisms may account for the decreased toxicity in the presence of BSA and FCS. Other serum factors such as cholesterol, triglycerides and phospholipids may also bind to silver nanoparticles.29 Very recently it was reported that BSA and silver nanoparticles rapidly form complexes by van der Waals and electrostatic forces.30 Furthermore, it is known that metal ion–albumin complexes are rapidly internalized by cells.30,31 The true nature of the protecting effect of proteins awaits further study, but it can already be stressed that toxicological studies and ''lethal concentrations'' for silver and silver nanoparticles must be taken with care as the agglomeration behavior and the presence of proteins are playing decisive roles. We thank the Deutsche Forschungsgemeinschaft (DFG) for financial support of this project within the Priority Program NanoBioResponses (SPP1313). We also thank the DAAD for generous funding within the program Acciones Integradas Hispano-Alemanas. </doc> ###",
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"content": "Extract sample context and measurement list. <doc> Department of Urban and Environmental Engineering, Graduate School of Engineering, Hokkaido University, North-13, West-8, Kita-ku, Sapporo 060-8628, Japan Received March 11, 2009. Revised manuscript received June 22, 2009. Accepted June 26, 2009. Although it has been reported that silver nanoparticles (Ag-NPs) have strong acute toxic effects to various cultured cells, the toxic effects at noncytotoxic doses are still unknown. We,therefore, evaluated in vitrotoxicity ofAg-NPs at noncytotoxic doses in human hepatoma cell line, HepG2, based on cell viability assay, micronucleus test, and DNA microarray analysis. We also used polystyrene nanoparticles (PS-NPs) and silver carbonate (Ag2CO3) as test materials to compare the toxic effects with respect to different raw chemical composition and form of silver. The cell viability assay demonstrated that Ag-NPs accelerated cell proliferation at low doses (<0.5 mg/L), which was supported by the DNA microarray analysis showing significant induction of genes associated with cell cycle progression. However, only Ag-NPs exposure exhibited a significant cytotoxicity at higher doses (>1.0 mg/L) and induced abnormal cellular morphology, displaying cellular shrinkage and acquisition of an irregular shape. In addition, only Ag-NPs exposure increased the frequency of micronucleus formation up to 47.9 ( 3.2% of binucleated cells, suggesting that Ag-NPs appear to cause much stronger damages to chromosome than PS-NPs and ionic Ag+. Cysteine, a strong ionic Ag+ ligand, only partially abolished the formation of micronuclei mediated by Ag-NPs and changed the gene expression, indicating that ionic Ag+ derived from Ag-NPs could not fully explain these biological actions. Based on these discussions, it is concluded that both \"nanosized particle of Ag\" as well as \"ionic Ag+\" contribute to the toxic effects of Ag-NPs. Nanomaterials areincreasingly beingmanufactured and used for commercial purposes because of their novel and unique physicochemical properties. These novel properties differ substantially from those bulk materials of the same composition. There are, however, rising concerns about the adverse effects of these materials on human health and environments. Some nanomaterials have been reported to produce reactive oxygen species (ROS) and exert cytotoxicity in vitro (1). Recent studies have shown that nanoparticles can readily pass through cell membranes (2, 3) and even biological barriers such as the blood-brain barrier and bloodtestis barrier (4), deposit in target organs, and interact with biological systems, which may create toxicity to living cells. Therefore, the establishment of principles and test procedures to ensure the safety of nanomaterials is urgently required. Of various nanomaterials, silver nanoparticles (Ag-NPs) are used most commonly in numerous consumer products including textiles, cosmetics, and health care products for exploiting its strong antimicrobial activity. Information on the toxicity of silver metals and silver salts, which have been used hitherto as silver-impregnated dressing and pharmaceuticals, is available (5), and they are considered to have no adverse effects on the human body when they used in reasonable amounts. However, despite widespread use, there is a serious lack of information concerning the toxicity of Ag-NPs to humans at the cellular and molecular level. There is growing evidence that Ag-NPs are highly toxic to various cultured cells. It has been reported that Ag-NPs exposures decreased viability, increased lactate dehydrogenase (LDH) leakage, or inhibited mitochondrial function in rat liver cells (6), mouse germline stem cells (7), human fibroblasts (8), and rat adrenal cells (9). Furthermore, Kim et al. demonstrated the dose-dependent changes of alkaline phosphatase and cholesterol values, which might be as a consequence of liver damage, in either the male or female rats following 28-day oral exposures to Ag-NPs (10). However, most of the studies evaluated the acute toxic effects of Ag-NPs at relatively high doses. It is required to evaluate the chronic toxicity at low doses, which could be developed by the prolonged internal exposure because Ag-NPs may remain in target organs for a long time. The genotoxicity such as mutagenicity and carcinogenicity of Ag-NPsis still largely unknown. In addition, the toxicity of Ag-NPs atmolecular level has not been reported yet so far. In this study, we therefore investigated toxic effects of Ag-NPs to human hepatoma derived cell line HepG2 that were exposed to Ag-NPs at low doses. For toxicity evaluations, cellular morphology, cell viability, and micronucleus formation were assessed under exposed conditions. Furthermore, DNA microarray analysis, which enables the examination of the expression of thousands of genes simultaneously and has been used in in vitro toxicogenomics, was performed to understand the cellular responses at molecular level. We evaluated the potential toxicity of Ag-NPs with emphasis on DNA damaging action or carcinogenicity by correlating cellular responses to gene expression patterns, which could provide a mechanistic understanding of the toxicity of Ag-NPs. Test Materials. Silver nanoparticles (Ag-NPs; 7-10 nm, stabilized with polyethylenimine) were purchased from Kyoto Nano Chemical Co., Ltd. (Kyoto, Japan). Polystyrene nanoparticles (micromerR, PS-NPs; 15 nm) was used as a control material because polystyrene itself has no apparent toxicity to human cells, but the PS-PNs are nanoscale particles that have nanoparticle properties, which displays the toxicity. To evaluate the toxic effect of ionic silver (Ag+) that could be released from the Ag-NPs, silver carbonate (Ag2CO3) was used as a controlmaterial. The Ag-NPs and Ag2CO3were purchased from COREFRONT Co., Ltd. (Tokyo, Japan) and Wako Pure Chemical Industries (Osaka, Japan), respectively. Cell Culture and Treatments. In this study, we used human hepatoma HepG2 cells, which retain normal cell functions and have been used in a number of toxicological studies. The cells obtained from the Riken Cell Bank (Tsukuba, Japan) were cultured in Eagle's minimal essential medium (MEM) (Nissui, Tokyo, Japan) supplemented with 1% nonessential amino acid (Invitrogen, Carlsbad, CA), 10% fetal bovine serum, and 60 mg/mL kanamycin at 37 °C and 5% CO2. In addition, to evaluate the contribution of Ag+ to the toxicity of Ag-NPs, 5 mM N-acetyl-L-cysteine was used as a strong Ag+ ligand. Neutral Red (NR) Uptake Assay. Viability of HepG2 cells after exposure to each material was determined by neutral red (NR) uptake assay. This assay was performed as described by Borenfreund and Puerner (11) with slight modification. Briefly, the cells were seeded in 96-well cell culture plate at a density of ca. 5.5 × 104 cells per well and incubated overnight. Following exposure to three test materials, the medium was replaced with MEM containing 0.005% neutral red dye. The plates were thenincubatedin a 5% CO2 incubator for 3 h at 37 °C. After incubation, the dye-containing medium was discarded. After washing with PBS, extractant solution (50% ethanol and 1% acetic acid) was added to each well. The microplates were shaken for few minutes, and the absorbance of solutions was measured at 540 nm using a microplate reader. All absorbance values were corrected against blank wells which contained growth media alone. Care was taken to ensure that the neutral red-containing plates and solutions were completely protected from light throughout the experimental procedure. Each assay involved eight wells per condition. Micronucleus Test. Cells were grown to 70% confluency in 60 mm culture dishes and were exposed to 1 mg/L of Ag-NPs for 24 h. The exposure doses were chosen as maximum concentrations, at which significant cytotoxicity was not observed by the neutral red uptake assay mentioned above. In addition, the cells were exposed to 1 mg/L of PS-NPs and 1.3 mg/L of Ag2CO3 (corresponding to 1 mg/L of Ag+) for comparison purposes. The cells exposed to the tested materials were trypsinized and incubated in a cold hypotonic solution (KCl 5.6 g/L) for 20 min and spread onto glass slides. After air-drying, the cells were fixed with methanol for 10 min and stained with 5% Giemsa. A total of 1000 binucleated cells were scored for the evaluation of the frequencies of micronucleus formation. Microarray Experiment. Cells were grown to 70% confluency in 60 mm culture dishes and were exposed to 1 mg/L of Ag-NPs, 1 mg/L of PS-NPs, and 1.3 mg/L of Ag2CO3 for 24 h. Following chemical exposure, the cells were washed with PBS and immediately subjected to RNA extraction. Three independent cultures were prepared for each treatment or control group. The cells were lysed directly on culture dishes, and total RNA was extracted using RNeasy Mini Kit (Qiagen, Hilden, Germany). Target preparation and hybridization were performed according to one-cycle eukaryotic target labeling assay protocols described in Affymetrix technical manual (Affymetrix, Santa Clara, CA). cDNA was synthesized from the total RNA by using One-Cycle cDNA Synthesis kit (Invitrogen, Carlsbad, CA) with a T7-(dT)24 primer incorporating a T7 RNA polymerase promoter. cRNA was synthesized from the cDNA and biotin-labeled by in vitro transcription using IVT Labeling kit (Affymetrix, Santa Clara, CA). Labeled cRNA was fragmented by incubation at 94 °C for 35 min in the presence of 40 mM Tris acetate, pH 8.1, 100 mM potassium acetate, and 30 mM magnesium acetate. Ten μg of fragmented cRNA was hybridized to a human genome focus array (Affymetrix, Santa Clara, CA) containing probes for 8795 human genes for 16 h at 45 °C. After hybridization, the microarrays were automatically washed and stained with streptavidin-phycoerythrin by using a fluidics station (Af- fymetrix, Santa Clara, CA). Finally, probe arrays were scanned with the Genechip System confocal scanner (Affymetrix, Santa Clara, CA). Microarray Data Analysis. Expression data of 12 samples (four treatments, n ) 3) stored as \"CEL file\" in the Gene Chip Operating Software (GCOS) (Affymetrix, Santa Clara, CA) were transferred into the Avadis 4.3 prophetic (Strand Genomics, Redwood City, CA). Signal intensity of probes were scaled and normalized by MAS5 algorism. These summarized data have been deposited to the National Center for Biotechnology Information (NBCI) Gene Expression Omnibus (GEO; http:// www.ncbi.nlm.nih.gov/geo), and are accessible through GEO series accession number GSE14452. From the results of detection call analyses, the genes with \"Present calls\" in three replications were selected and used in the subsequent steps. To identify differentially expressed genes, the unpaired t-test for control and respective treatment groups (n ) 3) was performed for each gene. From the results of these analyses, the genes with p < 0.05 and g2.0 fold change in either direction were identified as being differentially expressed. The genes that were differentially expressed by the test material treatments were functionally categorized based on gene ontology categories at level 6 and KEGG biological pathways by using web based gene ontology program Fatigo (http://fatigo.bioinfo.cipf.es). Cytotoxicity of Test Materials to HepG2 Cells. As the preliminary experiment, we assessed the cytotoxicity of three test materials (Ag-NPs, PS-NPs, and AgCO3) by measuring cell viabilities. The cell viabilities after 24 h exposure to the test materials were shown in Figure 1. Up to 0.5 mg/L of Ag-NPs, PS-NPs, and Ag2CO3, no significant cytotoxicity was observed, instead the cell viabilities increased up to around 120% relative to the nonexposed control. These results suggest that low dose of NPs accelerate cell proliferation. However, Ag-NPs exposure exhibited a significant cytotoxicity at higher doses (>1.0 mg/L), whereas PS-NPs and Ag2CO3 exposures did not produce a significant cytotoxicity. In this study, we chose 1.0 mg/L as an exposure dose for the following experiments, at which a significant cytotoxicity was not observed in the Ag-NPs exposure. control Ag-nanoparticle PS-nanoparticle Ag2CO3 Ag-nanoparticle + Cysteine frequencies of micrometeruclei (%) 2.1 ( 0.40 47.9 ( 3.2 2.5 ( 0.50 2.6 ( 0.36 29.3 ( 2.5 a For each treatment, triplicate biological samples were prepared, and 1,000 cells were analysed for each biological sample. Values are means ( SD. FIGURE 2. Venn diagrams illustrating shared gene expression in HepG2 cells under the exposures of three test materials (Ag-NPs, PS-NPs, and Ag2CO3). (SI) Figure S1 shows the general morphology of the HepG2 cells that were exposed to each test material at 1.0 mg/mL. There was no distinct change in cellularmorphology after 24 h exposure to PS-NPs and Ag2CO3 as compared with the control (nonexposed) cells. However, Ag-NPs exposed cells became abnormal in shape, displaying the widened intercellular spaces (cellular shrinkage) and pseudopodic form (acquisition of an irregular shape) (SI Figure S1B and E). Micronucleus Test for the NPs Exposed Cells. The frequencies of micrometerucleus formation in Ag-NPs, PS-NPs, or Ag2CO3 exposed cells are shown in Table 1. In the nonexposed (control) cells, micronuclei were found in 2.1 ( 0.40% of binucleated cells. Ag-NPs exposure remarkably increased the frequency of micronucleus formation up to 47.9 ( 3.2% of binucleated cells, indicating DNA damage and chromosome aberrations, while formations of micrometeruclei in PS-NPs and Ag2CO3 exposed cells were not significant (2.5 ( 0.50% and 2.6 ( 0.36% of binucleated cells, respectively). Genes Altered by Ag-NPs Exposure. Ag-NPs exposure altered the expression levels of 529 (induction: 236 and repression: 293) genes (Figure 2). To assess the effect of Ag-NPs exposure on cellular functions, we classified these altered genes functionally based on gene ontology (GO) categories of \"biological process\" (level 6). Figure 3 shows the major biological process, which assign functional characteristics, and the percentage of classified genes to total altered genes. From the results of this analysis, 521 genes could be annotated, and 255 biological processes were found. An important finding was remarkable inductions of genes classified in \"M phase\" (31 genes), \"microtuble-based process\" (19 genes), \"DNA repair\" (16 genes), \"DNA replication\" (24 genes) and \"intracellular transport\" (32 genes). The individual genes classified in these biological processes are shown in SI Table S1. Most of the genes classified in the \"M phase\", \"microtuble-based process\" and \"intracellular transport\", were involved in chromosome segregation, cell division, and proliferation. Furthermore, the genes categorized as \"DNA repair\" and \"DNA replication\" were involved in DNA biosynthesis and restoration of DNA after DNA damage. In this study, inductions of some well-known stressinducible genes were observed. Three metallothionein genes (MT1H; 4.5 fold, MT1X; 3.4 fold, and MT2A; 4.1 fold) and three heat shock protein genes (HSPA4L; 2.2 fold, HSPB1; 2.1 6048 9 ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 43, NO. 15, 2009 fold, and HSPH1; 2.1 fold) were significantly up-regulated in the cells exposed to Ag-NPs. Comparison of Ag-NPs-, PS-NPs-, and Ag2CO3-Altered Gene Profiles. In this study, we also performed DNA microarray analyses for the HepG2 cells exposed to PS-NPs and Ag2CO3 in the same manner to understand the cellular responses at molecular level. The exposure to PS-NPs and Ag2CO3 altered the expression levels of 189 (induction: 98 and repression: 91) and 304 (induction: 162 and repression: 142) genes, respectively. The overlaps of gene expressions among these two test materials and Ag-NPs are shown in Figure 2. In particular, 35 up-regulated genes and 27 downregulated genes were altered commonly in the same direction among all three chemical treatments. For further analyses, we compared the results of functional classification of altered genes among treatments with three test materials (Figure 3). In PS-NPs exposure, 188 genes were annotatable and 161 biological processes (level 6) were found. Of these, the up-regulations of the genes classified in \"M phase\" (14 genes), \"microtuble-based process\" (6 genes), \"DNA replication\" (4 genes) and \"intracellular transport\" (7 genes) were observed (SI Table S1). These classification patterns were similar to those in Ag-NPs exposurementioned above. In particular, 13 of 14 up-regulated genes classified in \"M-phase\" in the PS-NPs exposure overlapped with those in the Ag-NPs exposure, while the genes classified in \"DNA repair\" were not significantly overlapped (induction: 2 and repression: 5). In Ag2CO3 exposure, 298 genes were annotatable and 210 biological processes (level 6) were found. As shown in Figure 3 and SI Table S1, classification patterns of the altered genes were similar to those of Ag-NPs exposure. The inductions of genes classified in \"M phase\" (28 genes), \"microtuble-based process\" (17 genes), \"DNA repair\" (17 genes), \"DNA replication\" (24 genes), and \"intracellular transport\" (14 genes) were observed. Furthermore, we focused on the stress inducible genes that were remarkably induced by Ag-NPs exposure. Except for HSPB1, the genes up-regulated by Ag-NPs exposure mentioned above (MT1H,MT1X,MT2A,HSPA4L, andHSPH1) were not up-regulated by PS-NPs and Ag2CO3 exposure. Effects of Cysteine on Toxicity of Ag-NPs. In this study, 5 mM N-acetyl-L-cysteine was used as an Ag+ ligand to assess the contribution of Ag+ to the toxicity of Ag-NPs. In the presence of cysteine, no significant cytotoxicity was observed even at high concentrations of Ag-NPs (Figure 1). However, the cellular morphological change mentioned above was observed evenin the presence of cysteine (SI Figure S1E). Furthermore, the formation of micronuclei by Ag-NPs was partially counteracted by the addition of cysteine (formation frequency; 29.3 ( 2.5% of binucleated cells) (Table 1). The Ag-NPs exposure altered less number of total genes (induction: 213 and repression: 179) in the presence of cysteine. In these genes, only 165 (induction: 81 and repression: 84) genes were altered in the same direction in the absence of cysteine. The results of functional classification analysis of these genes were shown in Table S1 and Figure 3. The cysteine addition decreased the number of Ag-NPs induced specific genes that were classified into the \"M phase\" (22/31 genes: cysteine-treated/ nontreated cells) and \"DNArepair\" (11/16 genes: cysteine-treated/ nontreated cells) (SI Table S1). Furthermore, five stress inducible genes, which were significantly induced by Ag-NPs exposure, were downregulated (MT1H: -3.1 fold and MT1X: -3.1 fold) or did not exhibit significant expression level alteration (MT2A,HSPA4L, and HSPH1) in the presence of cysteine. To date, some studies have evaluated the acute toxic effects of Ag-NPs and demonstrated that Ag-NPs were highly cytotoxic to mammalian cells based on the assessment on mitochondrial function, membrane leakage of lactate dehydrogenase (LDH), abnormal cell morphologies (6, 7, 9). However, biological effects at noncytotoxic doses of Ag-NPs such as carcinogenesis are still unknown. The aim of this study was to evaluate potential toxicity of Ag-NPs at noncytotoxic doses and the general mechanism involved in the toxicity of Ag-NPs. For this purpose, we performed the neutral red uptake assay, micronucleus test and DNA microarray analysis for the HepG2 cells under noncytotoxic (100% cell viability detected by the neutral red uptake assay) exposure conditions of Ag-NPs. Furthermore, we used two test materials, PS-NPs and Ag2CO3, to evaluate the contribution of \"nanosized particle\" and \"ionic Ag+\" to the toxicity of Ag-NPs. The cytotoxicity of three test materials was assessed at various doses. Ag-NPs exhibited a significant cytotoxicity at high dose exposures (>1.0mg/L), whereas PS-NPs and Ag2CO3 had no measurable effects at the doses tested (Figure 1). In addition, abnormal cellular morphology was observed only in the Ag-NPs exposed cells (SI Figures S1B and E). Hussain et al. have shown that 5-50 mg/L of Ag-NPs exhibited a significant cytotoxicity in BRL 3A rat river cells. The cytotoxic doses determined in this study were on similar level with one reported in their report. Furthermore, it have been shown that Ag-NPs (5-10 mg/L, diameter: 15 nm) reduced mitochondrial function drastically and increased LDH leakage in the mammalian germline stem cells, whereas a significant cytotoxicity was not observed in Ag2CO3 exposed cells (7). Our experimental results agree with the result of this report. Intriguingly, noncytotoxic dose of Ag-NPs drastically increased cell viability. Thismay be a subsequence of hormesis, namely, stimulatory effects caused by low levels of potentially toxic agents. Furthermore, Ag-NPs and Ag2CO3, which did not produce a significant toxicity, also induced cell proliferation. It is obvious that the test materials used in this study would have some stimulant effects on human cells. In general, information on the genotoxicity of metal nanoparticles is limited. Exceptionally, it has been reported that ultrafine titanium dioxide (TiO2) induced micronuclei in Syrian hamster embryo fibroblasts (12). Micronuclei can be expressed in dividing cells as a result of chromosome breaks. At telophase, these fragments that did not reach the spindle poles during mitosis, form a separate and smaller nucleus. Micronuclei represent therefore a measure of DNA and chromosome breakage. In this study, the only Ag-NPs significantly increased the frequency of micronucleus formation (Table 1), suggesting that Ag-NPs have a potential to cause damage to chromosome. In contrast, PS-NPs and Ag2CO3 have no significant effect. Thus, the chromosome aberrations as well as cytotoxicty of Ag-NPs are likely to be mediated through a combined effect of \"nanosized particle\" and \"raw chemical composition of silver\". From the results of DNA microarray analysis, Ag-NPs induced larger number of genes than the other two test materials, indicating that As-NPs would affect various biological functions (Figure 2 and SI Table S1). In particular, only Ag-NPs induced well-known stress associated genes coding metallothionein (MT1H, MT1X, and MT2A) and heat shock protein (HSPA4L and HSPH1), which have been reported to be induced by cellular stresses such as heavy metal and various cytotoxic agent exposures (13, 14). From the results of functional classification of the altered genes, we highlighted the genes classifiedin biological process \"M-phase\". These genes are associated with cell cycle progression through mitotic (M) phase, and most of these genes are included in the \"cell cycle\" pathway based on the KEGG pathway mapping. In particular, we observed the increases in expression levels of checkpoint related genes (BIRC5, BUB1B, CCNA2, CDC25B, CDC20, and CKS2) (15-19) in the Ag-NPs, PS-NP, or Ag2CO3 exposed cells (SI Table S1). Abnormal expression of these genes would cause dysregulated cellular proliferation and play a critical role in carcinogenesis and tumor progression. Furthermore, induction of these genes has been observed by exposing to nongenotoxic carcinogens such as 12-O-tetradecanoylphorbol-13 acetate and tetrachloroethylene (20). In this study, the induction of these genes was found in all the cells exposed to three test materials including PS-NPs and would reflect the abnormal cell proliferation as shown in the cell viability assay (Figure 1). This suggests that the HepG2 cells likely respond to the nanosized particles regardless of their raw materials. In addition, induction of these genes by Ag2CO3 demonstrated that ionic Ag+ also promotes the cell proliferation. Therefore, the abnormal cell proliferating action of Ag-NPs would be mediated by both the nanosized particle and ionic Ag+. The genes classified in \"DNA repair\" were induced by only Ag-NPs and Ag2CO3, but not by PS-NPs. These Aginduced specific genes were involved in various DNA repair pathways activated by DNA damage, and its induction would be closely related to carcinogenesis. The previous report demonstrated that Ag-NPs induce an expression of RAD51 protein involved in DNA damage repair (21). In this study, induction of RAD51C gene, a member of the RAD51 family, was observed (SI Table S1). The RAD51 proteins including RAD51C are thought to promote DNA strand exchange and be involved in recombinational repair of damaged DNA (22, 23). Abnormal expression of RAD51 proteins has been reported in various tumor cells (24, 25). It has been reported that Ag+ binds with nucleobase covalently and increases DNA damage (26). The induction of DNA repair-associated genes by Ag2CO3 might reflect this interaction between Ag+ and DNA. However, the micronucleus formation was not significant in the Ag2CO3 exposed cells (Table 1). Thus, Ag+ binds with DNA, but does not cause damages to chromosome. Ag-NPs appeared to cause much more damages to chromosome than Ag+. Based on these discussions, it is concluded that both \"nanosized particle of Ag\" as well as \"ionic Ag+\" contribute to the DNA damaging action of Ag-NPs. In this study, we used cysteine, a strong Ag+ ligand, to assess the contribution of Ag+ to toxicity of Ag-NPs. Navarro et al. (27) have reported that cysteine abolished the inhibitory effects of Ag-NPs on photosynthesisin algae,Chlamydomonas reinhardtii and concluded that Ag-NPs contributed to the toxicity as a source of ionic Ag+. In this study, the addition of cysteine effectively inhibited the cell death (Figure 1) and the induction of stress-associated genes caused by Ag-NPs. These results suggest that ionic Ag+ contribute mainly to cytotoxic and stress associated effects of Ag-NPs. In addition, remarkable induction of cell proliferation by addition of cysteine (Figure 1) would be a consequence of growth stimulatory effects of NPs, which was also observed in PS-NPs exposure. This result was also supported by the induction of the genes classified in \"M-phase\" (Figure 3), which was only partially inhibited by cysteine. DNA damaging effect demonstrated by the micronucleus test was not completely counteracted (Table S1). Furthermore, the induction of the genes classified in \"DNA repair\" was only partially inhibited (Figure 3). These results suggest that the DNA damaging effect of Ag-NPs cannot be explained solely by the contribution of ionic Ag+ that is released from Ag-NPs. Thus, the nanosized particles of Ag alone have unique toxic effects to the cells. Metal ions including silver act as a catalyst and exhibit the ability to produce reactive oxygen species (ROS) in the presence of oxygen species, which is thought to be a mechanism of toxicity. The recent studies have indicated that Ag-NPs increased production of intracellular ROS (6). In addition, PS-NPs produced ROS in cell free medium (28). The ROS can act as signal molecules that promote cell cycle progression by affecting growth factor receptors, AP-1, NFkB, and so on (29-31) and induce the oxidative DNA damage. These mechanisms have been speculated to play important roles in carcinogenesis and tumor progressing actions of carcinogenic chemicals. Coincidentally, it has been reported that 2,3-dimethoxy-1,4-naphthoquinone (DMNQ), which has been known as a ROS generating chemical and frequently used as a model chemical for oxidative stress, induced the gene alteration patterns similar to one induced by Ag-NPs in this study (20, 32). Although little is known about relationship between carcinogenesis and ROS production by nanoparticles, it has been reported that nanoparticle carbon black induced DNA damage by ROS production, activating p53, proteinsinvolvedin DNA repair and regulation of cell growth and apotosis (33). Hence, it is speculated that the ROS production induces the genes associated with cell proliferation and DNA damage as shown in SI Table S1 and Figure 3. However, significant inductions of oxidative stress associated genes were not observed in this study, and a significant increase in intracellular ROS was not detected in the cells exposed to 1.0 mg/L of Ag-NPs by using a fluorescent probe 2′,7′-dichlorofluorescein diacetate (DCFH-DA) (34). (SI Figure S2). This is probably because the Ag-NPs concentration was too low to detect measurable ROS generation in the cells. It should be noted that such low dose of Ag-NPs caused a significant damage to chromosome (Table 1), reflecting a unique toxic effect of Ag-NPs. According to the criteria of the United States Environmental Protection Agency (EPA), silver is not classifiable as to human carcinogenicity (group D). Silver powder and colloidal silver do not induce cancer in animals, and silver chlorideis considered nonmutagenicin rec-assay. Thus, silver compounds have been generally considered not to have carcinogenicity in humans and animals. No evidence of the carcinogenicity of Ag-NPs has so far been reported despite the growing commercialization of Ag-NPs. In this study, however, the up-regulation of a number of the genes associated with DNA repair and the increase in micronuclei in the Ag-NPs exposed cells at relatively low doses (<1.0 mg/ L) clearly suggested the DNA damaging effects (chromosome aberration) of Ag-NPs. Both \"nanosized particles of Ag\" and \"ionic Ag+\" contribute to the toxic effects of Ag-NPs, DNA damaging action. The Ag-NP concentration assessed in this study would be higher than those occurring in air and water environment. However, since the internal kinetics of NPs has not been elucidated, the local concentration in tissues might reach higher level as the result of accumulation. In addition, since the physicochemical properties of Ag-NPs such as particle size, particle agglomeration, and dispersibility significantly influence the degree and actions of toxicity of Ag-NPs, further research is required to assess the effects of these variables. This research was carried out as a part of the 21st Century COE Program \"Sustainable Metabolic System of Water and Waste for Area-Based Society\". This study was also supported partially by a grant-in-aid (No. 19656129 and No.20360235) for Developmental Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan. Additional information including one table and two figures. This material is available free of charge via the Internet at http://pubs.acs.org. ES900754Q </doc> ###",
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"content": "Extract sample context and measurement list. <doc> Contents lists available at ScienceDirect journal homepage: www.elsevier.com/locate/btre C. Krishnaraj a, *, P. Muthukumaran b , R. Ramachandran a , M.D. Balakumaran a , P.T. Kalaichelvan a, ** aCentre for Advanced Studies in Botany, University of Madras, Guindy Campus, Chennai 600 025, Tamil Nadu, India b Centre for Biotechnology, Anna University, Guindy, Chennai 600 025, Tamil Nadu, India Article history: Received 23 April 2014 Received in revised form 14 July 2014 Accepted 8 August 2014 Available online 13 August 2014 Keywords: Acalypha indica Silver nanoparticles Gold nanoparticles MDA-MB-231 human breast cancer cells This study reports the in vitro cytotoxic effect of biologically synthesized silver and gold nanoparticles against MDA-MB-231, human breast cancer cells. Formation of silver and gold nanoparticles was observed within 30 min and the various characterization techniques such as UV–vis spectrophotometer, FE-SEM, TEM and XRD studies were confirmed the synthesis of nanoparticles. Further, MTT, acridine orange and ethidium bromide (AO/EB) dual staining, caspase-3 and DNA fragmentation assays were carried out using various concentrations of silver and gold nanoparticles ranging from 1 to 100mg/ml. At 100mg/ml concentration, the plant extract derived nanoparticles exhibited significant cytotoxic effects and the apoptotic features were confirmed through caspase-3 activation and DNA fragmentation assays. Thus, the results of the present study indicate that biologically synthesized silver and gold nanoparticles might be used to treat breast cancer; however, it necessitates clinical studies to ascertain their potential as anticancer agents. ã 2014 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/3.0/). Over the past few years, synthesis and characterization of nanoparticles has gained increasing momentum due to their large surface area to volume ratio because of which nanoparticles exhibit novel andnewproperties than theirmacroscopic counterparts.Thus, nanotechnology has immense potential to revolutionize in the biomedical research by developing new and improved products for clinical diagnosis and therapy. Several noble metal nanoparticles such as silver, gold, copper and platinumwere widely synthesizedby employing various procedures including physical, chemical and biological methods. The physical and chemical routes of nanoparticles preparation have many disadvantages and are not ecofriendly. Hence, researchers across the globe have searched for new and environmentally benign methods for the synthesis of biocompatible nanoparticles [29]. Incidentally, biological systems have long been known to reduce metal ions into nano-sized particles [7] and many researchers have recently reported the biogenic synthesis of silver and gold nanoparticles using a wide range of biological resources like bacteria [37], fungi[30,10]andplants [12,2].In theplantmediated greenchemistry approach, the reduction rate of metal salts is very fast and the procedure itself requires no specific conditions unlike the physical and chemical methods [29,32]. Besides, this biogenic method of nanoparticles synthesis appears to be reproducible and the particles, producedthroughthis environmentally friendly approach, are found highly stable[24].Hence, thisonepotgreenchemistryprocedurehas attractedthe attentionofbiologists andnanotechnologists inmyriad ways and is recently emerged as one of the active areas of current nanobiotechnological research. Breast cancer is the second leading cause of cancer death among women in the U.S. An estimated 39,620 breast cancer deaths and 232,340 new cases are expected among women in 2013 [5]. This data shows an increase of 100 breast cancer deaths and 1860 new cases compared to the previous report published in 2011 [4]. The existing cytotoxic agents used for the breast cancer treatment are found to be expensive and inefficient because they induce severe side effects due to their toxicity in noncancerous tissues [26,43]. Therefore, it is of urgent need to develop novel therapeutic agents http://dx.doi.org/ (P.T. Kalaichelvan). 2215-017X/ã 2014 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/3.0/). that are biocompatible and cost-effective. In recent times, nanotechnology based products such as nano-dresses, nanocars, skin creams, tennis rackets and balls have been increasingly introduced into the global market. To date, as many as 1628 nanobased products are being extensively used for various purposes throughout the world [34]. Inorganic nanoparticles have already been utilized in wound healing and in antibacterial applications [13]. Nowadays, silver and gold nanoparticles are emerging as promising agents for cancer therapy. The anticancer activities of nano-sized silver and gold particles have been evaluated against a variety of human cancer cells. However, very few reports were available against the breast cancer cells and most of these studies have mainly used chemically made nanoparticles [21,8,14]. Currently, there has only been a limited data existence for the cytotoxic effects of biologically synthesized silver and gold nanoparticles against human breast cancer cells [17,41]. The major objective of this work is to evaluate the cytotoxic effect of biosynthesized silver and gold nanoparticles against human breast cancer cell line. Our group has for the first time reported the biogenic synthesis of silver nanoparticles from Acalypha indica Linn leaves extract [28]. In continuation of this study, we screened the same plant for its ability to biosynthesize gold nanoparticles. Further, the cytotoxic effects of both silver and gold nanoparticles were tested against MDA-MB-231 cells by MTT assay and the possible mechanism for cell death was addressed through acridine orange and ethidium bromide (AO/EB) dual staining, caspase-3 and DNA fragmentation assays. Silver nitrate (AgNO3) and chloroaurate (HAuCl4) were purchased from Hi Media Laboratories Pvt. Ltd. Mumbai, India. MTT was obtained from Invitrogen, USA and acridine orange, ethidium bromide and all other fine chemicals were obtained from Sigma– Aldrich, St. Louis, USA. The fresh and healthy leaves of A. indica were collected from the Guindy campus of University of Madras, Chennai, India. Ten grams of freshly collected A. indica leaves were surface cleaned with running tap water followed by distilled water and boiled in 100 ml of distilled water at 60 C for 5 min. Then, the extract was filtered and used for the biogenic synthesis of both silver and gold nanoparticles. The biogenic synthesis of silver and gold nanoparticles was performed according to the standard published procedure with slight modifications [9]. The methods for the biosynthesis and characterization of silver nanoparticles from the leaves extract of A. indica were given in our previously published paper [28]. For gold nanoparticles biosynthesis,1 mM HAuCl4 was added to the broth containing 36 ml of leaf extract and 64 ml of distilled water at neutral pH. After this, the solution was kept at 37 C under static condition. Simultaneously, a control setup was maintained without adding HAuCl4. The pinkish violet colour formed after the addition of HAuCl4 was characterized using UV–vis spectrophotometer (Beckman DU-20 Spectrophotometer)in the range of 200– 700 nm. Further, the reaction mixture was subjected to centrifugation at 75,000 g for 30 min and the resulting pellet was dissolved in deionized water and filtered through Millipore filter (0.45mm). An aliquot of this filtrate containing gold nanoparticles was used for FE–SEM (Field Emission–Scanning Electron Microscopy), TEM (Transmission Electron Microscopy) and XRD (X-Ray Diffraction) analyses. For electron microscopic studies, 25ml of sample was sputter coated on copper stub and the size as well as shape of the gold nanoparticles was studied using FE-SEM and TEM. For XRD studies, dried gold nanoparticles were coated on XRD grid and the spectra were recorded by using Philips PW 1830 X-Ray generators operated at a voltage of 40 kV and a current of 30 mA with Cu Ka1 radiation. Human breast cancer cells (MDA-MB-231) were procured from National Centre for Cell Science, Pune, India. The cell lines were grown as a monolayer in Roswell Park Memorial Institute medium (RPMI) supplemented with 10% fetal bovine serum (FBS), penicillin/ streptomycin (250 U/ml),gentamycin (100mg/ml) and amphotericin B (1mg/ml) and incubated at 37 C in a humidified atmosphere of 5% CO2. Cells were grown confluence for 24 h before use. To determine the cytotoxic effect of both silver and gold nanoparticles, cell viability study was done with the conventional MTT-reduction assay with slight modifications [27]. Briefly, MDA-MB-231 cells were seeded in a 96-well plate at the density of 5 103 cells/well. The cells were allowed to attach and were grown in a 96-well plate for 24 h, in 200ml of RPMI with 10% FBS. After that the media was removed and replaced with suspension of various concentrations of AgNO3, HAuCl4, silver nanoparticles and gold nanoparticles viz., 1, 10, 50 and 100mg/ml (minimum 3 wells were seeded with each concentration). Equal concentrations of A. indica leaves extract were used as positive control and the cells were incubated for 48 h. After the addition of MTT (10ml, 5 mg/ml), the cells were incubated at 37 C for another 4 h. Optical density of the formazan product was read at 495 nm using scanning multi well spectrophotometer. The results were given as mean of three independent experiments. Acridine orange/ethidium bromide (AO/EB) dual staining was carried out to detect the morphological evidence of apoptosis in silver and gold nanoparticles treated cells. Twenty five microliters of treated and untreated cell suspension (5 106 cells/mL) was stained with 1ml of acridine orange and ethidium bromide dye mix (100mg/ml of acridine orange and ethidium bromide prepared in PBS separately) [42]. Then the samples were examined under fluorescent microscopy (Nikon Eclipse TS 100). Caspase-3 assay was carried out according to the procedure of Sutter et al. (2003) with slight modification [39]. The activity of caspase-3 was calculated from the cleavage of fluorogenic substrate Ac-DEVD-AMC (acetyl Asp-Glu-Val-Asp 7-amido-4 methylcoumarin). After 24, 36 and 48 h of incubation, silver and gold nanoparticles treated cell lysates were incubated with substrate solution (caspase-3 substrate Ac-DEVD-AMC 20 mg/ml, HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) 20 mM, glycerol 10%, dithiotheritol 2 mM, pH 7.5) for 1 h at 37 C and the cleavage of caspase-3 substrate was measured at an excitation wavelength of 390 nm and an emission wavelength of 460 nm. The activity was expressed as relative fluorescence unit (RFU). To investigate the internucleosomal DNA fragmentation caused by both silver and gold nanoparticles, DNA laddering assay was performed according to the standard procedure described by Su et al. (2005) with little modification [38]. A total of 1 106 cells was treated with silver and gold nanoparticles (100mg/ml) for 48 h and then collected by centrifugation. Further, the DNA was isolated using commercially available kit (Genei, Bangalore, India) following the manufacturer's instructions. DNA was resolved on 1.5% agarose gel (containing 3mg/ml of ethidium bromide in 1 TAE buffer of pH 8.5) at 90 V for 1.5 h and the bands were visualized using UV transilluminator. In this present study, gold nanoparticles were rapidly synthesized using A. indica leaves extract as bio-reductants. Similar to silver nanoparticles formation, the bio-reduction of HAuCl4 into gold nanoparticles was completed within 30 min of incubation. The very first indication for nanoparticles formation is colour change. A clear pinkish violet colour was formed within 30 min when 1 mM HAuCl4 was added into the aqueous leaves extract of A. indica, which indicates the biogenic synthesis of gold nanoparticles (Fig. 1). The intensity of pinkish violet colour was increased with the incubation period and it was due to the excitation of surface plasmon vibrations. On the other hand, control (leaf extract alone) showed no change of colour (Fig. 1). Very recently, Karuppaiya et al. (2013) have reported that the aqueous extract of Dysosma pleiantha rhizome rapidly biosynthesized gold nanoparticles within 20 min [25]. A characteristic absorption peak at 540 nm further confirmed the formation of nano-sized gold particles (Fig. 2). The formation of gold nanoparticles was started at 15 min and was completed at 30 min. Interestingly, the peak was found to be stable at the same wave length for up to 1 h, indicating that phytochemicals may have stabilized the synthesized gold nanoparticles (Fig. 2). Fig. 3a and b depict digitalized FE–SEM and TEM images of biosynthesized gold nanoparticles, respectively. These two images showed spherical shaped gold nanoparticles with a size of less than 30 nm. XRD analysis showed three distinct diffraction peaks at 38.1 , 44.1 and 64.1 which indexed the planes 111, 2 0 0 and 2 2 0 of the cubic face-centred gold. The obtained data was matched well with the Joint Committee on Powder Diffraction Standards (JCPDS) file no. 04–0784, which suggest the crystalline nature of gold nanoparticles (Fig. 4). The biogenic silver and gold nanoparticles were tested for their potent cytotoxic activity against MDA-MB-231, breast cancer cells. The results of the mechanistic studies indicated that silver and gold nanoparticles induced apoptosis through caspase-3 activation and DNA fragmentation. Different concentrations of AgNO3, HAuCl4, silver nanoparticles, gold nanoparticles and plant extract ranging from 1 to 100mg/ml wereused tostudy theviabilityofMDA-MB-231cells andthetoxicity was measured. Interestingly, HAuCl4, AgNO3 and A. indica leaves extract (positive control) treated cells did not show much toxic effects in all the tested concentrations; AgNO3 treated tumour cells showed more than 60% viable cells at 100mg/ml concentration (Fig. 5). Gold nanoparticles treated MDA-MB-231 cells exhibited slightly higher toxic effects than the silver nanoparticles at 1,10 and 50mg/ml concentrations; whereas, at 100mg/ml concentration, both silver and gold nanoparticles showed comparatively higher toxic effects (40%)than the other treated cells (Fig. 5). The results of this study suggest that the cytotoxicity of biologically synthesized silver and gold nanoparticles was increased with the increasing concentration of nanoparticles. Apoptotic morphological changes caused by both silver and gold nanoparticleswere studiedusing acridineorange/ethidiumbromide differential staining method. The stained cells were characterized to viable (light green), early apoptotic (bright green fluorescence and condensed chromatin), late apoptotic (orange fluorescence) and nonviable cells (red coloured fluorescence) (Fig. 6a–f). Both silver and gold nanoparticles treated cells showed condensed nuclei, membrane blebbing and apoptotic bodies. In contrast, the control cells showed intact nuclear architecture. However, very few apoptotic bodies were noticed in AgNO3 and HAuCl4 treated cells. To investigate whether apoptosis is mediated by caspase-3, cell lysates treated with AgNO3, HAuCl4, silver nanoparticles, gold nanoparticles and plant extract were analysed. Levels of caspase-3were found tobeelevated in thesilvernanoparticles treated tumour cells (Fig. 7). Plant extract treated cells exhibited slightly higher activity compared to gold nanoparticles treated ones. However, AgNO3,HAuCl4,treated cells showedmuchlower activity(Fig.7). The elevated level of caspase-3 was,further, confirmed bymeasuring the proteolytic activity of the fluorogenic peptide Ac-DEVD-AMC, a caspase-3specific substrate and its activitywas found to behighest at 48 h. The increased levels of caspase-3 activation suggest that silver and gold nanoparticles induce apoptosis in MDA-MB-231 breast cancer cells in a caspase-3-dependent manner. To investigate whether biologically synthesized nanoparticles induced cell death via apoptosis, DNA laddering assay was performed on agarose gel. A clear fragmented DNA ladders were observed in both silver and gold nanoparticles treated MDA-MB-231 cells whereas AgNO3 and HAuCl4 treated cells did not show such clear fragmented DNA ladders (Fig. 8). In addition, the untreated (control) cells did not show any prominent DNA ladders on the agarose gel. Therefore, the data obtained from this study confirms that both silver and gold nanoparticles induced cell death through apoptosis. In the recent years, biosynthesis of silver nanoparticles using plant extracts is getting more popular due to the strong antibacterial action of zerovalent silver and easy reduction of silver (I) salts. In our earlier study, silver nanoparticles were biosynthesized using aqueous leaves extract of A. indica as reducing and capping agents and those results were briefly discussed here [28]. The formation of silver nanoparticles was very rapid and it was completed within 30 min. The peak at 420 nm confirmed the biogenic synthesis of silver nanoparticles from A. indica leaves extract. Similarly, Jeyaraj et al. (2013) have recently reported that Podophyllum hexandrum leaves extract effectively synthesized silver nanoparticles at 420 nm [22]. Further, High Resolution – Transmission Electron Microscopy (HR-TEM) analysis confirmed the biosynthesis and the synthesized silver nanoparticles were predominantly in spherical shape with uniform size ranging from 20–30 nm. The XRD spectrum of biosynthesized silver nanoparticles was matched well with the JCPDS file no. 04–0783, which indicates the crystalline nature of face-centred cubic silver. These results were in good agreement with the recent reports. Interestingly, both silver and gold nanoparticles were formed within 30 min due to the rapid reduction of silver and chloroaurate ions by A. indica leaves extract. In contrast, Elavazhagan and Arunachalam (2011) have reported that Memecylon edule leaves extract took 1 h for the biosynthesis of gold nanoparticles while it was 3 h for silver [12]. However, in some studies, much faster rate of biosynthesis of silver and gold nanoparticles was observed. For instance, Dubey et al. (2010) have rapidly synthesized both silver and gold nanoparticles within 15 min from Sorbus aucuparia leaves extract [11]. Recently, Gangula et al. (2011) have reported that Breynia rhamnoides stem extract rapidly biosynthesized both silver and gold nanoparticles approximately 7 min and this is the much faster reduction process reported for the first time [16]. It is clear from these studies that the plant extract mediated biosynthesis is very simple, fast, low cost involvement, eco-friendly and safe for human therapeutic use [29,19]. Thus, this biogenic method of nanoparticles synthesis has much reduced impact to the environment and is recently emerged as viable alternative to conventional physical, chemical and even microbial methods. Silver and gold nanoparticles are being extensively synthesized using plant extracts, although the exact mechanism for this biogenic synthesis still remains to be completely unknown. However, a few hypotheses have been proposed to give some insights on the mechanical aspects of nanoparticles biosynthesis. Recent studies have shown that biomolecules such as protein, phenol and flavonoids present in the plant extract play an important role in the reduction of metals ions and capping of the nanoparticles [40]. Although the reduction of metal salts is environmentally benign, it is chemically a complex phenomenon involving an array of plant compounds such as vitamins, enzymes/ proteins, organic acids such as citrates, amino acids and polysaccharides [1]. The preliminary phytochemical screening of secondary metabolites has clearly revealed the presence of glucosides, flavonoids, phenolic compounds, alkaloids and carbohydrates in the leaves extract of A. indica (data not shown). We strongly believe that glucosides may be responsible for the bioreduction of both silver and chloroaurate ions. However, biosynthetic products or reduced cofactors may also play a key role in the reduction of respective salts to nanoparticles. In this present study, the cytotoxicity of silver and gold nanoparticles was increased with the increasing concentration of nanoparticles. This statement is true particularly in the case of MCF-7, another human breast cancer cell, which showed 100% cell Apoptosis is broadly considered as a distinctive mode of programmed cell death that eliminates genetically determined cells [15]. The induction of apoptosis is confirmed by two factors, (1) reduced and shrunken cells and (2) DNA fragmentation [36]. In this study, silver and gold nanoparticles treated cells showed apoptotic features such as condensed nuclei, membrane blebbing and apoptotic bodies at 48 h and these morphological changes were evident through AO/EB dual staining. Adding strengthen to the fact, silver and gold nanoparticles treated MDA-MB-231 cells showed clear fragmented DNA ladders, suggesting that cell death is due to apoptosis. In general, the fragmented DNA ladders indicate late apoptotic process in which caspase-3 plays a pivotal role [3,20]. The earlier studies have demonstrated that caspase-3 cascade activation is responsible for several apoptotic mechanisms [18]. Thus, it is obvious that DNA fragmentation and caspase-3 activation mediate the apoptotic process. In this present study, silver and gold nanoparticles treated MDA-MB-231 cells showed increased levels of caspase-3, indicating that apoptosis is mediated through caspase-3 cascade. These findings were coincided with the previous reports [17]. Caspase-3 activation may be initiated either through extrinsic pathway or intrinsic pathway due to the presence of toxicants in the surrounding environment [15,6]. In addition, caspase cascade activation is also reported to occur through the activation of granzyme B or death receptor or apoptosome [31]. In this study, although the silver nitrate caused cell toxicity was observed and the plant extract also up-regulated caspase-3 activity, however, only the gold and silver nanoparticles induced cell toxicity were specifically associated with all the observations of apoptosis including caspase-3 activity, AO/EB staining and DNA fragmentation. Apoptosis inducing agents that specifically target the tumour cells might have the potential to be developed as new antitumour drugs since apoptotic cell death does not induce an inflammatory response. The anti-inflammatory property of A. indica leaves extract was previously well studied [35]. As expected, both silver and gold nanoparticles biosynthesized from A. indica leaves extract did not show any inflammatory response, suggesting that nanoparticles targeted only the tumour cells. Based on the results obtained from these studies, it is quite apparent that biologically synthesized silver and gold nanoparticles have better therapeutic potentials than the reported chemically synthesized nanoparticles. Therefore, it might be worthwhile to explore the biosynthesized nanoparticles as a possible source of novel anticancer drugs. In this present study, silver and gold nanoparticles were rapidly synthesized using aqueous leaves extract of A. indica as novel source of bio-reductants. This single step procedure appears to be suitable for large scale production as it is simple, faster, costeffective, environmentally benign and safe for clinical research. Further, the plant extract derived nanoparticles exhibited strong cytotoxic effects against MDA-MB-231 cells, which suggest that biologically synthesized silver and gold nanoparticles might be used as novel anticancer agents for the treatment of breast cancer. However, the fate, transport and accumulation of nanoparticles inside the human body must be thoroughly studied prior to the approval to use as anticancer drug. The authors thank the Director, CAS in Botany, University of Madras for laboratory facilities. We are grateful to the Director, Centre for Biotechnology, Anna University for cell culture facilities. The authors are thankful to Dr. Udayakumar Muthulingam, Pachaiyappa's College, Chennai for taxonomical identification of the plant sample. The Head, SAIF, IIT-Madras is gratefully acknowledged for HR-TEM analysis. apoptotic effect of biologically synthesized silver nanoparticles using Podophyllum hexandrum on human cervical carcinoma cells, Colloids Surf. B 102 (2013) 708–717. </doc> ###",
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"content": "Extract sample context and measurement list. <doc> 1 Biopharmaceutics and Pharmaceutical Technology, Saarland University, Saarbruecken, Germany & 2 Merck KGaA, Darmstadt, Germany (Received 5 June 2008; accepted 29 September 2008) Recent opinions of the Scientific Committee on Consumer Products (SCCP) of the European Commission emphasize the missing validation of in vitro methodologies for nanomaterials and suggest that a review of the safety of insoluble nanomaterials presently used is required. Therefore the influence of Fluospheres† and silica nanoparticles, representing the class of organic and inorganic nanoparticles, respectively, on the lactate dehydrogenase (LDH) assay and the novel luminescence based Vialight† assay was tested. While LDH assay showed strong interactions with the tested silica particles, these problems may be overcome by novel methods like luminescence based assays. These findings suggest that even well characterized assay systems need a careful evaluation of the particle assay interactions when working with nanoparticles. Furthermore, particles based on the same material exhibit different biological properties depending on whether the material is used in micro- or nanometer range. Keywords: Cytotoxicity, silica, nanoparticles, LDH assay, luminescence assay Apart from many technical applications, nanoparticles of various materials are a promising raw material for a wide range of biological and pharmaceutical applications including therapeutics and medical diagnostics, as well as in food industry and for cosmetics (Allemann et al. 1998; Ahlin et al. 2002; Ito et al. 2005; Kukowska-Latallo et al. 2005; Chong et al. 2006). As a result of this multiplicity of application possibilities nanoscaled materials often come into contact with biological systems. Risk assessment and cytotoxicity evaluation are necessary to exclude negative effects resulting in tissue damage. Recent opinions of the Scientific Committee on Consumer Products (SCCP) of the European Commission suggest that a review of the safety of insoluble nanomaterials presently used in cosmetic products is required. According to the SCCP, nanomaterials should be treated as new chemicals from a risk point of view. Increased surface area and quantum effects are two principle factors that cause the properties of nanomaterials differing from bulk materials (SCCP 18 December 2007). But not only topical applied materials need to be characterised, also well established components of oral dosage forms, such as silica, require new evaluation. Since the 1940s silicium dioxide is present in tablets in the form of Aerosil†, an aggregate in the micrometer range formed by nanosized primary particles (Evonik 2007). Toxicity studies in rats showed no cytotoxic effects of these micrometer sized aggregates (Evonik 2008). Thus, not only an evaluation of micrometer sized aggregates but also of nanosized particles is necessary to determine the risks that a given material may present. The measurement principles to determine cytotoxic effects are manifold. However, optimal testing methods for nanomaterials have not yet been studied. The number of viability parameters addressed ranges from tests for metabolic activity and proliferation, the very basic activities of all living cells, to the determination of intracellular ATP. Assays like the uptake of neutral red give an impression of the amount of viable cells (Borenfreund and Puerner 1985). The ability to proliferate is an important mechanism which gives an impression of the health status of cells and can be addressed with the 5 bromo-2?-deoxyuridine (BrdU) assay (Gratzner 1982). The metabolic activity of cells is also an often assayed parameter for the determination of cell viability using the cleavage of tetrazolium salts, such as MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyl tetrazolium bromide), XTT or WST-1, to form a formazan dye (Mosmann 1983). The loss of cell membrane integrity can be addressed as a cell death parameter in different cytotoxicity assays such as the lactate dehydrogenase (LDH) or Toxilight† assay (measurement of adenylate kinase). Common to these assays is the detection of enzymes leaking from cells and using their catalytic function by colorimetric or chemoluminescent measurement. However, cell viability can be determined by the luminometric measurement of the intracellular ATP by the luciferin-luciferase assay. By artificial cell lysis ATP is released into the surrounding cell culture supernatant and then assayed. The light emission is linearly related to the ATP concentration (Crouch et al. 1993). As far as orally administered applications are concerned, influence of the administered substances on the gastrointestinal tract and on its barrier function has to be considered. The epithelial cell line CaCo-2 represents a widely accepted in vitro cell culture model for the intestinal mucosa displaying typical characteristics of the mucosal barrier, such as the formation of a confluent cell monolayer with functional tight junctions and the expression of characteristic transport systems (Artursson and Borchardt 1997; Fogh et al. 1977; Artursson et al. 2001). A number of cytotoxicity assays has been designed especially addressing the effect of test compounds on cell membrane integrity and hence on the barrier function of cells. Although cell viability assays analysing different cell death parameters are widely used and have been optimized for the use of microtiter plates to allow rapid and simultaneous measurement of many samples, current research shows that they still require validation and optimization for nanomaterials. Despite the fact that sensitivity and specificity of tetrazolium salt assays have been shown for different cell lines and test substances (Malich et al. 1997), recent publications suggest that nanomaterials interact with these assays and show the shortcoming of well characterised test systems (Worle-Knirsch et al. 2006; Laaksonen et al. 2007; Ulukaya et al. 2008). The decision which assay is suitable to assess the cytotoxic potential of a given compound or nanoparticle does not only depend on the cell death mechanism addressed but also on the material itself and its interactions with the assay components. Even a well characterized material may show different properties, if micro- or nanosized particles are compared. The object of this paper was to study the behaviour of two different nanoparticle types representing the classes of organic and inorganic nanoparticles in different cytotoxicity and viability assays and to evaluate a suitable assay for the cytotoxicity determination of the corresponding nanoparticles. Furthermore, it is shown that commonly used silica may exhibit different biological effects depending on the particle size. Dulbecco's modified Eagle's medium (DMEM) with 4.5 g/l glucose, L-glutamine and without pyruvate was purchased from Gibco/Invitrogen (Karlsruhe, Germany), non-essential amino acids (NeAA) from PAA (Pasching, Austria) and fetal bovine serum (FBS) from Sigma (Schnelldorf, Germany). Composition of Hank?s balanced salt solution (HBSS) was as follows: 136.9 mM NaCl, 5.4 mM KCl, 4.26 mM NaHCO3, 0.35 mM KH2PO4, 5.5 mM glucose, 10 mM HEPES, 1.26 mM CaCl2, 0.5 mM MgCl2*6H2O, 0.4 mM MgSO4*7H2O. HBBS was adjusted to pH 7.4 by means of NaOH. LDH assay was purchased from Roche (Mannheim, Germany), ATP stock solution, Vialight† Plus and Vialight† HS were purchased from Lonza (Wuppertal, Germany). CaCo-2 cells, clone C2Bbe1, were purchased at passage 46 from American Tissue Culture Collection (ATCC; Manassas, VA) and used at passages 58-72. Caco-2 cells were grown in a humidified incubator (378C in an atmosphere of 85% relative humidity and 5% CO2) to 90% confluency in 75 cm2 T-flasks with DMEM supplemented with 10% FBS and 1% non-essential amino acids (NeAA). Culture medium was changed every other day. After trypsinization, cells were seeded in 96-well plates at a density of 20,000 cells/well and incubated at 378C (85% relative humidity and 5% CO2) for eight days with change of culture medium every other day before being used in cytotoxicity assays. Silica nanoparticles with a size of 15 nm, as determined by transmission electron microscopy (data not shown), were a kind gift of Merck (Darmstadt, Germany). Aerosil† 200 was a gift from Evonik Degussa (Essen, Germany). Aerosil 200† is manufactured by flame hydrolysis and consists of microsized aggregates of 12 nm primary particles (Evonik 2007). FluoSpheres† with a size of 20 and 200 nm were purchased from Invitrogen (Karlsruhe, Germany). To ensure particle stability in physiological buffer, size and z-potential of all types of nanoparticles (dissolved in HBSS) were measured with the Zetasizer nano ZS (Malvern Instruments, Herrenberg, Germany) by means of dynamic light scattering (DLS) and Phase Analysis Light Scattering (PALS), respectively (Table I). The discrepancy in size measurement is a result of the fact, that dynamic light scattering measures the hydrodynamic diameter of the particles. LDH assay was performed with the LDH cytotoxicity detection kit. The cytoplasmic enzyme lactate dehydrogenase (LDH) is released through leaking membranes of dead or damaged cells. This enzyme can be detected in a two-step enzymatic test. The first step being the LDH catalyzed conversion of lactate to pyruvate, which results in the reduction of NAD to NADH/H and the second step being the reduction of 2-(4-Iodophenyl)-3-(4-nitrophenyl)-5 phenyl-2H-tetrazolium chloride (INT) to a watersoluble formazan dye. To test assay-particle interaction Fluospheres† and silica nanoparticles were diluted in HBBS in concentrations ranging from 10 ng/ml-10 mg/ml and 10 ng/ml-100 mg/ml, respectively; 100 ml particle dispersion were transferred to an optically clear 96-well plate and 100 ml of LDH reaction mixture were added. The assay was performed according to the manufacturer?s instruction in the absence of any cells. Fluorimetric measurement was performed with a Tecan SLT spectra (Tecan Deutschland GmbH, Crailsheim, Germany) at 492 nm. Vialight assay. ATP is present in all metabolically active cells. To detect the viability of cells a bioluminescent method was used in which the enzyme luciferase catalyses the formation of light from ATP and luciferin and the emitted light intensity is linearly related to the ATP concentration. For the determination of the cytotoxic potential of silica nanoparticles the particles were diluted in HBSS in concentration from 0.019-5 mg/ml. CaCo-2 cells were incubated for 4 h at 378C in an atmosphere of 85% relative humidity and 5% CO2 with different concentrations of nanoparticles. Two different assay kits were used: Vialight PLUS and Vialight HS. The Vialight PLUS assay was performed according to the manufacturer's instructions at room temperature with the exception that after cell lysis 20 ml of the samples were added to 80 ml HBSS buffer because of the high intracellular ATP levels. 100 ml of diluted sample were transferred to a white walled luminometer plate. Light intensity measurement was performed using a luminometer (MikroBeta† Trilux, Wallac, Shelton, CT, USA) and 1 sec integrated reading of the appropriate wells. Cells treated with HBSS only were used representing the values obtained for 100% viability and cells treated with 1% triton X-100 were used representing the values obtained for 0% viability. Vialight HS assay was performed in HBSS according to the manufacturer's instructions. To exclude cytotoxicity being caused by any soluble remains from particle preparation, silica nanoparticles were separated from the dispersant by centrifugation (30 min; 2000 g) in Centrisart† centrifugation tubes (Sartorius AG, Goettingen, Germany, cut-off 300 KDa). The supernatant was collected and used in a cytotoxicity assay according to the manufacturer's instructions and also served as a negative control. To test assay-particle interactions Fluospheres† and silica nanoparticles were diluted in HBSS in concentrations ranging from 30 ng/ml-20 mg/ml (Fluosphere†) and 30 ng/ml-30 mg/ml (silica nanoparticles), respectively. A 10 mM ATP stock solution was diluted in HBSS in concentrations of 0.003 mM, 0.03 mM, 0.3 mM and 3 mM. 50 ml of ATP dilution have been added to 50 ml of nanoparticle dilution resulting in an assay volume of 100 ml (final ATP concentration of 0.0015 mM, 0.015 mM, 0.15 mM and 1.5 mM, final nanoparticle concentration of 15 ng/ml-10 mg/ml in case of Fluospheres† and 15 ng/ ml-15 mg/ml in case of silica nanoparticles). To create a reference signal 50 ml ATP solution were added to 50 ml of HBSS buffer for each of the ATP concentrations. The assay was performed according to the manufacturers instructions by adding 50 ml of cell lysis reagent. After 10 min of incubation at room temperature, 100 ml were transferred to a white walled luminometer plate and 100 ml of ATP monitoring reagent was added. The generated luminescence signal of the ATP-particle mixture was measured using a luminometer and 1 sec integrated reading and is given as percentage of the artificial ATP signal. Data analysis. Graphic analysis was carried out using Sigmaplot† 9.0 (Systat software, Erkrath, Germany). Regression curves were determined using nonlinear regression procedure (four parameter logistic curve, [Equation 1]) The measurement of LDH leakage from damaged cells is commonly used as a marker for cell membrane integrity and cytotoxicity. The potential cytotoxic effects of silica particles, representing the class of inorganic nanoparticles, with a size of 15 nm and of commercially available Fluospheres†, representing the class of organic nanoparticles, with a size of 20 and 200 nm was assessed by LDH assay. In a first step, interactions of the test compounds with the LDH test system were addressed. Determination of substance interaction with the LDH assay reagents in absence of cells revealed, that the particles themselves led to a false positive signal. In case of the silica nanoparticles addition of the LDH reaction reagent caused precipitation. The occurring cloudiness interfered with the absorption measurement resulting in a higher absorption value in concentrations of 0.1-100 mg/ml. The measured absorption approached values of almost 2500% of the baseline value, thus being responsible for the occurrence of a false-positive signal (Figure 1). The turbid appearance of the 200 nm Fluospheres† particles also generated a higher absorption value for concentrations of 50 mg/ml and more. At a concentration of 10 mg/ml the absorption reached an almost 1800% increased value. To exclude the possibility that the red-fluorescence dye covalently linked to the particles may cause the increase in absorption, Fluospheres† with a size of 20 nm were used as control as they are missing the turbid appearance and show to be a clear, red coloured dispersion. Substance interaction tests exhibit a very slight increase in absorption for concentrations higher than 1 mg/ml, reaching a value of 210% of the baseline value for a concentration of 10 mg/ml. Regarding these findings it can be concluded, that the cloudiness in particle dispersions may interfere with absorbance measurements and is responsible for a false-positive signal in LDH assay which makes this assay not suitable for the tested silica particles and 200 nm Fluospheres†. Vialight. The Vialight† assay offers a novel testing method for cytotoxicity determining the intracellular ATP content of cells. ATP is necessary for luciferase to catalyse the formation of oxoluciferin and light, which can be assayed by bioluminescence measurement. By the addition of a cell lysis reagent the cell membrane is perforated, ATP leaks into the culture supernatant and is then measured by the luciferinluciferase reaction. The resulting bioluminescent emission of light is thereby linearly related to the ATP concentration. To evaluate the ATP concentration range where linearity is given, the assay was performed with artificial ATP concentrations from 0.0015 nM-1.5 mM ATP. The resulting light intensity (luminescence counts per secondLCPS) was then plotted double logarithmically against the ATP concentration. Linearity of the test system could be shown for ATP concentrations from 0.0015 mM- 1.5 mM ATP (Figure 2). To determine whether the generated luminescence signal would be affected by the nanoparticles, an artificial luminescence signal (0.0015 mM, The resulting luminescence was then calculated as percentage of the artificial ATP signal (Figure 3). The Fluospheres† themselves appeared as a red shimmering dispersion in concentrations of 0.15, 1.5 and 10 mg/ml, which was also cloudy in the two highest concentrations. A quenching of the luminescence signal could be detected for concentrations of 150 mg/ml and more (Figure 3A). Regarding the highest particle concentration of 10 mg/ml a clear drop in signal intensity was detectable resulting in a signal intensity of 20.5 (91.9)% of the 0.0015 mM ATP signal and 30.98 (90.49)% of the 1.5 mM ATP signal, respectively. This finding is in agreement with the results obtained for the Fluospheres† 20 nm particles (Figure 3B). A signal quenching was observed for particle concentration higher than 150 mg/ml resulting in a signal intensity of 29.69 (90,67)% of the 0.0015 mM ATP signal and 37.05 (92.72)% of the 1.5 mM ATP signal for a Fluospheres† concentration of 10 mg/ml. It can be concluded that the Vialight Plus† assay is not suitable for the cytotoxicity determination of the fluorescent 200 nm Fluospheres† nanoparticles, as the particles themselves are quenching the luminescence signal, which could be misinterpreted as a false positive result in a cytotoxicity assay. The results of substance interaction tests with the silica nanoparticles differ from the results obtained for the Fluorespheres† particles. SiO2 particle concentrations of 15 ng/ml to 15 mg/ml were tested in combination with four ATP concentrations Different Vialight† kits are commercially available, including the Vialight† Plus kit and the Vialight† HS kit. The Vialight† Plus assay kit is described by the manufacturer to have a more stable luminescence signal. This was verified by long time comparison of the EC50 values obtained from both assay types. Both assays were performed according to the manufacturer's protocol with the silica nanoparticles. The luminescence values were measured every 30 min over a period of 90 min. A dose-viability curve was established and the corresponding EC50 values calculated. Comparison of Vialight† HS and Plus assays revealed the expected enhanced signal stability for Vialight† Plus assay but also an increased signal accuracy. The luminescence signal generated by Vialight† Plus assay showed a very narrow standard deviation and remained unaltered for at least 90 min allowing the calculation of corresponding EC50 values over that period, while Vialight† HS assay showed a significant alteration in EC50 values and a broader standard deviation (Figure 4). As the Vialight† Plus assay was proven to exhibit the better signal accurancy and stability and to be unaffected by the silica nanoparticles, this assay was used to determine the cytotoxic effects of 15 nm silica nanoparticles. CaCo-2 cells were incubated for 4 h with particle concentrations from 0.019-5 mg/ml Although commercially available cytotoxicity assays have been well characterised and standardised it has been shown that cytotoxicity data calculated from these assays may differ significantly. Not only do different particle materials and modifications have an influence on the cytotoxicity and on the assay, but particle size itself is an influencing factor. Comparison of model agents such as triton X-100, chloroquine and sodium azid shows that the determination of cytotoxic effects depends on the used assay. The cytotoxic potential of membrane damaging agents like triton X-100 can be satisfactory assessed using LDH assay, but the same assay might be strongly influenced, when testing the cytotoxic potential of enzyme inhibitors like chloroquine (Weyermann et al. 2005). Furthermore, it has to be distinguished, which type of cells is used and whether effects on special cell organelles or general cytotoxicity are addressed. Assessing the toxicity of cadmium chloride (CdCl2) in vitro revealed different cytotoxicity profiles depending on the assay and used cell type. In HepG2 cells, a hepatoma cell line, the MTT assay revealed cytotoxicity of CdCl2 before effects with other tests systems where visible, as it was suggested that CdCl2 may have an effect on mitochondria in this cell line. However, HTC, another hepatoma cell line, showed a different result with neutral red uptake being the most sensitive cytotoxicity assay. Recent publications demonstrate that the choice of the best suitable assay does not only depend on the cell death mechanisms addressed, but also on potential assay-material interactions (Worle-Knirsch et al. 2006; Laaksonen et al. 2007). In this context single-walled carbon nanotubes (SWCNTs) have been shown to interact with MTT-formazan crystals, stabilizing their chemical structure and preventing them from being solubilized, a crucial step during analysis. No interaction was detected with other water-soluble formazan salts like WST-1, XTT or INT (Worle-Knirsch et al. 2006). To overcome disadvantages in analysing carbon nanotubes, a new clonogenic assay for the cytotoxicity testing of carbon-based nanomaterials has recently been described (Herzog et al. 2007). Furthermore, porous silica microparticles were shown to cause spontaneous redox reactions where MTT is reduced, while other silica based particles miss that ability (Laaksonen et al. 2007). The choice of drug has been shown to be a critical point in the reliability of cytotoxicity assays. In this context, the comparison of the inhibitory effect of different chemotherapeutics showed different inhibition levels depending on the assay and drug used. Thus, caution is necessary in the evaluation of chemotherapeutics as MTT assay gave rise to lower inhibition levels than an ATP based assay (Ulukaya et al. 2008). The findings presented in this study underscore these recently reported problems with colorimetric assays. Although well-described in literature, the LDH assay failed in the determination of the cytotoxic potential of 15 nm silica nanoparticles and 200 nm Fluospheres†. Addition of the reaction mixture to a silica nanoparticle gradient caused precipitation and a clouding of the normally clear dispersion resulting in an increasing absorbance, even in the absence of LDH. Precipitation may be the result of a change in pH from 7.4-8.6 when the LDH assay reagent is added to the cell culture supernatant. This leads to the conclusion, that the addition of the reaction mixture may alter the properties of a given dispersion and as a consequence leads to false-positive results. The cloudy appearance of a dispersion interacts with the absorbance measurement, which was also detected for the 200 nm Fluospheres particles. 20 nm Fluospheres, missing the cloudy appearance, showed only a slight increase in absorbance value in concentrations, where the dispersion is red coloured because of the covalently attached fluorescence dye. In contrast to absorbance measurements ATPbased cytotoxicity assays are based on the principle of bioluminescence detection. Being described as a potent method for the measurement of proliferation avoiding the use of radioisotopes (Crouch et al. 1993; Crouch 2000), the widely used luciferinluciferase assay is also a sensitive method for the determination of ATP levels in different cell types. Results of the ATP determination are available within 20 min after the incubation with test compound, which makes this assay a quick and simple method for cell viability determination. In order to assay the ATP content, cells are lysed by a suitable detergent or by lysis buffers provided within assay kits (e.g., Vialight†). The addition of lysis reagent to a silica nanoparticle gradient resulted in precipitation and clouding of the dispersion. To exclude the possibility of an interference of the cloudiness with the light emission, an artificial ATP signal was generated by addition of a defined ATP concentration. Linearity of the emitted light intensity can be shown for ATP concentrations from 0.0015-1.5 mM, thus easily providing a 4-log concentration range for cytotoxicity determination, which in other assay types must be achieved by additional background correction steps (Frgala et al. 2007). Although the cloudy appearance of the silica particle dispersion is not a hindrance for the test system, there are other disadvantages to overcome. As it was shown, the red fluorescent Fluospheres† led to a strong quenching of the luminescence signal, resulting in a false positive signal. Although only the 200 nm Fluospheres† are of cloudy appearance, both, the 20 nm and the 200 nm Fluospheres, cause a reduction of the signal intensity. So it is likely that reduced luminescence intensity is a result of the emissioned light interacting with the fluorescence dye. Even assays based on the same biological cell death parameter and measurement principle may differ significantly in their correct determination of cytotoxic effects, as could be shown for the Vialight† Plus and Vialight† HS assay. However, Vialight† Plus proved useful for silica nanoparticles, but not for fluorescently labelled particles. Size depending effects influencing the electronic structure of nanoparticles cause the properties of nanomaterials differing from bulk material (Nel et al. 2006). This has been shown to result in size dependent optical and electronic properties (Chiu 2004; Wang et al. 2005; Balamurugan and Toshiro 2006). Greater surface area per mass of nanosized particles compared with larger-sized particles of the same chemistry affects biological activity (Oberdo ̈rster et al. 2005b). The here described findings suggest that sizedependent effects also affect biological properties. Aerosil†200, commonly used in solid oral dosage forms, consists of aggregates in the micrometer range formed by 12 nm primary particles. Early toxicity studies in rats showed no cytotoxicity after oral administration (LD50 of more than 10000 mg/ kg) (Evonik 2008). This is in accordance with the results obtained by the luminescence based assay, where Aerosil†200 showed no cytotoxic effect on the colon carcinoma cell line CaCo-2. In contrast to this bulk material, a clearly detectable cytotoxic effect was observed for 15 nm silica nanoparticles, thus pointing out a discrepancy between the cytotoxic effect of 15 nm sized primary particles and micrometer sized aggregates of the same material. The potential of nanoparticles to agglomerate or deagglomerate in physiological media is sensitive to ions and proteins. Aggregation state and presence of protein coatings may alter the toxic effect of nanoparticles due to changes of surface coating or the transport mechanism (Schulze et al. 2008). Thus, surface charge, particle size and size distribution, as well as agglomeration state, chemical composition, and porosity are important characteristics in understanding the toxic effects of test materials (Oberdo ̈rster et al. 2005a). Recent publications underline the need for a categorization framework to aid hazard identification of nanomaterials. Furthermore, they suggest that nanomaterials will present hazards based on their structure and physicochemical properties, thus chal- lenging many conventional approaches to risk assessment (Maynard 2007; Hansen et al. 2008). The data presented in this study support this opinion and show that even well described materials like silica may interfere with standardised test systems. The choice of the best suitable assay does not only depend on the cytotoxicity parameter addressed but also on the particle itself and its interaction with different test principles. The new class of nanosized materials may have different effects and therefore a close examination of particle-assay interactions becomes necessary before any cytotoxicity assay can be applied. As recent publications reported the use of mesoporous silica nanoparticles for drug delivery applications (Chung et al. 2007; Salonen et al. 2005; Heikkila et al. 2007a,b) the findings presented herein clearly address the necessity for an evaluation of materials considered harmless when the border from micrometer to nanometer range is crossed. For the provision of silica nanoparticles and for financial support we thank Merck KGaA. Declaration of interest: The authors report no conflicts of interest. The authors alone are responsible for the content and writing of the paper. nanoparticles in 3T3-L1 cells and human mesenchymal stem cells. Biomaterials 28(19):2959-2966. This paper was first published online on iFirst on 12 November 2008. Failure of MTT as a toxicity testing agent for mesoporous silicon microparticles. Chem Res Toxicol, 20(12):1913-8. </doc> ###",
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