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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> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 84.01507}, {\"h\": 24.0, \"mgL\": null, \"viab\": 65.78499}, {\"h\": 24.0, \"mgL\": null, \"viab\": 54.64141}, {\"h\": 24.0, \"mgL\": null, \"viab\": 41.72621}, {\"h\": 24.0, \"mgL\": null, \"viab\": 12.871026}, {\"h\": 24.0, \"mgL\": null, \"viab\": 15.307593}]}", "role": "assistant" } ]	171.json
[ { "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> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 96.02428}, {\"h\": 24.0, \"mgL\": null, \"viab\": 96.874054}, {\"h\": 24.0, \"mgL\": null, \"viab\": 100.0607}, {\"h\": 24.0, \"mgL\": null, \"viab\": 92.2003}, {\"h\": 24.0, \"mgL\": null, \"viab\": 72.23065}, {\"h\": 24.0, \"mgL\": null, \"viab\": 43.12595}, {\"h\": 24.0, \"mgL\": null, \"viab\": 39.30197}, {\"h\": 24.0, \"mgL\": null, \"viab\": 18.057663}]}", "role": "assistant" } ]	170.json
[ { "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€ollerb 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> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 88.548386}]}", "role": "assistant" } ]	167.json
[ { "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. 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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> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 48.0, \"mgL\": null, \"viab\": 62.41379}, {\"h\": 48.0, \"mgL\": null, \"viab\": 66.206894}, {\"h\": 48.0, \"mgL\": null, \"viab\": 77.586205}, {\"h\": 48.0, \"mgL\": null, \"viab\": 80.0}]}", "role": "assistant" } ]	164.json
[ { "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 MgCl26H2O, 0.4 mM MgSO47H2O. 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> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 4.0, \"mgL\": null, \"viab\": 103.35135}, {\"h\": 4.0, \"mgL\": null, \"viab\": 103.35135}, {\"h\": 4.0, \"mgL\": null, \"viab\": 104.0}, {\"h\": 4.0, \"mgL\": null, \"viab\": 100.54054}, {\"h\": 4.0, \"mgL\": null, \"viab\": 88.64865}, {\"h\": 4.0, \"mgL\": null, \"viab\": 39.783783}, {\"h\": 4.0, \"mgL\": null, \"viab\": 16.864864}, {\"h\": 4.0, \"mgL\": null, \"viab\": 12.972973}, {\"h\": 4.0, \"mgL\": null, \"viab\": 15.351352}]}", "role": "assistant" } ]	159.json
[ { "content": "Extract sample context and measurement list. <doc> View Article Online View Journal This article can be cited before page numbers have been issued, to do this please use: M. O. Kim, S. A. Shah, G. Yoon, A. Ahmad, F. Ullah and F. Amin, Nanoscale, 2015, DOI: . This is an Accepted Manuscript, which has been through the Royal Society of Chemistry peer review process and has been accepted for publication. Accepted Manuscripts are published online shortly after acceptance, before technical editing, formatting and proof reading. Using this free service, authors can make their results available to the community, in citable form, before we publish the edited article. We will replace this Accepted Manuscript with the edited and formatted Advance Article as soon as it is available. You can find more information about Accepted Manuscripts in the Information for Authors. Please note that technical editing may introduce minor changes to the text and/or graphics, which may alter content. The journal's standard Terms & Conditions and the Ethical guidelines still apply. In no event shall the Royal Society of Chemistry be held responsible for any errors or omissions in this Accepted Manuscript or any consequences arising from the use of any information it contains. www.rsc.org/nanoscale Shahid Ali Shah PhD, Gwang Ho Yoon MS, Ashfaq Ahmad PhD, Faheem Ullah MS, Faizul Amin PhD and Myeong Ok Kim* Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. Department of Biology and Applied Life Science, College of Natural Sciences, Gyeongsang National University, Jinju, 660-701, Republic of Korea * Corresponding author Myeong Ok Kim, Prof. Ph.D, Head of Neuroscience Pioneer Research Centre, Department of Biology and Applied Life Science, College of Natural Sciences, Gyeongsang National University, Jinju, 660-701, South Korea Tel.: +82-55-772-1345 Fax: +82-55-772-1349 E-mail: The adverse effects of nanoscale-alumina (Al2O3-NPs) have been previously demonstrated in both in vitro and in vivo studies, whereas little is known about its mechanism of neurotoxicity. It is the goal of this research to determine the toxic effects of nano-alumina on human neuroblastoma SH-SY5Y and mouse hippocampal HT22 cells in vitro and on ICR female mice in vivo. Nano-alumina displayed toxic effects on SH-SY5Y cells lines in three different concentrations also increased aluminium abundance and induced oxidative stress in HT22 cells. Nano-alumina peripherally administered to ICR female mice for three weeks increased brain aluminium and ROS production, disturbing brain energy homeostasis, and led to the impairment of hippocampal dependent memory. Most importantly, these nano-particles induced Alzheimer disease (AD) neuropathology by enhancing the amyloidogenic pathway of Amyloid Beta (Aβ) production, aggregation and implied the progression of neurodegeneration in the cortex and hippocampus of these mice. In conclusion, these data demonstrate that nano-alumina is toxic to both cells and female mice and that prolonged exposure may heighten the chances of developing a neurodegenerative disease, such as AD. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. Key words; Nano-alumina, ICR mice, Alzheimer disease, ROS, phospho-tau, SH-SY5Y cells 2 plaques and neurofibrillary tangles (NFT), respectively17, 18. Aluminium can cross the BBB and accumulate in different brain regions, including the cortex and hippocampus, and induce oxidative stress by causing damage to membrane lipids, proteins and the antioxidative enzyme defence system19-23 . The purpose of this study was to test, for the first time, the adverse effects of nano-alumina on SH-SY5Y and HT22 cells line in vitro and simultaneously evaluate the hazardous consequences and cognitive impairment after peripheral administration of these nanoparticles to female ICR mice. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. The size and shape of nano-alumina was analysed through Transmission Electron Microscopy (TEM). ICR (Institute of Cancer Research) female mice (10 weeks of age) used in the present study were purchased from Sam Tako Inc., Korea. Nano-alumina was administered at 50 mg/kg intraperitoneally (i.p),9 two times a week for three weeks. Both treated and untreated animals were sacrificed three weeks after nano alumina administration. A great care was taken while handling the animals. All of the experimental procedures were approved (Approval ID: 125) by the local animal ethics committee (IACUC) of the Division of Applied Life Sciences, Department of Biology Gyeongsang National University South Korea. The following two types of cell cultures were used. 1) Mouse hippocampal HT22 neuronal cells 2) Human Neuroblastoma SH-SY5Y neuronal cells HT22 (a generous gift from Prof. Koh (Gyeongsang National University) and SH-SY5Y neuronal cells were purchased from KOREAN CELL LINE BANK, Seoul, Korea (KCLB). Both (HT22 and SH-SY5Y) neuronal cells were cultured in DMEM (Dulbecco's modified Eagle medium), containing high D-glucose, 10 % FBS and 1 % antibiotics (penicillin-streptomycin) at 37°C in humidified air containing 5 % CO2. For performing different assays, the cells were plated in 96-well plates at a required density of at least 70% confluence. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. For fluorescence-labeled nano-alumina formulation, we used a previously described method with some modifications to conjugate rhodamine-123/coumarin-6 to nano-alumina24. Briefly, 5 mg of rhodamine-123/coumarin-6 was dissolved in 25 ml of acetone, and added to 100 ml of nanoalumina under stirring. This mixture was then further stirred for 3 h at 60 °C and then cooled to room temperature. The solution was then centrifuged at 4000 rpm to remove unattached dye. This procedure was repeated three times to ensure that no free dye molecules remained in the final conjugate. The uptakes of rhodamine-123/coumarin-6 conjugated nano-alumina were studied in mouse bEnd3 cell line which served as an in vitro blood brain barrier (BBB) model (ATCC, USA) and human neuroblastoma SH-SY5Y cells, using confocal microscopy (Fluoview FV 1000, Olympus, Japan). Freshly prepared rhodamine-123/coumarin-6 conjugated nano-alumina suspension at concentration of 0.1 mg/ml was mixed with cell culture medium and added to the cells pre-cultured in 4-well cover glass chambers and incubated for 24 h. Following incubation, the cells were washed three times with PBS and were fixed with 4% NBP and rinsed with PBS and incubated with DAPI for 5 min. The slides were then rinsed with PBS and glass cover slips were mounted on glass slides with mounting medium, and fluorescent images were captured with confocal microscope. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. After 24 h of incubation for HT22 cells and 48 h of incubation for SH-SY5Y cells under the above-mentioned conditions, the culture medium was removed and replaced with medium containing nano-alumina at three different concentrations (100, 125 and 150 μg/ml). Incubation was continued for another 12 and 24 h. Control cells were cultured under similar conditions in culture medium without added nanoparticles. Mouse brain tissue was fixed by transcardial perfusion with 4% ice-cold paraformaldehyde. Post-fixation of the brains was continued for the next 72 h in 4 % paraformaldehyde and then in 20 % sucrose for 72 h at 40C. An Optimal Cutting Temperature (OCT) compound (A.O., USA) was used for mounting the brains followed by freezing them in liquid nitrogen and sectioning (14 μm) in the coronal plane with a CM 3050C cryostat (Leica, Germany). The tissue sections were mounted on Probe On Plus charged slides (Fisher, USA). To detect the aluminium accumulation in HT22 cells and in the brain of mice treated with nanoalumina, we performed Morin staining (3, 5, 7, 29, 49-Pentahydroxyflavone) as previously described29. The Morin solution was made by mixing 0.2% Morin reagent in 85% ethanol containing 0.5% glacial acetic acid. Slides containing HT22 cells and mouse brain tissues were immersed in a 1% aqueous solution of hydrochloric acid for 10 min, followed by washing and immersion in the Morin solution for 10 min and DAPI counterstaining for 5 min. Glass cover slips were mounted on slides with mounting medium, and images were taken using confocal laser scanning microscope (FV 1000, Olympus, Japan). Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. Procedure for Fluoro-Jade B (FJB) staining was followed as we reported earlier30. Overnight airdried brain tissue slides were immersed in a solution of 1% sodium hydroxide and 80% ethanol for 5 min, followed by immersion in 70% ethanol for 2 min, distilled water for 2 min, and 0.06% potassium permanganate solution for 10 min and a rinse with distilled water. Subsequently, the slides were immersed in a 0.01% FJB solution for 20 min, rinsed with distilled water and were allowed to dry for 10 min and immersed in DAPI solution for 10 min. Glass cover slips were mounted on the slides using mounting medium. A FITC filter on a confocal laser scanning microscope (FV 1000, Olympus, Japan) was used for capturing images. Amyloid plaques were investigated in the brain sections using Thioflavin S staining. Slides containing brain sections were washed twice in 0.01 M PBS for 10 min, then immersed in a fresh solution of 1 % Thioflavin S for 10 min at room temperature. Sections were then immersed in 70 % ethanol for 5 min, rinsed 2 times in water, counterstained with PI and covered with a cover slip. Both CycLex AMPK kinase (CycLex Co., Nagano, Japan) and Aβ1-42 assay (Invitrogen; Camarillo, CA) kits were used to measure the activated AMPK and Aβ-1–42 levels in both nanoalumina treated and untreated mice brain homogenates according to manufacturer's instructions. The original X-ray films of the western blots were scanned, and their densities analysed by using computer based Sigma Gel System (SPSS Inc.) software. Densities were normalized to the untreated control and all the experimental results in vitro repeated in triplicate were expressed as the mean ± SEM. Morphological images were analysed for their IOD (Integrated Densities) with Image J software. Prism 5 (Graph Pad Software, Inc., San Diego, California) was used for one way analysis of variance (ANOVA) followed by Student's t-test. Data were considered significant at p< 0.05, 0.01 and 0.001. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. The nano-alumina particle size was analyzed by transmission electron microscope (TEM). These nano particles are mixture of various sizes. The TEM images shown in Fig. 1 reveal that these particles were of oval shaped and of various sizes ranging from 30-60 nm. First, we analysed the toxic effects of nano-alumina in vitro. To investigate whether nanoalumina in three concentrations (100, 125 and 150 μg/ml) was toxic to human neuroblastoma SH-SY5Y neuronal cells in vitro, the Apo-Tox GloTM Triplex assay was performed. This assay consisted of three steps including 1). Cell viability, 2). Cytotoxicity and 3). Apoptosis (apoptotic marker caspase-3/7). The cells were exposed for 24 h to 100, 125 and 150 μg/ml of nanoalumina, and all concentrations caused a significant reduction in the number of viable SH-SY5Y cells, induced cell toxicity and increased the activation of caspase-3/7 significantly compared to the control (Fig. 2A). To further analyse the nano-alumina toxicity regarding oxidative stress, 100, 125 and 150 μg/ml of nano-alumina were investigated for producing reactive oxygen species (ROS). The results indicated that in contrast to untreated control cells, the three concentrations of nano-alumina significantly produced ROS in both SH-SY5Y and HT22 cells after exposure for 6 h (Fig. 2B). The uptakes of nano-alumina by mouse bEnd3 and SH-SY5Y cells were conducted by using confocal microscopic analyses. We investigated the intracellular retention, of rhodamine-123 conjugated nano-alumina in cultured mouse bEnd3 cells and coumarin-6-conjugated nanoalumina in cultured SH-SY5Y cells. Our results showed that both rhodamine-conjugated nanoalumina and coumarin-conjugated nano-alumina were efficiently internalized by mouse bEnd3 cells (Fig. 2C) and SH-SY5Y (Fig. 2D). These results indicate that nano-alumina can cross BBB and reach the brain and hence inducing its toxic effects to neurons after 24 h. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. To analyse whether HT22 cells can take up nano-alumina and increase their aluminium level, Morin staining was performed. Exposure of HT22 cells to nano-alumina caused an increased uptake of nano-alumina, which ultimately increased the aluminium abundance as evident from Fig. 2C. Furthermore, to assess oxidative stress induced by nano-alumina in HT22 cells, 8-Oxoguanine (8-OxoG) staining was performed using an anti-8-OxoG monoclonal antibody. The immunofluorescence images show that nano-alumina induced oxidative stress and produced a significantly high number of ROS in the treated cells in contrast to untreated HT22 cells (Fig. 2D). In the second phase, nano-alumina was peripherally administered for three weeks to ICR female mice and the mouse brain tissue was evaluated for the abundance of aluminium. The results indicate that exogenously administered nano-alumina significantly increased brain aluminium abundance compared to untreated control mice. These immunohistological evaluations for abundance of brain aluminium were conducted in the hippocampal CA1, CA3 and DG and cortical regions of the female mouse brain (Fig. 3A). Similarly, to analyse the extent of oxidative stress induced by nano-alumina, in vivo 8-OxoG staining was performed. It was evident from the immunostaining images that nano-alumina induced oxidative stress by increasing 8-OxoG expression in the brains of exposed mice. This trend was mainly observed in different parts of the hippocampus including cornu ammonis 1 and 3 (CA1 and CA3 respectively) and dentate gyrus (DG) and the cortical regions in treated mice, while no or only few 8-OxoG appearances were seen in the hippocampus and cortical regions in the untreated control mice (Fig. 3B). Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. After analysing the toxic effects of nano-alumina in vitro and brain aluminium abundance in vivo, we investigated the adverse effects of these nano-particles on behaviour and memory in female ICR mice using the Morris Water Maze (MWM) task. We analysed the escape latency time of these mice to measure spatial learning and memory before and after the injection of nano-alumina. As shown in Fig. 4A both treated and untreated mice had a similar escape latency time in finding the submerged platform, and there were no significant differences before nanoalumina administration. However, after nano-alumina exposure for three weeks, the mice showed significant memory impairment and a higher escape latency compared to a littermate control assessed on the first day. Furthermore, their escape latency in finding the submerged platform gradually decreased on day two and three, respectively, because there was no treatment during those days, but it was still higher than in untreated mice. Interestingly, when nano-alumina was injected on day four, the escape latency once again rose sharply compared to the untreated control, indicating that nano-alumina were involved in spatial memory impairment (Fig. 4B). After escape latency analysis probe test was performed by removing the platform. In the probe test, the time spent in the target quadrant by nano-alumina treated mice was significantly lower as compared to untreated control, indicating that these nanoparticles treatment partially impaired spatial learning and memory deficit in ICR female mice (Fig. 4C). We extended our study further to investigate Aβ production via the amyloidogenic pathway as a consequence of nano-alumina treatment in mice. The western blot results show that the administration nanoparticles to mice enhanced the amyloidogenic pathway of Aβ production. Nano-alumina upregulated the expression of the amyloid precursor protein (APP) and βsecretase BACE1 (beta-site amyloid precursor protein cleaving enzyme 1) activity, which significantly increased the generation of Aβ in treated mice compared to the untreated control. Moreover, it also caused downregulation of the α-secratase enzyme sAPP-α, which is responsible for the generation of non-toxic Aβ peptides through a non-amyloidogenic pathway (Fig. 4D). The levels of soluble Aβ1-42 in the brain homogenates was measured through ELISA method which revealed that nano-alumina significantly increased the production of Aβ1-42 (Fig. 4E). Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. The effect of nano-alumina on brain energy metabolism was evaluated in treated and untreated mouse brain homogenates via western blot. These results indicated that nano-alumina reduced the expression of AMPK by inactivating and dephosphorylating it at Thr172 in the brain compared to untreated mice, while the total AMPK level remain unchanged (Fig. 6A). Similarly, the AMPK activity was also reduced in the brain homogenates of nano-alumina treated mice analysed through the Cyclex AMPK activity assay method (Fig. 6B). Additionally, the expression of p-AMPK was investigated via immunofluorescence in the hippocampal CA1 and cortical regions of nano alumina treated and untreated mice. These images of immunofluorescence revealed that nano-alumina significantly inhibited the expression of p-AMPK, which supported our western blot results, suggesting that these nanoparticles are involved in the disturbance of brain energy metabolism (Fig. 6C). Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. The toxic effect of administered nano-alumina in inducing neurodegeneration was further examined by analysing the expression of apoptotic markers via western blot. Our results indicated that nano-alumina significantly upregulated the expression of various apoptotic markers, such as cleaved caspase-3 and cleaved PARP-1, in the hippocampus and cortical sections of the mouse brain, as shown in the western blot results in Fig. 7A. However, caspase-3 and PARP-1 were less expressed in untreated mice. Additionally, Flouro Jade B (FJB) staining was performed to investigate the toxicity of administered nano-alumina for inducing neurodegeneration in the hippocampus and cortical regions in mice. Investigation of the cortical and hippocampal CA1, CA3 and DG regions of nano-alumina-treated mice reveals that these nanoparticles induced neuronal cell death, as was evident from the number of positive FJB cells, whereas no such signals were observed in the control mouse brain sections (Fig. 7B). Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. This study demonstrated for first time the deleterious effects of nano-alumina exposure to a human neuroblastoma and mouse neuronal cell line in vitro and to a female mouse brain in vivo. These findings highlight that nano-alumina are responsible for the production of ROS and are very toxic to both human and mouse neuronal cells. Here we have demonstrated that the administration of nanosized alumina increased the abundance of metallic aluminium in the mouse brain, which indicated that this might be the initial stage inducing deleterious effects because mounting evidence has suggested that aluminium accumulation has severe toxic manifestations in the central nervous system. We used an ICR mice strain that is widely used in the neurosciences field, especially in the Morris water Maze (MWM). Indeed, earlier report31 demonstrates that there are sex differences between male and female mice in this strain, and their results indicate a poor MWM behaviour performance for male mice compared to female mice. Additionally, another report5 have shown that nanosized alumina induce behaviour impairment in ICR male mice. Additionally, various animal studies have shown that aluminium exposure causes aluminium accumulation in all regions of rat brain following chronic exposure, the maximum amount is seen in hippocampus, which is the site of memory and learning32, 33. A similar finding in previous studies also demonstrated that aluminium is responsible for inducing neuropathological, neurobehavioral, neurophysical and neurochemical alterations that ultimately result in memory impairment34, 35. Moreover, there is close association between high concentrations of aluminium in drinking water and AD36, but the exact mechanism is still unexplained. In this study, we demonstrated for the first time that nanoalumina induced AD-like pathology and not only enhanced toxic Aβ production via the amyloidogenic pathway in mice but also its accumulation and aggregation in the form of Aβ plaques. Along with increased Aβ production, nano-alumina caused the upregulation of the hyperphosphorylation of microtubule-associated tau proteins in treated mice, suggesting that ptau production and accumulation are Aβ-dependent. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. The literature indicates that Aβ interacts with signalling molecules that are involved in the hyperphosphorylation of microtubule-associated tau proteins37, 38. Hyperphosphorylated tau accumulation causes neurofibrillary tangles (NFT) that are rated to a highly toxic form of soluble tau. The accumulation of these proteins, including Aβ and p-tau, affects normal communications among neurons, causing the synapse disturbance that is an AD hallmark. Apart from senile plaques and neurofibrillary tangles, AD brains also show synaptic dysfunction, activated microglia, gliosis, activation of inflammatory markers, ROS production and neurodegeneration39 . Consistent with the previous studies, this work has noted that nano-alumina caused reduced expression of pre- and post-synapse related proteins, including synaptophysin and PSD 95, and affected their communication, inducing synaptic dysfunction in mice. Similarly, our results indicate that nano-alumina exposure induced oxidative stress and produced ROS in cells and in mice, and their accumulation and aggregation in the brain caused some serious CNS disorders, particularly AD. Other studies performed to analyse the toxicity of nano-alumina have demonstrated that these nanomaterials induce oxidative stress and inflammation40, 10. In this regard, the work of Li9 , which reported that these nanoparticles induced the activation of the glia in the brain even when peripherally administered, is very important. As the powerhouse of the cells, the mitochondria play a major role in energy metabolism, and it has been previously reported that aluminium exposure disrupts the normal functioning of mitochondria in neuronal cells. In this study, nanosized alumina disrupted energy metabolism in mice by deactivating and dephosphorylating AMPK at Thr172. These findings are similar to an earlier study that showed a decrease in ATP production in human hepatocytes after treatment with aluminium53. Our results suggest that ROS production and the deactivation of AMPK by nano-alumina are early events in the pathogenesis of AD. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. In conclusion, the present study postulates that nano-alumina are responsible for inducing toxic effects and decreasing cell viability by producing ROS in vitro, which can reach the brain and accumulate in exposed animals, inducing oxidative stress and neurodegeneration. Most importantly, our findings indicate that nano-alumina exposure make the CNS more vulnerable and accelerates multiple pathways, which may increase the chances of neurodegenerative disorders, such as AD. The authors declare no conflict of interest. This research was supported by the Pioneer Research Center Program through the National Research Foundation of Korea funded by the Ministry of Science, ICT & Future Planning (2012- 0009521). This manuscript is designed and written by Dr. Shahid Ali Shah. Dr. Shah also performed all the western blots, immunofluorescence and ELISA experiments described here. Dr. Ashfaq Ahmad, Mr. Faheem Ullah and Dr. Faizul Amin conducted the cellular uptake experiments. The results are compiled by Mr. Gwang Ho Yoon. We are very thankful to Prof. Myeong OK Kim (PhD) for her kind attitude and giving us an opportunity as she is the corresponding author and holds all the responsibilities related to this manuscript. Nanoscale Accepted Manuscript Published on 11 August 2015. Downloaded by Monash University on 25/08/2015 12:46:48. 53. R. Mailloux, J. Lemire and V. Appanna, Aluminum-induced mitochondrial dysfunction leads to lipid accumulation in human hepatocytes: a link to obesity. Cell Physiol. Biochem., 2007, 20, 627-638. Figure 1. Shown are the images taken through TEM of various sizes and shaped of nanoalumina. The particle size of nano-alumina in water suspension was in 30-60 nm range. Figure 7. Nano-alumina upregulated the expression of different apoptotic markers and induced neurodegeneration in the mice brain. The representative immunoblots of various (A) apoptotic markers, including cleaved caspase-3 and cleaved PARP-1, in the nano-alumina treated and control mice. The histograms containing their density details are also given here. (B) Representation of the green colour Fluoro Jade B (FJB) positive neuronal cells counterstained with DAPI (blue) in the cortical and hippocampal CA1, CA3, DG regions of the female ICR mice brain after exposing to nano-alumina for three weeks. FJB details were discussed in the materials and methods section. These images are representative of the staining obtained in the sections (n=5 animals/group). Significance = , *P < 0.01, 0.001. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 48.0, \"mgL\": null, \"viab\": 57.90178}, {\"h\": 48.0, \"mgL\": null, \"viab\": 46.724808}, {\"h\": 48.0, \"mgL\": null, \"viab\": 38.63421}]}", "role": "assistant" } ]	157.json
[ { "content": "Extract sample context and measurement list. <doc> Laura Braydich-Stolle,* Saber Hussain,† John J. Schlager,† and Marie-Claude Hofmann,1 Department of Biology, The University of Dayton, Dayton, Ohio 45469; †Applied Technology, Air Force Research Laboratory, Wright-Patterson Air Force Base, Dayton, Ohio 45433 Received May 13, 2005; accepted June 28, 2005 Gametogenesis is a complex biological process that is particularly sensitive to environmental insults such as chemicals. Many chemicals have a negative impact on the germline, either by directly affecting the germ cells, or indirectly through their action on the somatic nursing cells. Ultimately, these effects can inhibit fertility, and they may have negative consequences for the development of the offspring. Recently, nanomaterials such as nanotubes, nanowires, fullerene derivatives (buckyballs), and quantum dots have received enormous national attention in the creation of new types of analytical tools for biotechnology and the life sciences. Despite the wide application of nanomaterials, there is a serious lack of information concerning their impact on human health and the environment. Thus, there are limited studies available on toxicity of nanoparticles for risk assessment of nanomaterials. The purpose of this study was to assess the suitability of a mouse spermatogonial stem cell line as a model to assess nanotoxicity in the male germline in vitro. The effects of different types of nanoparticles on these cells were evaluated by light microscopy, and by cell proliferation and standard cytotoxicity assays. Our results demonstrate a concentration-dependent toxicity for all types of particles tested, whereas the corresponding soluble salts had no significant effect. Silver nanoparticles were the most toxic while molybdenum trioxide (MoO3) nanoparticles were the least toxic. Our results suggest that this cell line provides a valuable model with which to assess the cytotoxicity of nanoparticles in the germ line in vitro. Key Words: nanoparticles; toxicity; cell line; spermatogonia; stem cells. Nanotechnology involves the creation and manipulation of materials at the nanoscale level to create unique products that exploit novel properties. Recently, nanomaterials such as nanotubes, nanowires, fullerene derivatives (buckyballs), and quantum dots have received enormous national attention in the creation of new types of analytical tools for biotechnology and the life sciences (De Wild et al., 2003). Bionanomaterials, which are by definition in the 1–100 nm range, have been used to create materials that have novel physical/chemical properties and functions based on their advantageous, miniscule size. In particular, nanoparticles are now used to target synthetic peptides, proteins, oligonucleotides, and plasmids to specific cell type while protecting these macromolecules from enzymatic degradation (Chavany et al., 1994; Janes et al., 2001). In addition, nanoparticles have been proposed for the treatment of many diseases that need constant drug concentration in the blood or drug targeting to specific cells or organs (Moghimi et al., 2001; Panyam and Labhasetwar, 2003). In this respect, nanoencapsulated therapeutic agents such as antineoplastic drugs have been used with the aim of selectively targeting antitumor agents and obtaining higher drug concentration at the tumor site (Chawla and Amiji, 2002; Sahoo et al., 2004). This achievement appears to be important because many antineoplastic agents have several adverse side effects. Nanoparticles can be used to treat diseases that require a sustained presence of the drug at several anatomical sites (Panyam and Labhasetwar, 2003; Li et al., 2004). In addition, nanomaterials are of interest to defense and engineering programs because of their potential use in electronics, sensors, munitions, and energetic/reactive systems involved in the advancement of propulsion technology (Ringer and Ratinac, 2004). If formulated properly with other materials, nanomaterials may provide greater stability and efficiency for propellant systems. Despite the wide application of nanomaterials, there is a serious lack of information concerning the impact of manufactured nanomaterials on human health and the environment. Typically, after systemic administration, the nanoparticles are small enough to penetrate even very small capillaries throughout the body, and therefore they offer the most effective approach to distribution in certain tissues. Because nanoparticles can pass through biological membranes, they can affect the physiology of any cell in an animal body (Brooking et al., 2001). This consideration is of importance for stem cells, where the effects of nanoparticles on their potential for The Author 2005. Published by Oxford University Press on behalf of the Society of Toxicology. All rights reserved. For Permissions, please email: self-renewal and differentiation is unknown. Data available from toxicity studies of nanoparticles, in particular in adult stem cells, are limited. Gametogenesis is a complex biological process that is sensitive to environmental insult, for example, from chemicals (Adler, 2000; Iona et al., 2002). Chemical effects on germ cells and their maturation can inhibit fertility, cause cancer, and may have negative effects on the development of offspring. Mutagens, for example, produce heritable gene mutations, and heritable structural and numerical chromosome aberrations in germ cells (Allen et al., 1986; Tilly, 1998). The consequences of germ cell mutation for subsequent generations include the following: genetically determined phenotypic alterations without signs of illness; reduction in fertility; embryonic or perinatal death; congenital malformations with varying degrees of severity; and genetic diseases with varying degrees of health impairment (Brinkworth, 2000). Recent studies have shown that intravenous and/or intra-abdominal administration of nanoparticles to mice results in their accumulation in the cells of many tissues, including the brain and the testis, suggesting that they easily pass through the blood–brain and blood–testis barriers (Borm and Kreyling, 2004; Chen et al., 2003). The purpose of this study was to assess the suitability of a spermatogonial stem cell line as a model for the assessment of nanotoxicity in the male germ line in vitro. The effects of different types of nanoparticles on these cells was evaluated using light microscopy, cell proliferation and standard cytotoxicity assays. Our results suggest that this cell line provides a sensitive model with which to assess the cytotoxicity of nanoparticles in the germline. Furthermore, the relative toxicity of several types of widely used nanoparticles is discussed. The C18–4 cell line was previously established from type A spermatogonia isolated from 6-day-old mouse testes (Hofmann et al., 2005a). The cells were immortalized using the large T antigen gene and exhibit a phenotype characteristic of germline stem cells in vivo, such as the expression of the Ret and GFRa-1 surface receptors and of the nuclear factor Oct-4 (de Rooij and Russell, 2000; Pesce et al., 1998; Hofmann et al., 2005b). The cells are adherent and respond to the growth factor glial cell line–derived neurotrophic factor (GDNF) (Meng et al., 2000) by increasing their rate of proliferation (Hofmann et al., 2005a). The cells were maintained in D-MEM medium supplemented with 10% fetal calf serum (FCS), 1 mM sodium pyruvate, 2 mM glutamine, 50 U/ml penicillin, 50 mg/ml streptomycin, and 100 mM non-essential amino acids. All tissue culture reagents were purchased from Atlanta Biologicals, (Norcross, GA) or Fisher Scientific (Pittsburgh, PA). For morphological studies, the cells were seeded in 6-well plates at a concentration of 3 3 105 cells/well in 2 ml of complete medium. For the MTS and LDH assays, the cells were seeded onto 96-well plates at a density of 1 3 104 cells per well. The cells were incubated in a humidified incubator at 33-C and 5% CO2 atmosphere. They were cultured for 2 days (60% confluency) before the assays. For the apoptosis/necrosis assay, the cells were seeded onto 6-well plates at a density of 1 3 105 cells/well and cultured for 2 days (80% confluency) before the assays. Silver (Ag—15 nm), molybdenum (MoO3 —30 nm), and aluminum (Al— 30 nm) nanoparticles were obtained from the Air Force Research Laboratory, Brooks AFB, TX. The nanoparticles were made in a commercial pulsed-plasma reactor, which forms the particles in a gas phase process. As a positive control for toxicity, we used cadmium oxide, a large-sized material of ~1000 nm known for its cytotoxic properties (Goering et al., 2000) (Fluka Chemicals/Sigma, St Louis, MO). Silver, molybdenum, and aluminum nanoparticles were first dispersed in phosphate buffered saline (PBS) and used at final concentrations of 5, 10, 25, 50, and 100 lg/ml culture medium. Cadmium oxide was also dispersed in PBS and used at final concentrations of 1, 2, 5, 10, and 25 lg/ml. For comparison, we also assessed the effect of soluble species such as cadmium chloride, silver carbonate, aluminum chloride, and sodium molybdate. Those chemicals were obtained from Sigma, St Louis, MO. When cells reached 60% confluency, the different types of nanoparticles were added to the cells at different concentrations. After 48 h incubation, cell morphology was assessed with a Nikon Eclipse TS-100 phase-contrast microscope. LDH leakage. For the lactase dehydrogenase (LDH) leakage assay, we used the CytoTox 96 Non-Radioactive Cytotoxicity Assay of Promega (Madison, WI). This is a colorimetric assay that quantitatively measures LDH, a stable cytosolic enzyme that is released upon cell lysis. Released LDH in culture supernatants is measured with a 30-min coupled enzymatic assay that results in the conversion of a tetrazolium salt (INT) into a red formazan product. The amount of color formed is proportional to the number of lysed cells. Fourty eight hours after seeding, the cells were treated with the different nanoparticles at different concentrations, and incubated for another 48 h. Then, 50 ll of cell culture medium was collected from each well, diluted 1:1 with fresh medium, and plated into a new microtiter plate. Next, 50 ll of substrate solution was added to the wells, and the plates were incubated for 30 min at room temperature. Absorbance at 490 nm was measured with a standard microplate reader (Victor3 multilabel counter, PerkinElmer, Boston, MA). Each experiment was done in triplicate. The spectrophotometer was calibrated to zero absorbance using culture medium without cells. The relative LDH leakage (%) related to control wells containing cell culture medium without nanoparticles or PBS as a vehicle was calculated by [A]test/[A]control 3 100. Where [A]test is the absorbance of the test sample and [A]control is the absorbance of the control sample. Apoptosis/necrosis assay. To determine cellular apoptosis or necrosis, we used the Vybrant Apoptosis Assay Kit #4 (Molecular Probes, Eugene, OR). Kit #4 detects changes in cell membrane permeability with YO-PRO-1 dye, a green-fluorescent nucleic acid stain that is permeant to apoptotic cells but not to live cells. Necrotic cells are labeled with red-fluorescent propidium iodide. Cells were cultured with sublethal concentrations of nanoparticles for 48 h. They were then washed with PBS and incubated with 2 ll of YO-PRO-1 dye and 2 ll of propidium iodide in 2 ml of PBS per well, and further incubated for 30 min. After three washes, the cultures were covered with 1 ml of PBS, and the numbers of apoptotic and necrotic cells were immediately evaluated with a phase-contrast fluorescence microscope. Results were expressed as percent control cells cultured without nanoparticles. The general morphology of the C18–4 cells incubated with nanoparticles in phase-contrast microscopy is shown in Figure 1. The figure shows that the cells were well spread, and there was no distinct change in morphology after 48 h of incubation with any concentration of molybdenum nanoparticles relative to control cells (Fig. 1A and Fig. 1E). However, dramatic changes occur with cadmium oxide (Fig. 1B), which is considered as positive control for cytotoxicity in this report. Within 48 h of exposure at concentrations of 1 lg/ml CdO already, the cells shrink and become irregular. At concentrations higher than 5 lg/ml, they became necrotic and detached from the culture dishes. Figure 1C shows the dramatic changes induced by silver nanoparticles at concentrations of 10 lg/ml and above. In addition to necrotic areas, some cells retained an intact plasma membrane (arrows), indicating that apoptosis had occurred. Interestingly, aluminum nanoparticles did not induce shrinkage, necrosis, or apoptosis of the cells, at least at concentrations below 10 lg/ml. Instead, the particles accumulated in the cell cytoplasm, forming aggregates that were unable to enter the nucleus (Fig. 1D). The mitochondrial function—and by extension the viability of the germline stem cells—was measured by means of the MTS assay after culturing in presence of the nanoparticles for 48 h. As is evident from Figure 2, in general the cytotoxic effect of the nanoparticles on mitochondrial activity increases in relation to increasing concentration. Figure 2A shows that soluble cadmium chloride, a known toxicant, had no significant effect on mitochondrial activity at concentrations below 5 lg/ml. The EC50 was calculated at 21.3 lg/ml. In comparison, the cytotoxic effects of cadmium oxide were much stronger, with a significant inhibition of mitochondrial function starting at concentrations below 1 lg/ml. The EC50 was calculated at 0.5 lg/ml. Figure 2B shows that, on the one hand, silver carbonate, generally considered to be non-hazardous, had no significant cytotoxic effect on mitochondrial function and cell viability up to concentrations of 100 lg/ml. The EC50 of silver carbonate was calculated at 408 lg/ml. On the other hand, silver added to the cells as a nanoparticulate (diameter of 15 nm) reduced drastically mitochondrial function and cell viability. The toxic effect of silver nanoparticles started between 5 and 10 lg/ml, with an EC50 calculated at 8.75 lg/ml. Higher concentrations of silver nanoparticles could not be tested because of particle clumping and precipitation above 10 lg/ml. Figure 2C compares the influence of soluble and nanoparticulate molybdenum species on germline stem cell mitochondrial function and viability. In general, sodium molybdate is known to be moderately toxic for humans and other animal species. In the case of the C18–4 germline stem cells, the mitochondrial function and percentage viability of the germ cells decreased slightly at concentrations of 50 lg/ml and above, but the difference was not statistically different from the results obtained with lower concentrations. The EC50 of sodium molybdate was calculated at 322 lg/ml. In contrast, a significant decrease in mitochondrial function was seen when molybdenum was used as a nanoparticulate (30 nm in diameter). Our data indicate that in these cells, molybdenum nanoparticles exert toxic effects on cellular metabolic activity at concentrations of 50 lg/ml and above. The EC50 of molybdenum nanoparticles is 90 lg/ml. The effects of aluminum nanoparticles on mitochondrial function could not be assessed because the particles accumulated in the cells and formed cytoplasmic aggregates at low concentrations (Fig. 1D), and the light-scattering effect of the particles interfered with the spectrophotometric readings. Figure 3A shows that soluble cadmium chloride has no effect on the plasma membrane at any of the concentrations tested. However, a dose-dependent increase in LDH leakage was observed with cadmium oxide. The EC50 of cadmium oxide was calculated at 2.5 lg/ml. In comparison, 48 h of treatment of the C18–4 germline stem cells with nanoparticles produced a dose-dependent increase of LDH leakage at lower concentrations only. In general, LDH leakage reached a plateau at higher concentrations. Figure 3B shows that soluble silver carbonate does not affect the plasma membrane, which was expected. A slight increase in LDH leakage was observed with silver nanoparticles, indicating that these particles interfere with cell metabolism rather than disrupting the plasma membrane. Thus, they might promote cell apoptosis rather than necrosis (Fig. 1). The EC50 of silver nanoparticles was evaluated at 2.5 lg/ml. Higher concentrations of silver nanoparticles were not assessed because of particle clumping and precipitation at levels above 10 lg/ml. Figure 3C indicates that aluminum chloride did not affect the integrity of the plasma membrane at any of the concentrations tested. In contrast, a statistically significant increase of LDH leakage was observed with aluminum nanoparticles. Leakage is dosedependent at lower concentrations, and the values reach a plateau at around 25 lg/ml nanoparticles. The EC50 was measured at 4.7 lg/ml. Figure 3D shows the effects of sodium molybdate and molybdenum in nanoparticulate form on the C18–4 cells. Soluble sodium molybdate did not affect the integrity of the plasma membrane, whereas a significant increase of LDH leakage was observed with molybdenum nanoparticles. The leakage was dose-dependent at low concentrations until the values reached a plateau at 10 lg/ml nanoparticles. The EC50 was measured at 5 lg/ml. Because the cellular metabolic activity seemed affected by the nanoparticles but the plasma membrane remained relatively intact, we assessed the possibility of apoptosis caused by the nanoparticles, especially at lower concentrations. Using the Vybrant apopotosis assay that measures the number of cells in apoptosis versus necrosis, we confrmed that apopotosis took place at low nanoparticle concentrations. Figure 4A shows an increased number of apoptotic cells, and that increase is dosedependent at lower concentrations (1–5 lg/ml for cadmium oxide, and 10–50 lg/ml for the nanoparticulates tested). More cells became necrotic as the concentrations increased. In the case of molybdenum nanoparticles, a small number of apoptotic cells began to be observed starting at a concentration above 25 lg/ml, and few necrotic cells were observed at concentrations below 50 lg/ml. Figure 4B and 4C illustrates apoptosis in the C18–4 cells treated with molybdenum nanoparticles at a concentration of 50 lg/ml. Most cells looked intact in phase-contrast microscopy (Fig. 4B). However, these same cells exhibited positive nuclei in fluorescence microscopy when the YO-PRO-1 dye was used (Fig. 4C), but they were negative for propidium iodide that can penetrate only necrotic cells. Despite the many benefits of nanotechnology, some studies indicate that certain nanoparticles may cause adverse effects because of their small size and unique properties (Service, 2003; Hoet et al., 2004). Indeed, their size makes them highly mobile in both the human body and the environment. Nanomaterials can enter human tissues through several ports via the lungs after inhalation (Oberdorster, 2001), through the digestive system (Jani et al., 1990), and possibly through the skin (Kreilgaard, 2002; Lademann et al., 1999). Systemic distribution of nanoparticles has been demonstrated after inhalation and oral uptake (Jani et al., 1990; Oberdorster et al., 2002), and nanoparticles have been found to cross the blood–brain barrier, reaching the olfactory bulb and the cerebellum (Borm and Kreyling, 2004; Oberdorster et al., 2004). Chen and colleagues also reported that nanoparticles can penetrate the blood-testis barrier (Chen et al., 2003). Although organ- or cell-specific drug delivery through nanoparticles is a promising area of medicine, and nanoparticles might be used some day as sensors for intracellular mechanisms, few toxicology studies are available. Many of the artificially manufactured nanoparticles are made of nonbiodegradable pollutants, such as carbon black and metals, and the long-term behavior of such substances is not known. Toxicants that impair normal reproductive functions are an important public health issue. A decrease in semen quality of approximately 2% per year over the preceding 50 years has been reported for industrialized countries (Carlsen et al., 1992). It has been hypothesized that exposure to toxic chemicals is an important cause of the decline, although great regional differences exist for the same level of environmental contamination. Nevertheless, studies have shown that high exposure of men to various chemicals in certain occupational settings resulted in lower semen quality. For example, dibromochloropropane (DBCP), a nemotocide, resulted in lower sperm counts because of the destruction of undifferentiated spermatogonia (Potashnik et al., 1978). Therefore, it has now become critical to understand the molecular mechanisms leading to reproductive toxicity. Several in vivo animal models have been used to assess the testicular toxicity of many compounds. However, these models necessitate the sacrifice of animals at the end of experimentation, and they do not allow the manipulation or dissection of intracellular pathways to elucidate mechanisms of toxicity at the molecular level. In vitro model alternatives have been established, and some of them have tried to reproduce in the petri dish the complex cell–cell interactions that take place between the different germ cells and Sertoli cells (Hadley et al., 1985; Yu et al., 2005). However, these models are limited by the poor viability of the freshly isolated germ cells. In this study, we used a cell line with spermatogonial stem cell characteristics to evaluate the toxicity of different types of nanoparticles on the germline. We used three parameters widely used in toxicological studies, such as the ability of mitochondria to reduce MTS, the integrity of the plasma membrane, and the activation of apoptotic pathways. Our results indicate that the C18–4 cells provide a suitable test system for cytotoxicity in the germline. The MTS and LDH assays can be used for rank ordering of chemical and nanoparticle toxicity on mitochondrial function and plasma membrane integrity. Cadmium is a recognized toxicant that has been classified as a probable human carcinogen. It is a heavy metal that has the potential to cause lysosomal damage and DNA breakage in mammalian hepatocytes (Fotakis et al., 2005) and many other cells and tissues (Satoh et al., 2002). Cadmium also disrupts mitochondrial function both in vivo (Belyaeva et al., 2002) and in vitro (Pourahmad and O'Brien, 2000), and promotes apoptosis (Pulido and Parrish, 2003). In the testis, cadmium induces lysosomal damage in testicular Sertoli cells (Boscolo et al., 1985), but its main toxic effects appear in germ cells. In male rats subcutaneously injected with 0.6 mg cadmium chloride/kg body weight, histological examination of the testes revealed an accumulation of cadmium only in spermatogonia and spermatocytes, but not in somatic cells (Aoyagi et al., 2002). Subsequently, a decrease in the number of spermatogonia in relation to the time of exposure was observed, followed by a decrease in the number of spermatocytes and, ultimately, sperm cells. In humans, male infertility is strongly linked to cadmium exposure, but is rather due to a failure of the acrosomal reaction in sperm cells (Benoff et al., 2000). In our spermatogonial stem cell line, exposure of the cells to cadmium chloride induced a significant decrease in their metabolic activity. However, like hepatocytes (Fotakis et al., 2005), cadmium chloride had no effect on the integrity of the plasma membrane, as monitored by the LDH release assay. Importantly, the deleterious effects of cadmium were enhanced for cells exposed to the particulate, insoluble form of cadmium, such as cadmium oxide. These effects mimic well-documented data obtained on macrophages and other cell lines (Goering et al., 2000). In this case, not only mitochondrial function decreased drastically, but LDH was released in the cell environment as a function of cadmium oxide concentration. Because oxidative stress and lipid peroxidation have been reported after exposure to both cadmium chloride and cadmium oxide, the higher toxicity of cadmium oxide might be related to the size of the particles entering into contact with the plasma membrane. Furthermore, our study shows that the sensitivity of the C18–4 cells to cadmium oxide is comparable to the sensitivity of the BRL 3A liver cells regarding their metabolic activity (Hussain et al. in press) (Table 1). The membrane integrity of the C18–4 cells is less affected. This is corroborated by the fact that at lower concentrations cadmium oxide promotes apoptosis rather than necrosis in the C18–4 cells, thus leaving the plasma membrane intact. Because the C18–4 cells showed an increased sensitivity to the particulate form of cadmium rather than the soluble form, we next studied the effects of metal nanoparticles on these germline stem cells. As predicted, silver carbonate, which was used as a control, was not toxic as shown by the MTS and LDH leakage assays. In contrast, silver nanoparticles (15 nm) reduced mitochondrial function drastically and increased membrane leakage. Our data show that the C18–4 cells are more sensitive than the BRL 3A cells in that respect (Table 1). There are studies showing that silver nanoparticles could be used in bone cement or other implantable devices as antimicrobial agents (Alt et al., 2004), but our study shows that silver in nanoparticulate form could be toxic for the bone-lining cells and other tissues. Although molybdenum in soluble form is considered to be a mildly toxic substance, it did not significantly affect the metabolic activity or the membrane integrity of the C18–4 cells. Molybdenum as a nanoparticulate did not affect metabolic activity either, at least up to a concentration of 40 lg/ml (EC50 1⁄4 90 lg/ml). At higher concentrations (over 50 lg/ml), the molybdenum nanoparticles become significantly toxic. Interestingly, whereas the effect on mitochondrial function is mild, molybdenum nanoparticles seem to promote some plasma membrane leakage at very low concentrations (5 lg/ml and 10 lg/ml). The same pattern of toxicity is shown for aluminum nanoparticles; however, the morphology of the cells did not change, indicating that at these concentrations apoptosis still occurs. This finding was corroborated by the Vybrant assay. It is known that extreme and visible membrane damage occurs only in the late stages of apoptosis. In conclusion, we have demonstrated that the C18–4 germline stem cells are a valuable tool with which to study in vitro toxicity in the germline. The sensitivity of these cells to Ag nanoparticles is greater than that of BRL 3A liver cells, which are widely used in toxicity studies. However, in the case of cadmium and the other nanoparticles tested, the sensitivities of the two cell lines are comparable. The molecular mechanisms of nanoparticles toxicity are still poorly understood, and the availability of a cell line with which to gain an understanding of these processes in the germline is of paramount importance. We thank Jeffrey Calhoun for valuable technical assistance. The work was funded in part by the Lance Armstrong Foundation. Laura Braydich-Stolle is a Consortium Research Fellow funded by the AFRL Human Effectiveness Directorate. Conflict of interest: none declared. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 48.0, \"mgL\": null, \"viab\": 100.97087}, {\"h\": 48.0, \"mgL\": null, \"viab\": 100.97087}, {\"h\": 48.0, \"mgL\": null, \"viab\": 121.16505}, {\"h\": 48.0, \"mgL\": null, \"viab\": 61.359222}, {\"h\": 48.0, \"mgL\": null, \"viab\": 42.718445}]}", "role": "assistant" } ]	154.json
[ { "content": "Extract sample context and measurement list. <doc> Amanda M. Schrand,† Houjin Huang,† Cataleya Carlson,‡ John J. Schlager,‡ Eiji Oh sawa,§,\| Saber M. Hussain,,‡ and Liming Dai,† Department of Chemical and Materials Engineering, UniVersity of Dayton, 300 College Park, Dayton, Ohio 45469-0240, Applied Biotechnology Branch, Human EffectiVeness Directorate, Air Force Research Laboratory, Wright-Patterson AFB, Ohio 45433-5707, and NanoCarbon Research Institute, Ltd., Kashiwa-no-ha, Chiba 277-0882, Japan ReceiVed: September 28, 2006; In Final Form: NoVember 15, 2006 Finely divided carbon particles, including charcoal, lampblack, and diamond particles, have been used for ornamental and official tattoos since ancient times. With the recent development in nanoscience and nanotechnology, carbon-based nanomaterials (e.g., fullerenes, nanotubes, nanodiamonds) attract a great deal of interest. Owing to their low chemical reactivity and unique physical properties, nanodiamonds could be useful in a variety of biological applications such as carriers for drugs, genes, or proteins; novel imaging techniques; coatings for implantable materials; and biosensors and biomedical nanorobots. Therefore, it is essential to ascertain the possible hazards of nanodiamonds to humans and other biological systems. We have, for the first time, assessed the cytotoxicity of nanodiamonds ranging in size from 2 to 10 nm. Assays of cell viability such as mitochondrial function (MTT) and luminescent ATP production showed that nanodiamonds were not toxic to a variety of cell types. Furthermore, nanodiamonds did not produce significant reactive oxygen species. Cells can grow on nanodiamond-coated substrates without morphological changes compared to controls. These results suggest that nanodiamonds could be ideal for many biological applications in a diverse range of cell types. With the recent development in nanoscience and nanotechnology, carbon-based nanomaterials (e.g., fullerenes, nanotubes, nanodiamonds) are receiving much attention due to their remarkable mechanical, electrical, and thermal properties.1-3 The importance of carbon nanomaterials in biological applications has been recently recognized. Examples include their potential uses in drug, gene, and protein delivery; novel imaging methods, coatings for implantable/corrosion resistant materials; biosensors/biochips; purification of proteins; medical nanorobots; and many other emerging biomedical technologies.4-9 In particular, carbon nanomaterials have recently been examined with both in vivo studies10-11 and in vitro cell cultures.12-18 Although the scientific community has been so far primarily focused on the potential biological applications of fullerenes and/or carbon nanotubes, other carbon nanomaterials (especially nanodiamonds) are beginning to emerge as alternative candidates for similar and many other applications. Both carbon nanotubes and nanodiamonds can be similarly modified for nanocomposite and biological applications.19 It is envisaged that nanodiamonds may prove to be an even better drug carrier, imaging probe, or implant coating in biological systems compared to currently used nanomaterials due to its optical transparency, chemical inertness, high specific area, and hardness.4,20 Recent progresses in the dispersion of detonation nanodiamonds (NDs, 2-10 nm) in aqueous media made by Oh sawa and co-workers has facilitated the use of NDs in physiological solutions,21 whereas most previous studies have focused on polycrystalline chemical vapor deposited (CVD) diamond films for biomedical applications.6,7,22 It has been recognized that a bulk material with good biocompatibility may not be as well-tolerated by the body once it is in a fine particulate or nanometer-sized form. Although the CVD diamond thin films are generally regarded as biologically inert, noninflammatory, and biocompatible, are diamond nanoparticles also well tolerated by cells? The availability of the newly produced detonation NDs in aqueous dispersed forms facilitates their possible use in nanomedicine (e.g., medical nanorobots made of NDs) and biorelated studies.3 However, the environmental impact of the detonation NDs, especially on humans and other biological systems, has not been properly studied. In one recent study, Yu et al. investigated the biocompatibility of relatively large synthetic abrasive diamond powders (100 nm) in cell culture and found very low cytotoxicity in kidney cells.5 These authors visualized their diamond nanoparticles with fluorescent confocal microscopy to enter the cells and localize in the cytoplasm. For these nanodiamonds to be fluorescent, however, these authors performed the tedious procedures of electron beam irradiation and annealing of the nanoparticles. At the nanometer scale, the particle properties depend strongly on the size of the material.3 As a consequence, changes in the size-dependent cytotoxicity may be observed for diamond nanoparticles of different sizes. Due to the high surface-to- volume ratio associated with nanometer-sized materials, a tremendous surface chemistry effect on the nanoparticle properties is also expected.3 Previous studies in our AFB laboratory have primarily concerned metal nanoparticle toxicity using in vitro cell culture models for chemical toxicity screening.23-28 In the present study, we report the first biocompatibility study of small NDs (2-10 nm), synthesized by the detonation of carbon-containing explosives in an inert atmosphere,21,29 and possible effects of the surface chemistry by treating the NDs with acids or bases (-COOH, -COONa, -SO3Na; see ). In conjunction with tests to determine cytotoxicity, cells were grown on ND-coated substrates to examine their interactions and sustained viability over time. Nanodiamonds were generously supplied by NanoCarbon Research Institute Ltd. and were synthesized according to previously reported detonation techniques.21,29 Nanometer-sized carbon black (CB, Cabot) and submicron-sized cadmium oxide (CdO, Fluka Chemical Co.) were used as negative and positive controls, repsectively. All nanoparticles were UV-sterilized and then diluted to a stock concentration of 1 mg/mL in deionized water. Characterization of nanoparticle size and morphology was performed with transmission electron microscopy (TEM, Hitachi H-7600 W-tip). The chemical nature of the functionalized NDs was examined with X-ray photoelectron spectroscopy (VG Microtech ESCA 2000) using monochromatic Mg KR radiation at a power of 300 W, Fourier transformed infrared spectroscopy (FTIR, Perkin-Elmer, Spectrum One) and Raman spectroscopy (Renishaw, inVia reflex microRaman, 514.5 nm laser). ND substrates were prepared by drop casting solutions of ND onto collagen or poly(L-lysine) coated glass coverslips and drying for 1 h under UV light before plating cells. Neuroblastoma cells, a neuronal phenotype, were generously provided by Dr. David Cool's laboratory at Wright State University (Dayton, OH) and other cells were purchased from the American Type Culture Collection (ATCC). All cells were plated and grown according to standard cell culture techniques.26 After a desired growth period, cell cultures were dosed with freshly prepared nanoparticle working solutions at concentrations ranging between 5 and 100 μg/mL in cell culture media without serum. pH values were controlled between 7.2 and 7.6 for both control dosing media and nanoparticles in the cell culture solutions. Fluorescent microscopy was performed to examine mitochondrial membrane permeability (Rhodamine 123, JC-1, Invitrogen and Mijt -E-Ψ membrane permeability detection kit, BioMol). Both mitochondrial dyes entered the live cells after 15-30 min at 37 °C. If the mitochondrial membranes have been damaged after incubation with nanoparticles, then the dyes will disperse in the cytoplasm due to leakage, whereas intact mitochondrial membranes will retain and aggregate the dye. Fluorescence was visualized with TRITC and FITC filters on an Olympus IX71 epifluorescent microscope. Nanoparticletreated cell samples for TEM study were fixed with glutaraldehye/paraformaldehyde, stained with osmium tetroxide, dehydrated through analytical grade ethanol, embedded in resin, cured, and thin sectioned. For scanning electron microscopy (SEM, Hitachi S-4800), cells were first fixed and dehydrated as described above, mounted to aluminum stubs with doublesided carbon adhesive tape, and then air-dried and sputter-coated with gold. The 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) assay was conducted to assess cellular viability based on mitochondrial function30 with a slight modification of removing nanoparticles with centrifugation before microplate reading (vide infra). After 30 min of incubation with MTT, a purple color developed within the cells, indicating the cleavage of the tetrazolium salt (MTT) by mitochondrial reductase in live cells. The purple product (formazan crystals) was extracted into solution for homogeneous staining and the absorbance was read on a Spectromax 190 microplate reader at 570-630 nm. The percent reduction of MTT was compared to controls (cells not exposed to nanoparticles), which represented 100% MTT reduction. The CellTiter-Glo luminescent viability assay was performed to reconfirm data from the MTT assay. This assay provides a homogeneous method for determining the number of viable cells in culture based on quantitation of adenosine 5′ triphosphate (ATP), which indicates the presence of metabolically active cells. Oxidative stress was measured in relation to the generation of reactive oxygen species (ROS). Prior to dosing cells with nanoparticles, the fluorescent probe 2′,7′-dichlorohydrofluorescein diacetate (DCHF-DA, Sigma) was applied under a light controlled environment as described by Wang and Joseph.31 After nanoparticle treatment, the fluorescent intensity from each well was measured with a 485 nm excitation filter and a 530 nm emission filter on a SpectraMAX Gemini Plus microplate reader (Molecular Device) equipped with SOFTmax Pro 3.1.2 software (Molecular Devices Corp., Sunnyvale, CA). The positive control, hydrogen peroxide (30% H2O2, Fisher Scientific), was used to assess the reactivity of the probe. Biochemical assays (MTT and ROS) were done in triplicate and the results were presented as mean ( standard deviation. All nanoparticles in this study have average sizes ranging from 2 to 10 nm for nanodiamonds (ND), 20-30 nm for nanometer-sized CB, and up to hundreds of nanometers for cadmium oxide (CdO). Characterization of nanoparticle size distributions and morphologies was accomplished with transmission electron microscopy (TEM; see ). The morphology of neuroblastoma cells was round immediately after trypsinization. Upon attachment and growth, however, some of the cells developed elongated extensions and others remained round, which is characteristic of these cells. For morphological examination of cell-nanoparticle interactions, cells were incubated with media alone (control) or media containing nanoparticles at various concentrations. After 24 h of incubation with NDs or CB nanoparticles, cells appeared similar to control cells with some cells displaying an elongated morphology (Figure 1A-D). The ND and CB were irregularly shaped particles in the surrounding media, agglomerated at cell borders, along their processes, and within the cells at concentrations ranging from 5 to 100 μg/mL (Figure 1A-D). In contrast, cells incubated with the positive control CdO (Figure 1E) lacked cellular extensions, were reduced in size, had irregular cell borders, and formed vacuoles, which are morphological indicators of toxicity. For simplicity, only images of the highest concentrations of ND-raw and ND-COOH (100 μg/mL) are included to represent unfunctionalized and acid-functionalized NDs, respectively. Base-functionalized NDs, ND-SO3Na and ND-COONa, displayed similar morphological characteristics to ND-raw and ND-COOH. Additionally, only a low concentration (2.5 μg/mL) of CdO was shown to demonstrate its strong toxicity. To further examine interactions between the cells and nanoparticles, changes in mitochondrial membrane permeability and cytoskeletal architechure were examined with fluorescent microscopy (Figure 2). The Mijt -E-Ψ fluorescent reagent, when aggregated inside healthy mitochondria, fluoresces red whereas dispersion of the dye due to mitochondrial membrane disruption causes it to fluoresce green in the cytoplasm. Aggregation and retention of the mitochondrial dye inside healthy cells was shown in control cells (Figure 2A), cells incubated with 100 μg/mL ND-raw (Figure 2B), and to a lesser extent in cells incubated with 100 μg/mL CB (Figure 2C). Cells incubated with CB had noticeable dark CB nanoparticles attached to cell borders or internalized, which tended to block the fluorescent signal in certain areas compared to cells incubated with NDs (Figure 2C). The dispersion of the dye was apparent in cells treated with 2.5 μg/mL CdO, suggesting toxicity and the initiation of apoptosis or programmed cell death (Figure 2D). Therefore, these results show that the mitochondrial membrane was clearly maintained in cells incubated with ND in support of the biocompatibility of NDs. In the case of cytoskeletal alterations after incubation with NDs, distinct branching and extension of multiple neurites was found after 24 h (Figure 2F) compared to control (Figure 2E), CB (Figure 2G), or CdO (Figure 2H) treated cells. Again, noticeable black CB nanoparticles could be seen attached to the cells (Figure 2G) or cell shrinkage due to the toxic effects of CdO (Figure 2H). Because individual NDs were too small to be resolved inside the cells with conventional light or fluorescent microscopy, the internalization of NDs into the cells after 24 h was examined with TEM. As can be seen in Figure 3A,B, NDs were found inside the cells after incubation with 25 μg/mL of ND-COOH for 24 h. Although these representative images do not specify the exact location of the ND particles, they appear to localize in the cytoplasm in aggregates approximately 500 nm in size (Figure 3A,B). Although the internalization of nanodiamonds into the cells could provide new nanotherapeutics with the nanodiamonds as biological transporters32-34 or internalized cell killers,35 as is the case of carbon nanotubes, they may be difficult to clear from the cell. The possible long-term pathologic effects of the internalized nanodiamonds on the cells and the related long-term and acute toxicology data on the animal model are still pending for further investigation. Additionally, cells were plated on ND substrates to determine if they could adhere and grow while maintaining morphologies similar to control cells. After 96 h, cells grown on the ND- COOH coated collagen substrates (Figure 3D) were viable with morphologies similar to control cells (Figure 3C), suggesting that these substrates allow cell attachment and growth. To more quantitatively determine the toxicity or biocompatibility of NDs, colorimetric (MTT), luminescent (Cell-titer Glo), and fluorescent (ROS) assays were performed. The MTT assay was selected to assess the mitochondrial function of neuroblastoma cells. Mitochondria are vulnerable targets for toxic injury by a variety of compounds because of their crucial role in maintaining cellular structure and function via aerobic ATP production.24 The reduction of MTT dye occurs only in functional mitochondria, therefore, a decrease in MTT dye reduction is an indication of damage to mitochondria. CdO exhibited strong toxicity with its capability to sharply decrease cell viability, compared to the control, leading to a strong concentration dependence over the whole concentration range from 5 to 100 μg/mL (Figure 4A). Nanometer-sized carbon black showed slightly reduced cell viability, but no significant difference from controls at concentrations up to 100 μg/mL. However, cells incubated with various concentrations of functionalized or unfunctionalized nanodiamonds had some slightly higher values, but again no significant difference in viability compared to controls at concentrations up to 100 μg/mL (Figure 4A). To further confirm the low cytotoxicity of NDs, three other cells types (macrophages, keratinocytes, and PC-12 cells) were investigated and found to display similar trends of low cytotoxicity. A comparison between neuroblastoma cells and macrophages demonstrated that NDs display no toxicity even to cells, which may have different methods of internalization (Figure 4B). To further verify the biocompatibility of nanodiamonds, we measured luminescence, corresponding to ATP production, as a marker of cell viability (see ). Similar trends for nanoparticle toxicity were found, but surprisingly, the luminescence value for ND-raw was lower than nanometersized CB, though these values were not significantly different from the control. It is worthwhile to point out here that our recent work (see ), along with others,4 has indicated that nanoparticles could interfere with various colorimetric cytotoxicity assays, such as the MTT and CellTiter-Glo luminescent viability assays, probably due to light scattering and/or direct nanoparticle interaction with the chemical components of the assays. All data in Figure 4A,B were taken after the incorporation of an additional centrifugation step to remove the nanoparticles from the plate before reading. To investigate nanoparticle-induced oxidative stress as mechanistic changes, we assessed the generation of reactive oxygen species (ROS).36 ROS are naturally generated byproducts of cellular redox/enzymatic reactions such as mitochondrial respiration, phagocytosis, and metabolism. However, they can also unregulate generation, leading to a condition known as oxidative stress, which can cause numerous pathological conditions.37 Increases in intracellular ROS (oxidative stress) represent a potentially toxic insult which, if not neutralized by antioxidant defenses (e.g., glutathione and antioxidant enzymes) could lead to membrane dysfunction, protein degradation, and DNA damage.38-42 The fluorescence intensity of dichlorofluorescein (DCF), an oxidized form of 2′,7′-dichlorofluorescein, can be used as a measure of the cumulative production of ROS over a period of nanoparticle exposure. We found that CB shows a higher level of ROS production than ND (Figure 4C). The positive control for this assay, hydrogen peroxide (H2O2), showed a dosedependent increase in ROS production (Figure 4D). The relatively low level of ROS generation produced in cells incubated with NDs is consistent with the MTT and ATP results. Therefore, these results further support the biocompatibility of NDs and suggest that ND does not induce ROS generation in this in Vitro cell model system. In summary, we have demonstrated that 2-10 nm nanodiamonds, with and without surface modification by acid or base, are biocompatible with a variety of cells of different origins, including neuroblastoma, macrophage, keratinocyte, and PC-12 cells. Several methods for assessing toxicity were used to rigorously test the cytotoxicity of the nanodiamonds (2-10 nm) using carbon black (20-30 nm) and cadmium oxide (100- 1000 nm) as negative and positive controls, respectively. Although the cell types used may have different mechanisms of internalization of the nanodiamonds and the long-term effect of the internalized nanodiamonds on the cells needs to be further investigated, the resultant retention of mitochondria membrane along with low levels of ROS suggests that once inside the cell the nanodiamonds remain nonreactive. In conjunction with the toxicity testing of nanodiamonds, cells were grown on NDcoated substrates to examine their interactions and sustained viability over time, which provided further assurance for the utility of nanodiamonds as biologically compatible materials. Acknowledgment. We thank Col. J. Riddle for his strong support and encouragement for this research. TEM work was performed at the Nanoscale Engineering Science and Technology Laboratory (NEST), University of Dayton. A.S. is funded by the Biosciences and Protection Division, Air Force Research Laboratory, under the Oak Ridge Institute for Science and Education, Oak Ridge, TN, and the Dayton Area Graduate Studies Institute (DAGSI). L.D. and E.O. thank NEDO International Cooperative Grant (2004IT081) for financial support. Available: Experimental details for nanoparticle characterization, nanoparticle interference assays, and bacterial growth in nanodiamond-containing solutions. This material is available free of charge via the Internet at http:// pubs.acs.org. (1) Baughman, R. H.; Zakhidov, A. A.; de Heer, W. A. Science 2002, 297, 787-792. (2) Zhang, M.; Fang, S.; Zakhidov, A. A.; Lee, S. B.; Aliev, A. E.; Williams, C. D.; Atkinson, K. R.; Baughman, R. H. Science 2005, 309, 1215-1219. (3) Dai, L., Ed. Carbon Nanotechnology: Recent DeVelopments in Chemistry, Physics, Materials Science and DeVice Applications; Elsevier: Amsterdam, 2006. (4) Huang, L.-C. L.; Chang, H.-C. Langmuir 2004, 20, 5879-5884. (5) Yu, S. J.; Kang, M.-W.; Chang, H.-C.; Chen, K.-M.; Yu, Y.-C. J. Am. Chem. Soc. 2005, 127, 17604-17605. (6) Yang, W.; Auciello, O.; Butler, J. E.; Cai, W.; Carlisle, J. A.; Gerbi, J. E.; Gruen, D. M.; Knickerbocker, T.; Lasseter, T. L.; Russell, Jr., J. R.; Smith, L. M.; Hamers, R. J. Nat. Mater. 2002, 1, 253-257. (7) Poh, W. C.; Loh, K. P.; Zhang, W. D.; Triparthy, S.; Ye, J.-S.; Sheu, F.-S. Langmuir 2004, 20, 5484-5492. (8) Bondar, V.; Pozdnyakova, I. O.; Puzyr, A. P. Phys. Solid State 2004, 46, 758-760. (9) Pantarotto, D.; Briand, J.-P.; Prato, M.; Bianco, A. Chem. Commun 2004, 16-17. (10) Muller, J.; Huaux, F.; Moreau, N.; Misson, P.; Heilier, J.-F; Delos, M.; Arras, M.; Fonseca, A.; Nagy, J. B.; Lison, D. Toxicol. Appl. Pharmacol. 2005, 207, 221-231. (11) Gharb, N.; Pressac, M.; Hadchouel, M.; Szwarc, H.; Wilson, S. R.; Moussa, F. Nano Lett. 2005, 5, 2578-2585. (12) Ding, L.; Stilwell, J.; Zhang, T.; Elboudwrej, O.; Jiang, H.; Selegue, J. P.; Cooke, P. A.; Gray, J. W.; Chen, F. F. Nano Lett. 2005, 5, 2448- 2464. (13) Cui, D.; Tian, F.; Ozkan, C. S.; Wang, M.; Gao, H. Toxicol. Lett. 2005, 155, 73-85. (14) Monteiro-Riviere, N. A.; Nemanich, R. J.; Inman, A. O.; Wang, Y. Y.; Riviere, J. E. Toxicol. Lett. 2005, 155, 377-384. (15) Manna, S. K.; Sarkar, S.; Barr, J.; Wise, K.; Barrera, E. V.; Jejelowo, O.; Rice-Ficht, A. C.; Ramesh, G. T. Nano Lett. 2005, 5, 1676-1684. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 14.606205}, {\"h\": 24.0, \"mgL\": null, \"viab\": 13.747017}, {\"h\": 24.0, \"mgL\": null, \"viab\": 2.5775657}, {\"h\": 24.0, \"mgL\": null, \"viab\": 2.5775657}, {\"h\": 24.0, \"mgL\": null, \"viab\": 1.1455847}]}", "role": "assistant" } ]	153.json
[ { "content": "Extract sample context and measurement list. <doc> Available online at www.sciencedirect.com jou rnal h om ep age: www.elsevier.com/locate/etap a Department of Biophysics, College of Basic Medical Sciences, Second Military Medical University, Xiangyin RD, Shanghai 200433, China b Department of Chemistry, College of Chemistry and Molecular Engineering, East China University of Science and Technology, Meilong RD, Shanghai 200237, China c Department of Physics, Henan Normal University, Henan 453007, China Article history: Received 20 September 2012 Received in revised form 26 February 2013 Accepted 8 March 2013 Available online 21 March 2013 Keywords: Nanosized titanium dioxide (nano-TiO2) Toxicity A549 cell Mitochondria The nanosized titanium dioxide (nano-TiO2) is an important nanoscale compound applied in many different fields because of its superior performance. Here, an anatase nano-TiO2 showed cytotoxicity in a dosage-dependent manner, which was in accordance with changes of A549 cell ultrastructure, A549 cell viability and intracellular ATP level. The lungs of rats treated with single intratracheal instillation of nano-TiO2 were injured, which was demonstrated by changes of alveolar epithelial cell ultrastructure, lung tissue pathology and lung tissue MDA level. The results of this study indicated that nano-TiO2 should be related to the generation of intracellular reactive oxygen species (ROS), which injured mitochondria and prevented the synthesis of ATP. The cells were approaching to apoptosis eventually. In macroscopic view, the lungs inevitably suffered. 2013 Elsevier B.V. All rights reserved. In recent years, a large number of nanomaterials with special physical and chemical properties have been synthesized and applied in many different fields, such as nanocatalysis (Vinod, 2010), nanoceramics (Hoda et al., 2012) and biomedicine (Jung and Jong, 2011). Among the various nanomaterials, nanosized titanium dioxide (nano-TiO2) was chemically inert, noncorrosive and photocatalytic, thus nano-TiO2 was widely applied in consumer products, such as pigments (Taavitsainen and Jalava, 1995), photocatalysts (Sun et al., 2004) and cosmetic products (Marta et al., 2010). However, the fact that nano-TiO2 could enter the human body through different routes such as inhalation (respiratory tract), ingestion (gastrointestinal tract), dermal penetration (skin) and injection (blood circulation) (Liu et al., 2010) was demonstrated by many experts. Lots of investigations had shown that nano-TiO2 was able to cause injuries in various cells such as human hepatocellular carcinoma cell (SMMC-7721), human liver cell (HL-7702), rat hepatocarcinoma cell (CBRH-7919) and rat liver cell (BRL-3A) (Sha et al., 2011). Nano-TiO2 showed toxicity in many animal organ types, including lung (Freyre-Fonseca et al., 2011; Liu et al., 2009; Rehn et al., 2003), kidney (Gui et al., 2011), liver (Cui et al., 2011; Palaniappan and Pramod, 2011) and brain (Shin et al., 2010). The toxicity of nano-TiO2 was much attributed to the generation and accumulation of reactive oxygen species (ROS) which resulted in an inflammatory response (Gonzalez et al., 2008; Jaeger et al., 2012; Park et al., 2008). The role of cellular ROS induced by nano-TiO2 had been gained much attention, but the exact mechanism was not fully described. Studies of mitochondrial injury induced by nano-TiO2 which might be related to the mechanism were rarely reported yet. The present study was designed to investigate the effect of nano-TiO2 in A549 cells and rats. Cell ultrastructure, cell viability and intracellular ATP level were detected to access the cytotoxicity of nano-TiO2. Three methods including alveolar epithelial cell ultrastructure, lung tissue pathology and lung tissue MDA level were taken to check the effect of nano-TiO2 in rats. The effects of nano-TiO2 were comprehensively understood by experiments which were carried out both in A549 cells and rats. ATP assay and lipid peroxidation MDA assay helped us to reveal the possible mechanism of the toxicity induced by nano-TiO2. All chemicals and reagents were obtained from Sinopharm Chemical Reagent Beijing Co., Ltd. (Shanghai, China) unless otherwise stated. Modified Eagle Medium (MEM), fetal bovine serum (FBS) were purchased from Thermo Fisher Scientific (Shanghai, China). Nano-TiO2 particles were supplied by Shanghai Institute of Ceramics, Chinese Academy of Sciences (Shanghai, China). The cell counting kit-8, ATP assay kit and lipid peroxidation MDA assay kit were purchased from Beyotime Institute of Biotechnology (Jiangsu, China). Nano-TiO2 particles were sterilized by heated at 120 ◦C for 2h, then suspended in MEM (for A549 cells) or normal saline (for rats) to a stock solution and kept at 4 ◦C until used. Before application, the suspension was ultrasonicated for 30min in a break every 10min for vortex. A stable suspension of nano-TiO2 was obtained in this way and used immediately. Morphology and structure of nano-TiO2 were analyzed by TEM (H-7500, Hitachi, Japan) imaging. A549 cells were obtained from China Center for Type Culture Collection (Beijing, China). Cells were grown in MEM supplemented with 10% (v/v) FBS. Cells were maintained at 37 ◦C in a humidified atmosphere of 5% CO2 and passed at 80% of confluence. Twenty healthy male rats (200 ± 2 g) were purchased from Experimental Animal Center of Chinese Academy of Sciences (Shanghai, China). The rats were kept in a positive pressure air-conditioned unit for animal housing. Distilled water and sterilized food were available. The rats were allowed to acclimate 5 days prior to experimental use. The rats were randomly divided into four groups, including a control group treated with normal saline and three experimental groups treated with 0.1, 1.0 and 10.0mg/mL nano-TiO2 suspension, respectively. Rats were anesthetized with chloral hydrate, and then nano-TiO2 suspension was instilled into the rat's trachea using an indwelling needle. Three days later, all rats were dissected for the following study. Cell viability was detected by cell counting kit-8 according to the manufacturer's instruction. Briefly, A549 cells were seeded in 96-well culture plates with 5 × 103 cells in 100-L MEM per well, after 24h of cell attachment, A549 cells were treated with nano-TiO2 for 4h. Six replicate wells were used for each control and tested concentrations. After incubated for 4h, 10-L 2)-2Htetrazolium, monosodium salt (WST-8) solution was added to each well and the cells further incubated at 37 ◦C for 1h in a 5% CO2 humidified incubator. Absorbance was quantified at 450nm using a microplate reader (Synergy2, Bio-tek, USA). The results of the treated groups were compared to the control group and represented as the percentage of the control value. Intracellular ATP level was a sensitive readout of mitochondrial state and determined using a firefly luciferase-based ATP assay kit. Briefly, cells treated with nano-TiO2 for 4h were lysed and centrifuged at 12,000 × g for 5min in 24-well plates, 100-L supernatant was mixed with 10-L ATP detection working dilution. Luminance was measured by a luminometer (Synergy2, Bio-tek, USA). ATP content was calculated according to a standard curve (Chen et al., 2009). Malondialdehyde (MDA), a product of lipid peroxidation, was analyzed by using an assay kit (Beyotime Institute of Biotechnology). All procedures were completely complied with the manufacturer's protocol. The concentration of MDA could be measured at a wavelength of 532nm by reacting with thiobarbituric acid (TBA) to form a stable chromophoric production. MDA contents were calculated according to a standard curve. Lungs obtained from rats were washed with phosphate buffer solution (PBS) and fixed with 2.5% glutaraldehyde. All fixed lung tissues were routinely processed, embedded in paraffin, sectioned at 3m, and stained with hematoxylin and eosin (H&E) for light microscopic examination. Samples obtained from experiments were fixed with 2.5% glutaraldehyde and post-fixed with 1% OsO4. Then all samples were routinely washed with PBS, dehydrated in a graded series of acetone, and embedded in epoxy resins. Ultrathin sections were cut with an ultramicrotome (UC6, Leica, Germany), deposited on copper grids, stained with uranyl acetate and lead citrate and observed by a transmission electron microscope (H-7650, Hitachi, Japan). All numerical data are presented as mean ± standard deviation (SD) of three separate experiments. Data were analyzed by a one-way ANOVA followed by Dunnett's t-test for comparisons between groups. p < 0.01 was considered significant compared to control. The morphology and size of anatase nano-TiO2 were shown in Fig. 1. There was no surface coating on nano-TiO2. Nano-TiO2 with ultrasonicating (Fig. 1B and C) exhibited better distribution in size than those without ultrasonicating (Fig. 1A). There was no significant difference whether the dissolvent was normal saline (Fig. 1B) or culture media (Fig. 1C). The size of nano-TiO2 particles which were ultrasonicated ranged from 20 to 50nm. A549 cells were treated with four concentrations (50, 100, 200 and 300g/mL) of nano-TiO2 for 4h.Changes of A549 cell ultrastructure were shown in the images of Fig. 2. The A549 cell Fig. 4 – The mitochondria state was represented by intracellular ATP content. The intracellular ATP content decreased gradually after the treatment with four concentrations of nano-TiO2. Data were expressed as the mean ± SD. *p < 0.01 compared with the control. in non-treated control group was normal (Fig. 2A). Clusters of nano-TiO2 enveloped by plasmalemma emerged in treated groups. The higher concentration of nano-TiO2, the more nano-TiO2 appeared in the A549 cell. Mitochondrial cristae were dissolved in a dosage-dependent manner. What's more, mitochondrial cristae which were much closer to nano-TiO2 were dissolved more seriously. The effect of nano-TiO2 on A549 cell survival was assessed by cell counting kit-8 and the result was presented as a percentage of control group viability. A549 cells were treated with four concentrations (50, 100, 200 and 300g/mL) for 4h, and then the viability was checked. A dose-dependent decrease in cell viability was demonstrated by CCK-8 assay as shown in Fig. 3. The viability of A549 cells cultured with 300g/mL nano-TiO2 for 4h was decreased to 68.5%. Intracellular ATP content reflected the state of mitochondria. As shown in Fig. 4, nano-TiO2 led to a concentrationdependent decrease in intracellular ATP content. Cellular ATP levels from 2.72 ± 0.11 to 1.55 ± 0.05mol/mL was detected after A549 cells were treated with four concentrations (50, 100, 200 and 300g/mL) of nano-TiO2 for 4h. The lungs obtained from rats were processed for the histopathological observation (Fig. 5). In control group (Fig. 5 A), the alveolar wall was thin and alveolar structure was normal. After single intratracheal instillation of different concentrations of nano-TiO2, alveolar wall thickened in treated groups. Some alveoli were ruptured and formed big bubbles especially in 10mg/mL nano-TiO2 treated group (Fig. 5D). The amount of red blood cells which appeared in the lung tissue increased with the increasing concentration of nano-TiO2. In 10mg/mL nano-TiO2 treated group (Fig. 5D), nano-TiO2 could be seen obviously in the lung tissue. The ultrastructure of alveolar epithelial cells obtained from rats which treated with single intratracheal instillation of different concentrations of nano-TiO2 was shown in Fig. 6. In control group (Fig. 6A), the cell was normal. In 0.1mg/mL and 1.0mg/mL treated groups (Fig. 6B and C), nano-TiO2 enveloped by plasmalemma and a few lysosomes appeared in the cell. In 10mg/mL treated group (Fig. 6D), lots of nano-TiO2 could be seen in the cell and the amount of lysosomes increased compared to the other treated group. MDA was the representative product of lipid peroxidation and the content could reflect the rate and strength of lipid peroxidation. Rats were treated with three concentrations of nano-TiO2 (0.1, 1.0 and 10mg/mL) with single intratracheal instillation. Nano-TiO2 led to a concentration-dependent increase of intracellular MDA content as shown in Fig. 7. MDA content of 10mg/mL treated group was more than control group by 35%. At present, the biological effects of nanometer materials have been largely reported. The toxicity of nanometer materials was much attributed to the generation and accumulation of reactive oxygen species (ROS). Mitochondria carried out aerobic respiration. Most of oxygen was combined with electrons which came from electronic transmission chain at inner mitochondrial plasmalemma and formed H2O after a series of oxidation–reduction reactions. However, a small amount of oxygen was formed superoxide which was the major source of ROS in cells. The cumulative ROS attacked the mitochondria and restrained the functional activity of mitochondria, as a result, the cell was approaching to apoptosis. A549 cells were exposed to four concentrations (50, 100, 200 and 300g/mL) of nano-TiO2 for 4h. Intracellular nano-TiO2 enveloped by plasmalemma indicated that nano-TiO2 could be swallowed into the A549 cell by endocytosis. Nano-TiO2 showed cytotoxicity according to the cell viability experiment and intracellular ATP level. Nano-TiO2 caused a dose-dependent decrease of intracellular ATP level which must be related to the generation of ROS induced by nano-TiO2. Mitochondria surrounded by a high concentration of ROS were damaged and ATP generation was blocked. The cell was approaching to apoptosis because of lack of energy, and as a result A549 cell viability inevitably dropped. Since nano-TiO2 could be swallowed by A549 cells and showed cytotoxicity, nano-TiO2 should cause some harmful influence on rats. Rats were treated with single intratracheal instillation of nano-TiO2 concentrations. Thick alveolar and big bubbles indicated the lungs injury caused by nano-TiO2. The increasing number of red blood cells indicated the worse inflammation of lungs. The pathology result was in accordance with the ultrastructure of alveolar epithelial cells. The higher concentration of nano-TiO2, the more lysosomes appeared in the cell which was related to the cell apoptosis. The results of lung tissue lipid oxidation accessed by MDA content confirmed that the damage of lung tissue was indeed related with the generation and accumulation of ROS. In conclusion, nano-TiO2 really showed toxicity not only in cells but also in rats. Nano-TiO2 stimulated the generation of ROS which damaged the mitochondria and the generation of ATP in mitochondria was disturbed. The cell was approaching to apoptosis. In macroscopic view, the function of lungs was injured and the inflammation occurred. Based on the results reported in this study, the role of nano-TiO2 was shown in Fig. 8. The authors declare that there are no conflicts of interest. This work was financially supported by National Key Technology R&D Program (31-1122ZCKF) and Quality Public Welfare Industry Research (201210284-2). Chen, K., Zhang, Q.Y., Wang, J., Liu, F.J., Mi, M.T., Xu, H.X., Chen, F., Zeng, K.H., 2009. Taurine protects transformed rat retinal ganglion cells from hypoxia-induced apoptosis by preventing mitochondrial dysfunction. Brain Res. 1279, 131–138. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 4.0, \"mgL\": null, \"viab\": 87.06105}, {\"h\": 4.0, \"mgL\": null, \"viab\": 78.76518}, {\"h\": 4.0, \"mgL\": null, \"viab\": 76.395164}, {\"h\": 4.0, \"mgL\": null, \"viab\": 68.59472}, {\"h\": 4.0, \"mgL\": null, \"viab\": 86.71875}, {\"h\": 4.0, \"mgL\": null, \"viab\": 78.515625}, {\"h\": 4.0, \"mgL\": null, \"viab\": 69.53125}, {\"h\": 4.0, \"mgL\": null, \"viab\": 56.640625}]}", "role": "assistant" } ]	148.json
[ { "content": "Extract sample context and measurement list. <doc> ARTICLE IN PRESS Biomaterials 28 (2007) 2572–2581 www.elsevier.com/locate/biomaterials Thomas R. Pisanic IIa,-, Jennifer D. Blackwella , Veronica I. Shubayevb,d, Rita R. Fin ̃onesc , Sungho Jinc a Department of Bioengineering, University of California, San Diego La Jolla, CA 92093, USA b Department of Anesthesiology, University of California, San Diego School of Medicine La Jolla, CA 92093, USA c Mechanical and Aerospace Engineering Department, University of California, San Diego La Jolla, CA 92093, USA d San Diego Veteran Affairs Medical Healthcare System, La Jolla, CA 92093, USA Received 29 October 2006; accepted 31 January 2007 Available online 11 February 2007 Magnetic nanoparticles (MNPs) have shown great promise for use as tools in a wide variety of biomedical applications, some of which require the delivery of large numbers of MNPs onto or into the cells of interest. Here we develop a quantifiable model cell system and show that intracellular delivery of even moderate levels of iron oxide (Fe2O3) nanoparticles may adversely affect cell function. More specifically, we show that exposure to increasing concentrations of anionic MNPs, from 0.15 to 15 mM of iron, results in a dosedependent diminishing viability and capacity of PC12 cells to extend neurites in response to their putative biological cue, i.e. nerve growth factor. The cytotoxicity results of biomaterials in our model system imply that more study into the acute and long-term effects of cellular Fe2O3 internalization is both warranted and necessary. r 2007 Elsevier Ltd. All rights reserved. Keywords: Nanoparticle; Magnetism; Neural cell; Cytotoxicity; Biocompatibility The delivery of magnetic nanoparticles (MNPs) to or into various cell types has become an area of increasing interest in the biomedical sciences [1,2]. For example, cell labeling with MNPs is now a widely used means of in vitro cell separation, where a high gradient magnetic field is used to isolate cells of interest, tagged with magnetic particles via phenotypic markers, from a heterogeneous population [3,4]. Targeted delivery is used to deliver drugs or genes regio-specifically by attaching them to MNPs and locally concentrating the resulting complexes in vivo to the desired locale [5,6]. Similarly, magnetic hyperthermia, the local concentration of MNPs and subsequent heating via AC magnetic fields, has shown promise as a potentially viable cancer therapy [7,8]. And it has additionally been shown that if cells are labeled with large enough numbers of MNPs, that these cells can be located, tracked and recovered using imaging techniques such as high-resolution magnetic resonance imaging (MRI) [9,10]. While these imaging technologies boast exceeding sensitivities, cells of interest must be labeled with relatively large amounts of magnetic material in order to be readily detected. Due to this constraint, much effort has been dedicated to the development of efficient mechanisms of delivery of MNPs into cells of interest. Many groups have investigated the use of various organic coatings as a means of optimizing the delivery of MNPs to or into cells. Toward these ends, MNPs have been coated with ligands to cell surface receptors [11,12] in order to utilize a receptor-mediated endocytotic pathway, amphiphilic coatings such as polyethylene glycol (PEG) [13,14] or dendrimers [15] to allow their free passage into cells and also with well-known transfection agents such as the HIV-derived TAT protein [9,10]. In several cases it has been shown that even a simple dimercaptosuccinic acid (DMSA) coating can improve uptake efficiency by three orders of magnitude, presumably by engendering the MNPs with an anionic charge, resulting in nonspecific adsorption to the cell surface followed by endocytosis into the cell [16–18]. Indeed these techniques succeed in delivering large numbers of MNPs into the cells (up to 3–4 orders of magnitude over fluid phase endocytosis), but a valid concern arises over the effects that large intracellular concentrations of iron oxide (Fe2O3) might have on normal cell behavior. In addition to magnetic tracking, many biomedical applications of MNPs such as targeted drug delivery and hyperthermia require very large amounts of MNPs to be injected or targeted to the cells or area of interest. While historically Fe2O3 nanoparticles are considered to be well tolerated in vivo [19,20], the effects of moderate to high intracellular concentrations of Fe2O3 nanostructures upon cell function remain to be adequately elucidated. Although a few studies have been performed investigating the acute cytotoxicity of MNPs and their qualitative effects upon cellular morphology [13,20–24], little work has focused on quantifying the effects that Fe2O3 internalization has upon cell behavior and, in particular, the ability of cells to appropriately respond to biological cues. For example, Curtis et al. studied the effects that various nanoparticle surface coatings had on cellular morphology, uptake efficiency, cytotoxicity and, in one case, cell mobility [13,21,22]. Indeed, these papers have shown that nanoparticles and the surface coatings employed can have a dramatic effect upon the relative cell behavior and morphology. More quantitatively, Hussain et al. [25] presented data on the effects that various metal/metal oxide nanomaterials had upon liver cell function in vitro. Here we demonstrate with an established biomedically used Fe2O3 nanoparticle and surface coating preparation, viz. anionic MNPs (AMNPs), that changes in cell behavior and phenotype can be quantified and directly correlated with the level of AMNP exposure. Enhanced endocytosis via [anionic] DMSA coating is a simple, efficient and well-characterized method of intracellular delivery of Fe2O3 nanoparticles. While several studies have clearly shown little to no in vivo toxicity of either of these components individually [19,20,26], only recently have they been combined to deliver large numbers of nanoparticles into cells [16–18,27]. In order to clearly evaluate the effects that Fe2O3 nanoparticle internalization might have upon cell behavior, phenotype and response to biological cues, a readily quantifiable model cell system was chosen. The PC12 pheochromocytoma clonal cell line has served as a paradigm for neurobiological and neurochemical studies [28]. A prominent characteristic of PC12 cells is their readily quantifiable, rapid and reversible response to nerve growth factor (NGF), resulting in the extension of neurite-like processes up to 1 cm in length, comparable to those of sympathetic neurons [29]. Here we examine and quantify the specific effects that DMSA-coated MNPs have upon cultured PC12 cells and evaluate their ability to respond to NGF as compared to control cells. The focus of this paper is thus on the cytotoxicological evaluation of a previously characterized nanostructure intended and accepted for biomedical uses and its effect upon a readily quantifiable model cell system. Maghemite nanoparticles were synthesized as previously described in [30]. Two starting solutions were made by adding 5.406 g of FeCl3 (Sigma-Aldrich, St. Louis, MO) to 20 ml of deionized (DI) water and, separately, adding 1.988 g of FeCl2 (Sigma-Aldrich) to 5 ml of a 2 N solution of HCl (Sigma-Aldrich). These solutions were then added to 100 ml of DI water under vigorous stirring, to which 120 ml of a 2 M ammonia solution was added and remained under vigorous stirring for 5 min, during which, a black precipitate formed, indicating the synthesis of Fe3O4. This solution was centrifuged at 900g for 5 min, the supernatant was removed and the nanoparticle solution was redispersed in DI water. This solution was centrifuged at 900g for 5 min and resuspended in DI water. This washing process was repeated 4 additional times and the centrifugate was finally resuspended in 35 ml of DI water, resulting in an opaque black solution. This solution was heated to 80 1C and the MNPs were oxidized, by bubbling oxygen for 2 h, to Fe2O3, as indicated by a dark brown color. The mass of resulting maghemite was determined through drying and weighing to be 1.68 g of Fe2O3 in 50 ml of DI water. DMSA coating was performed as previously described [31]. Separately, 0.26 g of DMSA (Sigma-Aldrich) was added to 313 ml of deoxygenated DI water (via 1 h of nitrogen bubbling) and 205 ml of deoxygenated DI water was added to the maghemite solution. These solutions were deoxygenated for an additional 2 h at room temperature. The maghemite solution was then vigorously stirred while bubbling nitrogen and the pH of both solutions was adjusted to 3.0 with HNO3 prior to mixing the solutions under vigorous stirring with constant nitrogen bubbling. The reaction was allowed to proceed for 30 min and the resulting mixture was spun down at 800 g for 5 min and resuspended in 200 ml of DI water. Under constant stirring, the pH of this solution was adjusted with NaOH to 9.25–9.5 and maintained for 30 min before lowering the pH to 7.4 with HCl. The resulting solution was centrifuged at 1000g for 10 min to remove any precipitated aggregates and the resulting opaque dark brown supernatant was deoxygenated, sterile filtered and stored under nitrogen at 4 1C. The final stock concentration of Fe2O3 was adjusted with DI water to 150 mM Fe concentration. The rat pheochromocytoma cell line PC12M was derived from the PC12 line by Marc Montminny (Salk Institute, La Jolla, CA) and exhibits increased adherence to polystyrene dishes. The line was grown in highglucose Dulbecco's modified Eagles medium (DMEM) (Gibco, Grand Island, NY) containing 10% fetal bovine serum (FBS) (Invitrogen, Carlsbad, CA), 5% heat-inactivated horse serum (Omega Scientific Inc.), 100 U/ml penicillin and 100 mg/ml streptomycin (Gibco) at 37 1C in 5% CO2 until reaching 70% confluence. PC12 cells were plated from culture at a concentration of approximately 20,000 cells/ml into 6 or 12 well plates, at volumes of 2 and 1 ml, respectively, in high-glucose DMEM (Gibco) containing 10% FBS (Invitrogen), 5% heat-inactivated horse serum (Omega Scientific Inc.), 100 U/ml penicillin and 100 mg/ml streptomycin (Gibco) at 37 1C and allowed to adhere for 2 days. The following day, the cells were washed and the media was replaced with serum and phosphate-free HEPES-buffered DMEM (Invitrogen) in order to prevent particle agglomeration. To this solution a concentrated stock solution of AMNPs ([Fe] 1⁄4 150 mM) was added, yielding final iron concentrations of 15, 1.5 mM and 150 mM, as well as a phosphate-free HEPES-buffered DMEM only control. After overnight incubation, the cells were washed and the modified DMEM was replaced with media containing 1% FBS, 5% heat-inactivated horse serum and 1% penicillin-streptomycin; NGF (Sigma-Aldrich) was added to a working concentration of 100 ng/ml to induce sprouting. The cells were cultured with this media, including changes every other day, for the remainder of each experiment. For AMNP only analyses, the samples were dried onto a carbon TEM grid and were observed using a transmission electron microscope (TEM) (FEI Sphera Tecnai T2 ). For cell samples, cells were plated as described above into 6 well size Petri dishes. Twenty-four hours following exposure to AMNPs, cells were fixed in an aqueous solution of 100 mM sodium cacodylate (Sigma-Aldrich) containing 2% paraformaldehyde (Sigma-Aldrich) and 2% glutaraldehyde (Sigma-Aldrich) at 37 1C for 2 min and then allowed to cool to room temperature for 25 min. The cells were then washed 3 times in ice-cold 100 mM cacodylate buffer and kept on ice. Cells were then post-fixed in ice cold 100 mM sodium cacodylate containing 1% osmium tetroxide (Sigma-Aldrich) for 30 min on ice and then washed 3 times with ice cold DI water followed by a 30 min post fixation in 2% uranyl acetate (Fluka, Seelze, Switzerland) and another three washes with ice cold DI water. The samples were then serially dehydrated in ethanol at room temperature and finally embedded in Durcupan (Fluka), which was sectioned prior to TEM analysis. The samples were then placed onto a carbon TEM grid and were observed using a TEM (JEOL 200CX). Cells were plated as described above in 12 well plates. Prior to, 2, 4 and 6 days following AMNP exposure, the cells were first washed in PBS at 37 1C and left in 1 ml of PBS/well. Calcein AM and ethidium homodimer-1 (ethyl-D) were used from a LIVE/DEAD Viability/Cytotoxicity Kit (Invitrogen) and diluted with PBS to a final concentration of 10 mM each. Each well was aspirated and 20 ml of both stain solutions was added to each well and incubated at 37 1C for 20 min. The samples were analyzed via fluorescent microscopy with appropriate fluorescent filter cubes (FITC cube for calcein, TRITC cube for ethyl-D). A total of at least 300 cells were counted per condition per sample per timepoint, with green fluorescence (emission 515 nm) indicating living cells and red fluorescence (emission 635 nm) indicating dead cells. Cell detachment was quantified by taking media samples from the culture wells 2, 4 and 6 days following AMNP exposure and determining the cell density, and thus total detached cells/well, using a standard hemocytometer (Hausser Scientific, Horsham, PA). The PC12 cells were plated as described above into 6 well plates containing sterile glass coverslips; 2 , 4 and 6 days following AMNP exposure the coverslips from particular wells of each exposure condition were removed and extracted in a solution of 10 mM 2-morpholinoethanesulfonic acid (MES) buffer (Sigma-Aldrich), 138 mM KCl, 3 mM MgCl2, 2 mM EDTA and 320 mM sucrose for 45 s, followed by fixation in the same buffer containing 4% formaldehyde for 15 min. The coverslips were then washed once in Tris buffered saline (TBS) (150 mM NaCl, 20 mM Tris-HCl, pH 7.4) and stored in TBS at 4 1C until staining. At the time of staining, the samples were permeabilized in a solution of TBS containing 0.5% Triton-X 100 (TX) for 10 min and then rinsed 3 times in TBS containing 0.1% TX for 4 min each rinse. The samples were then blocked in a solution of TBS-0.1% TX with 2% BSA (Sigma-Aldrich), and 0.1% sodium azide (Sigma-Aldrich) for 10 min, followed by incubation for 20 min with mouse anti-tubulin (Molecular Probes, Eugene, OR) diluted to 1 mg/ml in the blocking buffer. The samples were then washed four times in TBS-0.1% TX and fluorescein conjugated anti-mouse secondary antibody was then added at 5 mg/ml of blocking buffer and incubated for 20 min. The samples were washed 4 times in TBS-0.1% TX and then incubated with rhodamine-phalloidin diluted in blocking buffer to 1 mg/ml for 20 min and then washed twice in TBS-0.1% TX. Finally the samples were rinsed once in TBS, once in DI water, drained and mounted onto glass slides. The samples were then analyzed by standard fluorescent microscopy (DMIRB microscope; Leica, Wetzlar, Germany) at 100 with fluorescein and rhodamine filter cubes (Leica), for microtubules and actin, respectively. Six days following exposure to AMNPs, the 6 well plate samples were analyzed via phase contrast microscopy at a magnification of 10 - . Micrographs of random areas of the culture plates were taken and subsequently analyzed with ImageJ (NIH, Bethesda, MD) for number of neurites per cell, length per neurite and number of intercellular processes per cell. Depending on the growth characteristics, 30–50 cells were analyzed for each parameter for each experimental condition. Six days following exposure to AMNPs, cells were lysed in a 50 mM Tris-HCl buffer containing 10% glycerol, 0.1% sodium dodecyl sulfate (SDS) (Sigma-Aldrich), 1% TX, 150 mM NaCl, 1.5 mM MgCl2, and protease inhibitors [2 mg/ml aprotinin, 50 mM leupeptin, 1 mM pepstatin, 10 mM phenylmethylsulfonyl fluoride (PMSF) and 5 mM EDTA]. The samples were then reduced in 10% b-mercaptoethanol and centrifuged at 13,000 RPM for 5 min. Thirty mg of protein was added to each lane of 10% SDS-polyacrylamide gel and underwent electrophoresis at 160 V for 90 min. The separated protein was then transferred to a nitrocellulose membrane at 50 V for 1 h in a Tris-glycine transfer buffer (Invitrogen) containing 12 mM Tris base, 95 mM glycine, and 20% methanol. The membranes were then blocked in 5% nonfat dry milk (Bio-Rad, Hercules, CA) for 1 h and incubated overnight at 4 1C with anti-GAP-43 (Sigma-Aldrich) antibody diluted 1:1000 in a 1% BSA solution. Horseradish peroxidase conjugated goat anti-mouse IgG (ECL Kit; Amersham, Arlington Heights, IL) was then applied for 1 h at room temperature. The blots were developed in luminol and exposed to Hyperfilm ECL (Amersham). The blots were stripped and normalized by reprobing with a gel loading (b-actin) control. The molecular weights were compared with prestained low-range standards (Bio-Rad). The final AMNP product was a highly concentrated aqueous ferrofluid, stable over a wide range of pH and salt concentrations. Fig. 1A shows a TEM image of the dried final ferrofluid with average nanoparticle diameters between 5 and 12 nm. The observation of apparent aggregation in Fig. 1A is an artifact of the drying process for standard TEM sample preparation, as the AMNP fluid itself is highly stable against aggregation in water and many other aqueous-based buffer solutions. As previously stated, AMNPs have been shown to be readily taken up by a variety of cells via an endocytotic mechanism. The internalization of AMNPs by PC12 cells was verified using TEM. Fig. 1B shows large numbers of AMNPs free within the cytoplasm and contained inside numerous endosomes, accumulated in the perinuclear region within the cells. It was additionally noted that the intracellular levels of AMNPs were qualitatively proportionate to the added concentrations of the particles (data not shown). In the initial assessment into the effects of AMNP internalization, standard cytotoxicological evaluations of acute toxicity to the cells were performed using a fluorescent live/dead stain. The stain consists of the dyes calcein AM and ethylidium homodimer-1 (EthD-1), which stain for live and dead cells, respectively. Live cells are stained by membrane-permeant non-fluorescent calcein AM which is converted by intracellular esterases of live cells to the brightly fluorescent calcein, yielding a uniform green fluorescence inside the cells. EthD-1 is non-permeant and is extruded by the intact membrane of live cells; in compromised membranes of dead cells, however, EthD-1 directly binds to nucleic acids and undergoes a 40-fold increase in red fluorescence. Fluorescent images of cells stained with the live (green) and dead (red) stain 6 days after AMNP exposure are shown in Figs. 2A (15 mM iron concentration) and 2B (Control cells). Assessment of cell viability (n 1⁄4 300 per timepoint) through a series of experiments demonstrates statistically significant reductions in PC12 cell viability after exposure to the particles (Fig. 2C). Almost all observed cell death occurs within the first 48 h following incubation with the particles and the total observed cell death increases as a function of AMNP exposure concentration. In addition to cell death, significant cell detachment was also hemocytometrically quantified from the media and observed to correlate to AMNP exposure. A plot of total cell detachment per well as a function of time post exposure to various AMNP concentrations is shown in Fig. 2D. While previous reports have also shown qualitative disruption of the cytoskeleton due to exposure to high concentrations of MNPs [13,21,22,24], this study investigates methods of quantifying phenotypic changes in cells as a function of exposure to increasing nanoparticle concentration. Normally, PC12 cells are characterized by their rapid and reversible response to NGF, resulting in the extension of neurite-like processes up to 1 cm in length, comparable to those of sympathetic neurons [29]. This profound and easily observable response makes PC12 cells an ideal model system for the study of neural stem cell development and differentiation, neurite regulation and outgrowth, and as a general model of cell response to biochemical cues. These changes can be readily quantified and, based on previous literature [32], the following parameters were used to compare the phenotypic response of cells exposed to AMNPs: frequency of neurite lengths, the number of neurites or ''sprouts'' extending from the soma, the number of branches per neurite, the number of intercellular processes and quantification of NGF-induced expression of specific proteins. The most obvious effect that exposure to the AMNPs has upon the cells is their ability to generate mature neurites. The average number of neurites that sprouted from each [remaining] live cell (dead cells were not included) was measured at each of the three concentrations and compared with media only control (Fig. 4). Fig. 4A shows a typical NGF-stimulated PC12 cell in culture with arrows showing the location of its neurites (N) and a line delineating the length of an extended neurite. The data from these evaluations, plotted in Fig. 4B, shows the effect of AMNPs on the generation of neurites. Of the live cells evaluated, those exposed to 0.15, 1.5, and 15 mM AMNP iron concentrations on average produced 2.67, 1.9, and 0.97 neurites per cell (n 1⁄4 30 cells/condition), respectively, as compared with 2.79 in the control cells. The lengths of the neurites which did extend were further evaluated, as shown in Fig. 4A. Of those cells that did sprout neurites (non-sprouting cells were not included), not only were there fewer neurites per cell, but also the lengths of the neurites that did extend were dramatically affected by exposure to AMNPs. In Fig. 4C, an inverse correlation between the level of AMNP exposure and the ability of the cells to respond normally to NGF and extend neurites into the periphery is seen. The histogram shows the frequency of neurite lengths of cells exposed to AMNPs at the tested concentrations. Notice the increasing inability of PC12 cells exposed to higher AMNP concentrations (X1.5 mM iron) to develop mature extended neurites, once again clearly indicating a negative relation between AMNP exposure and the production of neurites. Similar to most neuronal type cells, the neurites of PC12 cells typically branch out in order to establish intercellular contacts. Such branching is vital to the formation of neuromuscular connections, synaptogenesis, neurite extension, regeneration and signal transduction associated with these processes. As such, formation of intercellular contacts is considered a metric of PC12 cell and neuronal morphology. Additionally, it has been readily established that disruption of these abilities results in diminished in vivo function and utility of neuronal type cells. Typical intercellular contacts (IC) for control cells (0 mM Fe) are shown in Fig. 5A. Fig. 5B shows that nanoparticle exposure reduces the ability of neurites to develop intercellular contacts and thus physically interact with one another. When plated at a standard density, the cells exposed to AMNPs fail to develop intercellular contacts at the same frequency as control cells. For each condition, the contacts (as indicated in Fig. 5A) of over 250 cells were tallied and divided by the total number of cells. Even exposure to the most moderate of AMNP concentrations results in an over 30% reduction in intercellular contacts per cell (1.01 contacts/cell for control compared with 0.67 contacts/cell in 0.15 mM treated cells), likewise, higher concentrations further diminish intercellular communication (1.5 mM:0.31 contacts/cell; 15 mM:0.09 contacts/cell). Lastly considered were the effects of AMNP exposure on the expression of protein related to axonal sprouting and neuronal function in order to determine whether the effects of exposure were strictly morphological or phenotypic as well. Growth-associated protein-43 (GAP-43) is a neuronal protein associated with axonal growth, neuronal plasticity Consistent with earlier literature [26], it was independently verified that DMSA concentrations (7.5, 75 and 750 mM) that are equivalent to or higher than the net concentrations of DMSA in experiments have little to no measurable effect on the parameters tested in these studies. While this certainly does not exclude the possibility that DMSA might have an effect upon these parameters when carried intracellularly into the cell, it does confirm that the effects seen here are not the result of DMSA exposure alone. This study indicates that even temporary exposure to Fe2O3 AMNPs results in a dose-dependent reduced ability of PC12 cells to appropriately respond to nerve growth factor. PC12 cells exposed to AMNPs show reduced viabilities, increased cytoskeletal disruption, and a diminished ability to form mature neurites in response to NGF exposure as compared to control cells. This may have significant implications for in vivo and phenotypic dependent in vitro uses of AMNPs and Fe2O3 MNPs in general. Overall, it was shown that the exposure of cells to even moderate concentrations of MNPs can adversely affect cell function, phenotype and viability. These findings also indicate and confirm previous reports that the presence of intracellular Fe2O3 nanoparticle constructs can result in significant changes in cell behavior and viability [21–23,34]. In light of these results and taking into account the results of other investigators, it seems that nanoparticle surface coatings, while perhaps innocuous themselves, can drastically affect the behavior of cells exposed to nanostructures coated with these agents. While the standard concentration of iron Fe3+ ions where cytotoxic effects normally begin to appear is roughly 4 mM [35], only the highest (15 mM) of the tested MNP concentrations exceeded this value, yet significant negative effects were seen at concentrations over 20-fold lower than the toxic level. It seems that there are three plausible explanations for this observation: first, that coordination of the coating agent with the nanostructure facilitates entry into or interaction with cells of both the nanostructure and surface chemicals, thereby magnifying any interactions (positive or negative) with cellular components; a second, alternative explanation is the variance in the effectiveness of the coatings to shield the nanostructures from [potentially adverse] interactions with cellular components; thirdly, there may also be a combination of these effects. The underlying causes of certain biological effects and interactions may be easier to diagnose than others, such as cytotoxic effects from known toxic nanostructures like cadmium selenide (CdSe)-based quantum dots, where toxicity differences between various coatings can almost certainly be explained through the second posited explanation [36]. In the present case, the observed cytotoxic effects are more difficult to diagnose, but are possibly due to free radical generation through Fenton and/or Haber-Weiss reactions whose effects only become noticeable at higher intracellular concentrations of iron [37]. Another possible explanation for the significant cytotoxicological, morphological and phenotypic effects of AMNP internalization is the apparent migration of the AMNP containing endosomes to the proximal perinuclear region of the PC12 cells, as shown in the TEM image in Fig. 1B, which may drastically impede transcriptional regulation and protein synthesis, such as GAP-43, resulting in loss of cell phenotype and possibly cell death. Yet the mere presence of intracellular Fe2O3 nanoparticles, even at the lowest It should be noted that in the case of ferrofluid cytotoxicological studies, the implementation of a bare MNP control, while desirable, would raise more questions than it would answer. Firstly, an MNP coating is required for colloidal MNP stability in aqueous biological solutions making it unsuitable as an accurate comparison. Bare MNPs are highly unstable in saline solutions; form optically visible aggregates, and produce an interaction of such disparate physical characteristics so as to be irrelevant as a proper control against a stably dispersed ferrofluid. Furthermore, the effect of oxide nanoparticle solubility has recently been shown to directly affect the cytotoxicity of the material [35]. And lastly, the implementation of a different stabilizing coating would beg the question of which coating to utilize and what effect the other coating itself would have upon the cells. In summary, this work has shown that exposure to increasing concentrations of anionic magnetic nanoparticles results in a dose-dependent diminishing ability of PC12 cells to differentiate in response to nerve growth factor. The results of our model system may act as a caveat for the use of AMNPs in neuronal experiments and in biomedicine in general. These results further imply that more study into the effects of cellular iron oxide internalization is both warranted and necessary and act to further bolster recent calls for increased attention and interest into the toxicity of nanomaterials [38,39]. Failure to fully and properly evaluate nanostructures on an individual case-by-case basis may lead to lack of parameter control in in vitro experiments, as well as incorrect assumptions concerning their biocompatibility and biosafety of their in vivo use. We are sincerely grateful to Dr. Eduardo Macagno for his insightful conversations and general advice. We would also like to thank Chiara Daraio and Mariana Loya for all of their help and expertise in the TEM work. We additionally acknowledge the assistance of Norm Olson for TEM analysis using the UCSD Cryo-Electron Microscopy Facility supported by NIH Grants 1S10RR20016 and GM033050 to Dr. Timothy S. Baker and a gift from the Agouron Institute to UCSD. This work was supported by The University of California, San Diego and The American Society Engineering Education (ASEE), National Defense Science and Engineering Graduate (NDSEG) Fellowship. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 48.0, \"mgL\": null, \"viab\": 69.0}, {\"h\": 48.0, \"mgL\": null, \"viab\": 95.45}, {\"h\": 48.0, \"mgL\": null, \"viab\": 90.26}, {\"h\": 96.0, \"mgL\": null, \"viab\": 74.53416149}, {\"h\": 96.0, \"mgL\": null, \"viab\": 73.37662338}, {\"h\": 96.0, \"mgL\": null, \"viab\": 70.77922078}, {\"h\": 144.0, \"mgL\": null, \"viab\": 18.63354037}, {\"h\": 144.0, \"mgL\": null, \"viab\": 4.545454545}, {\"h\": 144.0, \"mgL\": null, \"viab\": 11.68831169}]}", "role": "assistant" } ]	145.json
[ { "content": "Extract sample context and measurement list. <doc> Ajay Kumar Gupta* and Stephen Wells Abstract—Superparamagnetic iron oxide nanoparticles have been used for many years as magnetic resonance imaging (MRI) contrast agents or in drug delivery applications. In this study, a novel approach to prepare magnetic polymeric nanoparticles with magnetic core and polymeric shell using inverse microemulsion polymerization process is reported. Poly(ethyleneglycol) (PEG)-modified superparamagnetic iron oxide nanoparticles with specific shape and size have been prepared inside the aqueous cores of AOT/n-Hexane reverse micelles and characterized by various physicochemical means such as transmission electron microscopy (TEM), infrared spectroscopy, atomic force microscopy (AFM), vibrating sample magnetometry (VSM), and ultraviolet/visible spectroscopy. The inverse microemulsion polymerization of a polymerizable derivative of PEG and a cross-linking agent resulted in a stable hydrophilic polymeric shell of the nanoparticles. The results taken together from TEM and AFM studies showed that the particles are spherical in shape with core–shell structure. The average size of the PEG-modified nanoparticles was found to be around 40–50 nm with narrow size distribution. The magnetic measurement studies revealed the superparamagnetic behavior of the nanoparticles with saturation magnetization values between 45–50 electromagnetic units per gram. The cytotoxicity profile of the nanoparticles on human dermal fibroblasts as measured by standard 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay showed that the particles are nontoxic and may be useful for various in vivo and in vitro biomedical applications. Index Terms—Biological cells, drug delivery systems, magnetic materials, nanotechnology. SUPERPARAMAGNETIC iron oxide nanoparticles with tailored surface chemistry have been widely used experimentally for numerous in vivo applications such as magnetic resonance imaging (MRI) contrast enhancement, tissue repair, immunoassay, detoxification of biological fluids, hyperthermia, drug delivery, cell separation, etc. [1]–[4]. All these biomedical and bioengineering applications require that these nanoparticles have high magnetization values and size smaller than 100 nm with overall narrow particle size distribution, so that the particles have uniform physical and chemical properties. In addition, these applications need special surface coating of S. Wells is with Liquid Research Limited, Gwynedd LL5 7UP, U.K. Digital Object Identifier the magnetic particles, which has to be not only nontoxic and biocompatible but also allow a targetable delivery with particle localization in a specific area [5]. Because of large surface area to volume ratio, the magnetic nanoparticles tend to agglomerate and adsorb plasma proteins. The body's reticuloendothelial system (RES), mainly the kupffer cells in the liver, usually take up these nanoparticles due to the hydrophobic surface. Surface coverage by amphiphilic polymeric surfactants such as poloxamers, poloxamines and poly(ethylene glycol) (PEG) derivatives over the nanoparticles significantly increases the blood circulation time by minimizing or eliminating the protein adsorption to the nanoparticles. PEG is widely used as a coating material for nanoparticles due to the following properties: 1) easy excretion through the kidney; 2) low interfacial free energy water; 3) excluded volume effect; 4) nonimmunogenic properties; and 5) nonantigenic properties. In addition, it has been demonstrated that PEG-modified nanoparticles can interact with cell membranes resulting in enhanced cellular response [6]. For biomedical applications, the size, charge, and surface chemistry of the magnetic particles is particularly important and strongly affects both the blood circulation time as well as bioavailability of the particles within the body [7]. In addition, magnetic properties and internalization of particles depend strongly on the size of the magnetic particles [8]. For example, following systemic administration, larger particles with diameters greater than 200 nm are usually sequestered by the spleen as a result of mechanical filtration and are eventually removed by the cells of the phagocyte system, resulting in decreased blood circulation times. On the other hand, smaller particles with diameters of less than 10 nm are rapidly removed through extravasation and renal clearance. Particles ranging from circa 10 to 100 nm are optimal for subcutaneous injection and demonstrate the most prolonged blood circulation times. The particles in this size range are small enough both to evade RES of the body as well as penetrate the very small capillaries within the body tissues and, therefore, may offer the most effective distribution in certain tissues [9]. An advance in the use of magnetic particles for biomedical applications depends on the new synthetic methods with better control of the size distribution and of particle surface characteristics. The most common method for synthesis of magnetite particles is by coprecipitation from a solution of Fe(II) and Fe(III) salts in an appropriate ratio using alkali metal hydroxides. Smaller and more uniform particles can be synthesized by using the microemulsion approach with a good control over iron oxide amount and magnetic properties [10]. Synthesis of hydrophilic magnetic polymeric nanoparticles with magnetite core and polymeric shell is possible using an inverse microemulsion polymerization process [11]. The strategy of utilizing inverse microemulsion approach to modulating the surface of magnetic nanoparticles with PEG is based on the following prior observations: Superparamagnetic iron oxide nanoparticles of narrow size range can be easily produced and coated with various polymers, providing convenient, readily targetable MRI agents. The object of this study is to present a novel approach to prepare more uniform magnetic polymeric nanoparticles with magnetite core and polymeric shell inside the aqueous cores of reverse micelles. The magnetic polymeric particles prepared have been characterized by various physicochemical methods and the cytotoxicity of the surface modified nanoparticles on human dermal fibroblasts in vitro has been assessed, as compared to those underivatized particles using standard 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. Magnetic polymeric nanoparticles prepared using this method were found to be nontoxic and, thus, may serve as an improved way of drug delivery and targeting. 1) Synthesis of Magnetic Nanoparticles in Reverse Micelles: In order to achieve a narrow particle size distribution, magnetite nanoparticles were prepared by using an inverse microemulsion approach. Highly monodispersed iron oxide nanoparticles were synthesized by using the aqueous core of aerosol-OT (AOT)/n-Hexane reverse micelles (without microemulsions) in N atmosphere. The reverse micelles have an aqueous inner core, which can dissolve hydrophilic compounds, salts, etc. A deoxygenated aqueous solution of the ferric and ferrous salts (molar ratio 2:1, 1 M) was dissolved in the aqueous core of the reverse micelles formed by 0.05 M AOT in n-hexane. Chemical precipitation was achieved by using a 1 M deoxygenated solution of sodium hydroxide. The reaction was carried out in nitrogen atmosphere at low temperature (4 C–6 C) with vigorous stirring. The hexane was evaporated and the particles were recovered by precipitation in an excess of an acetone-methanol mixture (9 : 1 ratio), followed by dialysis using 12-kD cutoff dialysis membrane against double-distilled water to remove unreacted iron salts. The surface of the particles was neutralized with 0.01 M HCl, and the particles were dried in a vacuum oven at 70 C–80 C. Particles were then coated with PEG to form the stable dispersion of the magnetite nanoparticles. 2) Surface Modification of Magnetic Nanoparticles With PEG: PEG is hydrophilic and is widely used in biological research, as it protects surfaces from interacting with cells or proteins. Thus, coated particles may result in increased blood circulation time. Ten-milligram magnetite particles were dispersed in 1.0 ml of deoxygenated water by sonication for 30 min. The aqueous dispersion of magnetic nanoparticles was dissolved in the aqueous cores of reverse micelles together with polymerizable derivative of PEG as a monomer (i.e., MA-PEG) MBA as cross-linking agent under nitrogen gas. Additional amounts of water may be added in reverse micellar solution in order to get the host micellar droplet of desired size. In a typical experimental protocol, to a 50-ml 0.05-M AOT solution in hexane, 500 l of magnetite solution (10 mg/ml), 100 l of MA-PEG (5 mg/ml), and 10 l of MBA (0.5 mg/ml) were dissolved. The solution was stable and brownish transparent at this stage. The nitrogen gas was bubbled through this solution to remove the dissolved oxygen. After 30 min, 20 l of 2% ammonium persulphate as an initiator was added. The polymerization of monomers was carried out by free radical polymerization mechanism at 37 C for 8 h. After polymerization, the particles were purified from unreacted monomers and other toxic reactants by dialysis against distilled water and dried in an oven before characterization. 1) Fourier Transformed Infrared (FTIR) Spectral Studies: The FTIR spectrum was recorded in the transmission mode on a Nicolet Impact 410 spectrometer. The dried samples of magnetite or PEG-modified magnetic particles were grounded with KBr and the mixture was compressed into a pellet. The spectrum was taken from 4000 to 400 cm . 2) Transmission Electron Microscopy (TEM) Studies: The average particle size, size distribution, and morphology were examined using a Zeiss 902 TEM at a voltage of 80 kV. The aqueous dispersion of the particles was drop-cast onto a carboncoated copper grid, and the grid was air dried at room temperature before loading into the microscope. 3) Atomic Force Microscopy (AFM) Studies: The aqueous dispersion of the nanoparticles was put on a glass coverslip and the coverslip was air dried at room temperature. Once dry, the samples were analyzed using the Nanoscope III scanning probe microscope (Digital Instruments, Santa Barbara, CA). 4) Vibrating Sample Magnetometer (VSM) Analysis: Magnetic properties and magnetic particle size measurements have been done using a VSM (Liquid Research Ltd., Gwynedd, U.K.) on liquid samples. 5) Total Iron Determination: In order to determine the total concentration of iron present in the nanoparticles, the nanoparticles were completely dissolved in 30% v/v HCl for 2 h at elevated temperatures (50 C–60 C). The iron concentration was determined by spectrophotometric measurements at 340 nm using a Shimadzu UV-160 A UV-visible recording spectrophotometer. Infinity telomerase-immortalized primary human fibroblasts (hTERT-BJ1, Clontech Laboratories, Inc., Hampshire, U.K.) were seeded onto 13-mm glass coverslips in a 24-well plate at a density of 1 10 cells per well in 1 ml of complete medium for 24 h, after which the growth medium was removed and replaced with the medium containing nanoparticles. For control experiments, medium with no particles was used. The medium used was 71% Dulbecco's modified Eagle's medium (DMEM) (Sigma, Dorset, U.K.), 17.5% Medium 199 (Sigma, U.K.), 9% fetal calf serum (FCS) (Life Technologies, Paisley, U.K.), 1.6% 200 mM L-glutamine (Life Technologies, Paisley, U.K.), and 0.9% 100 mM sodium pyruvate (Life Technologies, Paisley, U.K.). The cells were incubated at 37 C in a 5% CO atmosphere. The fibroblast cells were seeded onto 13-mm glass coverslips at 10 000 cells/ml in a 24-well tissue culture plate. After the cells were attached to the coverslips, cell medium was exchanged with the fresh medium containing nanoparticles, and cells were cultured at 37 C. After 24 h, medium was removed and the cells were washed with phosphate-buffered saline (PBS) followed by viability staining using calcein AM (2 M, Molecular Probes, Leiden, The Netherlands) and ethidium homodimer (4 M, Molecular Probes, Leiden, The Netherlands) for 1 h at room temperature. All samples were viewed on a fluorescence microscope. The MTT assay is a simple nonradioactive colorimetric assay to measure cell cytotoxicity, proliferation, or viability. MTT is a yellow, water-soluble tetrazolium salt. Metabolically active cells are able to convert this dye into a water-insoluble dark blue formazan by reductive cleavage of the tetrazolium ring [13]. Formazan crystals, then, can be dissolved in an organic solvent such as dimethylsulphoxide (DMSO) and quantified by measuring the absorbance of the solution at 550 nm, and the resultant value is related to the number of living cells. To determine cell cytotoxicity/viability, the cells were plated at a density of 1 10 cells/well in a 96-well plate at 37 C in 5% CO atmosphere. After 24 h of culture, the medium in the wells was replaced with the fresh medium containing nanoparticles in varying concentrations. After 24 h, 20 l of MTT dye solution (5 mg/ml in phosphate buffer pH 7.4, MTT Sigma, Dorset, U.K.) was added to each well. After 4 h of incubation at 37 C and 5% for exponentially growing cells and 15 min for steady-state confluent cells, the medium was removed and formazan crystals were solubilized with 200 l of DMSO and the solution was vigorously mixed to dissolve the reacted dye. The absorbance of each well was read on a microplate reader (Dynatech MR7000 instruments) at 550 nm. The spectrophotometer was calibrated to zero absorbance, using culture medium without cells. The relative cell viability (%) related to control wells containing cell culture medium without nanoparticles was calculated by test control . The statistical analysis of experimental data utilized the student's t-test and the results were presented as mean standard deviations. Statistical significance was accepted at a level of 0.05. The surfactant (for example, AOT) when dissolved in nonpolar solvents like hexane forms reverse micelles where hydrophobic tails of surfactants are assembled toward the bulk nonpolar solvent and the hydrophilic head is directed inside enclosing a aqueous core [14]. The magnetic nanoparticles were synthesized by coprecipitation of ferrous and ferric salts solution by concentrated sodium hydroxide solution inside the aqueous cores of reverse micellar droplets. Smaller and more uniform particles were prepared by precipitation of magnetite at low temperature in the presence of nitrogen gas. The synthesis of magnetic nanoparticles in oxygen-free environment not only protects the oxidation of iron oxide particles but also reduces the size of the particles as compared with methods without removing oxygen [15]. The size of the inner aqueous core of reverse micelles is in nanometer range, so the magnetic nanoparticles prepared inside these nanoreactors were found to be very small in size (less than 15 nm) with narrow size distribution. The advantage of utilizing this type of microemulsion system for nanoparticle formation is that the size of nanoparticles can be controlled by modulating the size of aqueous micellar core [12]. Magnetic nanoparticles have a large ratio of surface area to volume and, therefore, tend to agglomerate in order to reduce their surface energy by strong magnetic dipole–dipole attractions between particles. The colloidal suspension of magnetite particles, however, can be stabilized by coating the particle surfaces with high molecular weight polymers such as PEG, polyvinylalcohol (PVA), dextran, etc. Such coatings have been postulated as necessary for effective stabilization for ferrofluids [16]. The aqueous dispersion of magnetic nanoparticles along with the aqueous solutions of MA-PEG and cross-linking agent were dissolved in the aqueous core of the reverse micelles, and since these cores are hydrophilic, the cross linking of MA-PEG with MBA and their subsequent absorption on the nanoparticles' surface takes place inside these droplets, giving a uniform and highly stable polymeric coating. The magnetic polymeric particles synthesized in reverse micelles under controlled conditions of temperature and oxygen have shown fairly narrow size distribution. The colloidal solution of magnetic particles coated with PEG showed very high stability at neutral pH and no sedimentation was observed even after two months of storage at room temperature, whereas uncoated magnetic particles did not form a stable colloidal suspension and sedimented within a week. (a) (b) TEM and AFM studies: TEM picture of the magnetic particles was taken to determine the shape, size, and uniformity of the particles. TEM picture of the magnetic nanoparticles synthesized in inner aqueous core of the water-in-oil microemulsions shows that these particles have a very small size of around 10- to 15-nm diameter with narrow size distribution [Fig. 3(a)]. The size of the particles after coating was about 40- to 50-nm diameter, as was determined by TEM studies [Fig. 3(b)]. AFM was performed to study the shape, size, and surface appearance of the nanoparticles. A drop of diluted aqueous solution was placed on a glass coverslip and dried out at room temperature for 24 h. Fig. 4 represents the AFM image of PEG-modified nanoparticles after water evaporation showing the core shell structure and size homogeneity of the nanoparticles. The average particle diameter of the particles was found about 50 nm, which was in close agreement with the size obtained by TEM studies. Determination of total iron content in magnetic nanoparticles requires total dissolution of the oxides and is achieved by Fig. 5 shows the relative magnetization curve as a function of magnetic field for the uncoated particles. From the figure, no hysteresis curve was observed, which indicates the characteristic superparamagnetic behavior of the particles. The saturation magnetization value of the magnetite nanoparticles was found between 45 and 50 electromagnetic units per gram (emu/g). In addition, the magnetization decreases from the plateau value and reaches zero when the magnetic field is removed. The behavior shows that the iron oxide particles correspond to the single-crystal domain exhibiting only one orientation of the magnetic moment and are magnetite in structure. The size distribution was calculated using the equation based on a log-normal function [17]. The average particle size was found to be around 12.92 0.48 nm, which is in close agreement with the size obtained from TEM measurements. It was found from VSM studies that the magnetic particles shows no diamagnetic contribution and are small enough to exhibit superparamagnetic behavior; thus, they are of particular interest for drug targeting systems, as they do not retain any magnetism after removal of a magnetic field. The cell viability staining using calcein AM/ethidium homodimer showed that the cells exposed to PEG-coated nanoparticles were more than 99% viable. Live–dead cell viability assay is a two-color fluorescence assay that is based on the simultaneous determination of the numbers of live and dead cells. Live cells have intracellular esterases that converts nonfluorescent, cell-permeable calcein acetoxymethyl (calcein AM) to the intensely fluorescent green calcein which is retained within the cells. On the other hand, ethidium homodimer enters the damaged membranes of dead cells and is fluorescent red when bound to nucleic acids. The MTT assay for cell viability evaluation has been described as a suitable method for detection of biomaterial toxicity [13]. The MTT assay relies on the mitochondrial activity of fibroblasts and represents a parameter for their metabolic activity. The proliferation/viability of fibroblasts was measured by MTT assay after culturing for 24 h and showed that cell proliferation was more favorable in case of PEG-coated particles than with uncoated ones. All nanoparticles affected the metabolic activity in concentration dependent manner when they were added in the concentration range of 0–1000 g/ml to the cells. Cytotoxicity of the nanoparticles increased in relation to increasing concentration, as shown in Fig. 6. PEG-coated nanoparticles revealed no cytotoxic effects to cells and they remained more than 100% viable relative to control at concentration as high as 1 mg/ml. These samples increased the viability of the cells to about 10%–40% depending on the nanoparticle concentration in the medium. The increased cell viability can be explained by nutrient effect [18]. Incubation with uncoated plain magnetic particles, however, the fibroblasts showed significant loss in viability of about 25%–50% observed at concentrations of 250 g/ml. Below this concentration, cellular metabolic activity did not change much in comparison with control cells. Nanoparticle–cell interaction depends on the surface aspects of materials, which may be described according to their chemistry, hydrophilic/hydrophobic characteristics, or surface energy. These surface characteristics determine how the nanoparticles will adsorb to the cell surface and more particularly determine the cell behavior on contact. It was observed from cell viability studies that the plain uncoated nanoparticles reduced cell viability significantly as compared to the cells that were not exposed to the nanoparticles. One possible explanation for this large decrease in cell viability may be that these nanoparticles are taken up by the cells as a result of endocytosis or are promoting apoptosis (programmed cell death) due to weak cell adhesive interactions with the nanoparticles. The low toxicity of nanoparticles coated with PEG may be attributed to the fact that PEG is hydrophilic, and it protects surfaces from interacting with cells or proteins. It has been demonstrated that PEG-modified nanoparticles can interact with cell membranes, resulting in enhanced cellular response, as these coatings on the nanoparticles are biocompatible, nonimmunogenic, and nonantigenic [19]. PEGs at present are the most important material for tissue engineering and other biomedical applications including RES avoidance and blood residence prolongation of nanoparticles. PEG consists of hydrophobic parts and hydrophilic tails. The strong anchoring of the PEG molecules on the surface of a particulate carrier results in the steric stabilization of the particles. The steric stabilization improves the particle stability in the biological milieu against interactions with macromolecules (e.g., opsonins) and cells, thus imparting prolonged circulation in blood and reduced RES uptake to particulate carrier system [20]. Biodistribution studies performed in mice have shown that the blood circulation time increases as the molecular weight of the PEG increases (from 1900 up to 20 000 Da) [21]. A wide variety of superparamagnetic iron oxide nanoparticles have been prepared to date and tested for many preclinical applications and in clinical trials [22]–[25]. Most iron oxides have a relatively short blood half-life and their primary application is for imaging of the liver, spleen, and gastrointestinal tract. Surface-modified iron oxide nanoparticles having long blood circulation times, however, may prove very useful for imaging of the vascular compartment (magnetic resonance angiography), imaging of lymph nodes, perfusion imaging, receptor imaging, and target-specific imaging. In this paper, PEG-modified superparamagnetic iron oxide nanoparticles about 50 nm in size with a narrow size distribution have been prepared using an inverse microemulsion polymerization method and characterized in vitro by various physicochemical means. The magnetic polymeric nanoparticles have a core–shell structure with magnetic core and polymeric shell. The colloidal solution of nanoparticles shows high stability. These nanoparticles exhibit superparamagentic behavior and are nontoxic and, hence, may be useful for relevant drug delivery and other biomedical applications. The authors would like to thank Prof. A. S. G. Curtis, University of Glasgow, Glasgow, U.K., for encouraging them to work in his laboratory. Ajay Kumar Gupta received the M.Sc. degree in chemistry and the Ph.D. degree from the University of Delhi, Delhi, India, in 1997 and 2001, respectively. He is currently with the Centre for Cell Engineering, University of Glasgow, U.K., working on the synthesis of superparamagnetic nanoparticles of a specific shape and narrow size distribution with tailored surface chemistry, which may be useful for various biomedical applications. He has worked on drug delivery systems using nanotechnology, and successfully devised a nanoparticulate carrier system using polymeric micelles for ophthalmic delivery of hydrophobic drugs. He has U.S. and European patents and has filed Indian patents for these formulations. The technical know-how has been transferred to a pharmaceutical company, Panacea Biotech, New Delhi, India, for further commercialization of the formulation. Stephen Wells, photograph and biography not available at the time of publication. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 97.3923}, {\"h\": 25.0, \"mgL\": null, \"viab\": 84.941284}, {\"h\": 26.0, \"mgL\": null, \"viab\": 72.867615}, {\"h\": 27.0, \"mgL\": null, \"viab\": 67.59135}, {\"h\": 28.0, \"mgL\": null, \"viab\": 49.814953}]}", "role": "assistant" } ]	144.json
[ { "content": "Extract sample context and measurement list. <doc> Kivilcim Buyukhatipoglu, M.S.,1 Robert Chang, M.S.,1 Wei Sun, Ph.D.,1 and Alisa Morss Clyne, Ph.D.1,2 Tissue engineering may require precise patterning and monitoring of cells and bioactive factors within the scaffold. We investigated a new hybrid nanobioprinting technique that facilitates manipulation and tracking of cells and bioactive factors within a three-dimensional tissue construct. This technique combines the initial patterning capabilities of syringe-based cell deposition with the active patterning capabilities of superparamagnetic nanoparticles. Superparamagnetic iron oxide nanoparticles, either in the alginate biopolymer or loaded inside endothelial cells, were bioprinted using a solid freeform fabrication direct cell writing system. Bioprinting did not impact cell viability when nanoparticles were in the alginate. However, both control and printed samples with 0.1 or 1.0 mg=mL nanoparticles in the alginate showed a 16% or 35% viability loss at 36 h after printing, respectively. Nanoparticle loading in cells decreased cell viability to 11% and bioprinting decreased viability to an additional 29% at 36 h. No changes were observed in any samples after 36 h, suggesting that cell viability stabilized following the initial nanoparticle toxicity effect. Nanoparticles in the alginate and those loaded in cells were moved using an external magnet, depending on biopolymer viscosity, and imaged by microcomputed tomography. The hybrid nanobioprinting method can noninvasively manipulate and track bioactive factors and cells within tissue engineering structures. Tissue engineering is an interdisciplinary field that uses engineering and life science principles to advance our knowledge of tissue growth, which is then applied toward the development of biological tissue substitutes to restore organ function.1 Tissue engineering may require precise patterning of cells and bioactive components to recreate the complex, three-dimensional (3D) architecture of native tissue. These cells and bioactive factors may then need to be repositioned during tissue growth in vitro or after implantation in vivo to achieve the desired tissue properties. Certain biological components that are valuable in tissue development, such as stem cells, growth factors, or nanoparticles themselves, may induce unwanted effects when the tissue-engineered construct is implanted. It may be beneficial to remove these cells and bioactive components entirely prior to implantation for biosafety concerns. Further, it is difficult to noninvasively image and track cells and bioactive factors once they are incorporated into the tissue engineered construct, much less when they are implanted in vivo. Visualization of how the tissue components move and interact is critical to improving our understanding of tissue development. Many biofabrication techniques have been developed to incorporate living cells into functionalized scaffolds in a re- producible, 3D pattern.2,3 Rapid prototyping,4,5 inkjet-based cell printing,6–8 and microcontact printing9,10 are among the commonly used cell deposition systems for tissue engineering applications. These biofabrication methods allow initial deposition of scaffold and cells in a predefined pattern. However, the methods are often expensive, are time consuming, require chemically modified surfaces, or cause cell damage due to high temperatures and pressures used in the deposition process. We developed a direct cell writing system for the freeform construction of biopolymer-based 3D tissue scaffolds and cell-embedded tissue constructs.11 The direct cell writing system uses micronozzles driven by pneumatic microvalves to deposit living cells, scaffold material, and bioactive components such as growth factors in controlled amounts with precise spatial positioning. The system requires no preprocessing, is computer controlled to rapidly produce sample replicates, and operates at room temperature and low pressure to maximize cell viability. Recently, several new approaches have been proposed to actively pattern cell constructs using external forces, including dielectrophoresis,12 an optical trap,13 or superparamagnetic nanoparticles in a magnetic field.14,15 Superparamagnetic iron oxide nanoparticles have been of primary interest for both in vivo and in vitro applications because they exhibit magnetic behavior only in the presence of a magnetic field.16 These nanoparticles can be conjugated with proteins or loaded inside cells, are relatively nontoxic, and can be imaged by magnetic resonance imaging (MRI) or computed tomography (CT). In vivo, superparamagnetic nanoparticles have been used to target drugs to a treatment site to increase drug efficiency and reduce systemic effects,17 to enhance gene delivery to target cells because nanoparticles easily cross cell membranes,18,19 and to detect vascular tissues such as tumors, because iron oxide nanoparticles appear dark on MR images.20,21 In vitro, superparamagnetic nanoparticles have been used to create high resolution, two-dimensional (2D) cell patterns on nonfunctionalized surfaces.14 More recently, Frasca et al. used magnetic fields and magnetic field gradients to achieve 3D cell patterning.15 However, the ability of this technique to create complex 3D shapes is highly limited because the only method of shape control is with a magnetic field gradient from magnets placed under the scaffold material. We propose a technique combining the initial patterning capabilities of the direct cell writing system with the active patterning capabilities of superparamagnetic nanoparticles. This new hybrid technique would allow biofabrication of a complex 3D tissue scaffold of magnetically labeled cells and bioactive factors, which could then be manipulated and tracked within the tissue engineering construct. In this study, superparamagnetic iron oxide nanoparticles were bioprinted either in an alginate scaffold or inside endothelial cells using the multinozzle direct cell writing system. Cell viability was assessed for various nanoparticle and alginate concentrations at a predefined printing nozzle size. The nanoparticles in both alginate and endothelial cells were manipulated using a magnetic field. Finally, the nanoparticles were patterned inside 3D biopolymer scaffolds and imaged using a microCT scanner. Bioprinting of superparamagnetic iron oxide nanoparticles could help create more versatile tissue engineering structures, as well as improve our understanding of cell behavior in 3D tissue culture. Sodium alginate powder (FMCBioPolymer, Drammen, Norway) was dissolved in deionized water at 0.5%, 1%, 2%, and 3% (w=v) concentrations. An ionic crosslinking solution was prepared by dissolving calcium chloride (BDH Chemicals, Poole, UK) in deionized water. NanoArc magnetic iron oxide nanoparticles (Alfa Aesar, Ward Hill, MA) of 20–40 nm diameter were used in all experiments. Spherical nanoparticles of the given size were selected to obtain maximum uptake efficiency.22 Sodium alginate–magnetic nanoparticle solutions were prepared by vigorously mixing sodium alginate with increasing concentrations of iron oxide nanoparticles to achieve a homogeneous nanoparticle distribution. Porcine aortic endothelial cells (PAECs) were isolated by the collagenase dispersion method and maintained in lowglucose Dulbecco's modified Eagle's medium supplemented with 5% fetal bovine serum, 1% penicillin–streptomycin, and 2% glutamine (Invitrogen, Carlsbad, CA). Culture medium was changed every 48 h, and cells between passages 4 and 9 were used. Prior to printing, cells were gently mixed at a concentration of 1.5105 cells=mL in sodium alginate solution to ensure uniform cell distribution. For magnetically labeled cells, PAECs in 100-mm tissue culture dishes were loaded with different nanoparticle concentrations and incubated at 378C in a 5% carbon dioxide incubator for 24 h. Nanoparticle uptake by cell was confirmed by transmission electron microscopy. Our transmission electron microscopy images also suggest that the majority, if not all, of the cells take up nanoparticles. A proprietary solid freeform fabrication-based direct cell writing system (Fig. 1A) was developed to create 3D tissue constructs by dispensing cells and biopolymers into predefined patterns.11,23 The direct cell writing system used in this study operates at room temperature and low-pressure conditions to facilitate deposition of living cells, growth factors, or other bioactive compounds in controlled amounts with precise spatial positioning. Pneumatic microvalves (EFD, East Providence, RI) were used to apply a low printing pressure of 2 and 5 psi to minimize cell death due to the dispensing process11,23 (Fig. 1B). Sodium alginate was chosen as the scaffold biopolymer. Alginate–nanoparticle–cell mixtures with 0, 0.1, or 1.0 mg=mL nanoparticle concentration were printed using 250-mm nozzles. Control samples were dispensed in the system but without using nozzle tips. All samples were dispensed as 0.3 g of bulk material with a sample size of three, and each experiment was repeated a minimum of two times. Data presented are from one representative experiment. After dispensing, each sample was immediately submerged in a 5.0% (w=v) calcium chloride crosslinking solution for 5 min, placed in supplemented media, and returned to the incubator. Samples in the long-term study were crosslinked daily to maintain both cell immobilization and alginate structural integrity. Representative images of printed bulk samples and cell distribution in alginate bulk samples are presented in Figure 1C, D, and E. Alamar blue quantitatively measures cell metabolic activity by using an oxidation–reduction indicator that fluoresces and changes color in metabolically active cells.24 Crosslinked alginate–cell solutions in six-well plates were incubated with 2 mL supplemented media and 200 mL Alamar blue solution (AbD Serotec, Oxford, UK). After 4 h of incubation at 378C in 5% carbon dioxide atmosphere, 100 mL of media from each well was transferred into a 96-well flat-bottomed black assay plate, and fluorescence was measured at 535=590 nm in a GENios microplate reader (Tecan, Ma ̈nnedorf, Switzerland). About 3104 cells were calibrated to a fluorescence intensity reading of 35,000. As the Alamar blue assay measures the mean metabolic activity of the cell population, cell viability was confirmed using a live=dead assay (Invitrogen) as per manufacturer's instructions. iron boron (NdFeB) magnet with a surface field of 6450 Gauss (K&J Magnetics, Jamison, PA) was placed under the 60-mm cell culture dishes. Specific patterns of nanoparticles and magnetically labeled cells were also printed using the cell-dispensing system. A rectangular NdFeB magnet with a surface field of 6450 Gauss (K&J Magnetics) was used to move nanoparticles to a specified location either in a new pattern or while maintaining the original printed pattern. Movement ofmagnetic nanoparticles and themagneticallylabeled cells by the applied magnetic field was imaged using a 4-megapixel CCD camera (Alpha Innotech, San Leandro, CA). A 1.5 mm1.5 mm area of 0.1 mg=mL magnetic nanoparticles was printed within a 5 mm5 mm2 mm of 2% (w=v) alginate construct and imaged using a microCT scanner (SkyScan 1172, Skyscan, Kontich, Belgium). MicroCT allows nondestructive evaluation of the internal structure and composition of the sample based on changes in X-ray absorption. Image resolution was set at 2.16 mm with a 1-mm aluminum filter. The rotation angle was 1808 with a rotation step of 0.18. The samples were statistically compared using Student's t-test. Statistical significance was established at either p < 0.05 (#) or p < 0.01 (*). Two-way analysis of variance (ANOVA) was used to compare changes over time, with statistical significance established at p < 0.0001. Bioprinting magnetic nanoparticles along with cells in a biopolymer scaffold may provide an effective means to track and manipulate bioactive factors in tissue-engineered structures. We now show that while nanoparticles themselves slightly decreased endothelial cell viability, bioprinting had no significant effect (Fig. 2A). At 0 and 12 h after printing, cell viability did not change significantly for unprinted or printed cells with 0 or 0.1 mg=mL nanoparticles in a 1% (w=v) alginate solution. However, at 36 h after printing, PAECs with 0.1 or 1.0 mg=mL nanoparticles were 16% or 35% less viable than cells without nanoparticles, respectively. The viability loss was independent of the printing process. Cell viability continued to decrease with time up to 60 h after cell printing (ANOVA, p < 0.0001). In a long-term assay (Fig. 2B), endothelial cell viability similarly decreased nearly 22% with 1.0 mg=mL iron oxide nanoparticles in the alginate at 72 h after printing compared with samples without nanoparticles (ANOVA, p < 0.0001). No further cell viability decrease was observed from 72 to 144 h, showing that cells maintained their viability following the initial nanoparticle toxicity effect. Increased nanoparticle concentration decreased cell viability, but no additional decrease was observed with printing (Fig. 2A). PAECs encapsulated in alginate with 1.0 mg=mL nanoparticles showed 20% lower viability than cells with 0.1 mg=mL nanoparticles and 36% lower than the control, suggesting a nanoparticle concentration-dependent effect on cell viability. This decreased viability was observed at 36 and 60 h after printing, but the printing process itself did not affect cell viability. To confirm that the Alamar blue measured cell viability, but not a change in cell metabolism, a live=dead assay was performed on printed samples. The live=dead data agreed well with the Alamar blue results. We next investigated whether alginate concentration, which effectively alters biopolymer viscosity, affected printed cell viability. Immediately following printing, there was a 20% viability decrease for cells printed with nanoparticles in 2% (w=v) alginate when compared with the 1% (w=v) alginate (Fig. 3). Twelve hours after printing, lower viability was also observed for control cells with nanoparticles in the 2% (w=v) alginate. This decreased cell viability for cells with nanoparticles in the 2% (w=v) alginate solution was no longer observed at later time points, primarily because cell viability decreased in the samples with nanoparticles in 0.5% or 1% alginate. Interestingly, in cell samples without nanoparticles, cell viability decreased for both control and printed cells without nanoparticles in the 2% (w=v) alginate solution at 36 and 60 h (Fig. 3C, D). Overall, cells without nanoparticles in the 0.5% and 1% (w=v) alginate solutions dem- onstrated an increase in Alamar blue fluorescence over time, which could represent increased cell number or increased cell metabolism. No cell samples in alginate with nanoparticles, and no cell samples in 2% alginate, showed this increase in viability with time. This effect also was independent of printing. Magnetically labeled cells, internally loaded with iron oxide nanoparticles, could be used to track and move cells printed within a tissue-engineered structure. The viability of nanoparticle-loaded cells was examined after printing in 1% alginate, with an initial dispensing pressure of 5 psi. Both control and printed samples without nanoparticles showed increased viability at time points up to 60 h. However, a steep decrease in cell viability was observed from 0 to 36 h for both control and printed cells loaded with either 0.1 or 1.0 mg=mL nanoparticles (Fig. 4A). Printed cells showed the most dramatic change, with a 40% decrease in the Alamar blue fluorescence when compared with printed cells without nanoparticles at 36 h. This viability change was in direct contrast to the lack of printing effect for samples with nanoparticles in the alginate. Although early cell viability was significantly decreased, there was no significant change at time points after 36 h, suggesting stabilization of the remaining cell population. When printing pressure was decreased to 2 psi, cell viability increased almost 20% (Fig. 4B). The nanoparticles were magnetically manipulated within the alginate to determine if nanoparticles could be used to move bioactive factors after printing. The 1.0 mg=mL nanoparticles were homogenously distributed in 1%, 2%, and 3% (w=v) alginate, printed in bulk samples, and left as a viscous liquid or crosslinked with calcium chloride to form a gel (Fig. 5A, C, E, G, I, K). The nanoparticles printed in either 1% or 2% (w=v) alginate without calcium chloride moved toward the NdFeB magnet placed under the cell culture dish within a minute (Fig. 5B, D; arrows indicate nanoparticles at the magnet edge). However, no nanoparticle movement was observed in the 3% (w=v) alginate solution, likely because of the high alginate solution viscosity (Fig. 5F). When the samples were crosslinked with calcium chloride, nanoparticles similarly moved toward the magnet edge in the 1% and 2% (w=v) alginate, but not in the 3% alginate (Fig. 5H, J, L). However, the nanoparticles moved more slowly, and less spatial repositioning of nanoparticles was observed. We next investigated if cells loaded with magnetic nanoparticles could be moved within the alginate biopolymer. PAECs magnetically labeled with nanoparticles were initially homogenously distributed in 0.5% and 1% (w=v) alginate (Fig. 6A, E, I; higher magnification in Fig. 6B, F, J). Magnetically labeled cells moved toward the NdFeB magnet placed under the cell culture dish (Fig. 6C, G, K). At higher magnification, individual cells were seen at the magnet edge (arrows in Fig. 6D, H, L). Isolated nanoparticles can also be seen in the alginate, which are likely artifacts of incomplete nanoparticle removal from the cell solution when it was mixed with alginate. Magnetically labeled cells continued to cluster at the magnet edge in the crosslinked alginate, but no movement was observed in alginate concentrations higher than 1%. Specified patterns of nanoparticles and magnetically labeled cells were printed and moved using a magnetic field. The 1% alginate with iron oxide nanoparticles was printed in a pattern (Fig. 7A), and a magnetic field was used to move the nanoparticles to the printed pattern tips (Fig. 7B). The basic shapes (lines and rectangles) of either nanoparticles (Fig. 7C, D, G, H) or magnetically labeled cells (Fig. 7E, F) were moved to new locations while maintaining the original pattern. Magnetic nanoparticles printed within 3D alginate scaffolds were imaged by microCT to determine if nanoparticle printing would allow noninvasive tracking of bioactive factors and cell location in tissue-engineered structures. A nanoparticle– alginate prepolymer solution was encapsulated in alginate biopolymer solution using layer-by-layer deposition with the solid freeform fabrication-based direct cell writing system. Printed nanoparticle clusters were clearly visible by microCT scanning of the 3D tissue scaffold (Fig. 8, arrows). Enhanced nondestructive imaging of cellular and biochemical interactions within 3D tissues would advance the knowledge of tissue development, and the ability to precisely pattern cells and bioactive factors throughout the tissue growth process would improve fabrication of complex tissues. A combination of bioprinting, which allows initial patterning, and superparamagnetic nanoparticles, which allow tracking and repatterning, could help realize these tissue engineering goals. We now show that magnetic nanoparticles can be bioprinted for tissue engineering applications. Printing nanoparticles in the alginate biopolymer did not impact cell viability more than the presence of nanoparticles. Further, while printing cells loaded with nanoparticles did decrease cell viability, the viable cells were stabilized shortly after printing. Magnetically labeled biofactors and cells could be moved within the alginate structures in the presence of an externally applied magnetic field, or imaged nondestructively using microCT. Sodium alginate was used in all experiments because it is nontoxic, remains as viscous liquid at room temperature, and crosslinks to form a gel under mild conditions. However, endothelial cells are unable to specifically interact with alginate, which prevents cell anchorage and attachment in the biopolymer.25 The cells have a rounded morphology because they are encapsulated in, not adhered to, the polymer scaffold. As the printing system operates at room temperature, we are currently unable to print polymers such as collagen, to which the cells would attach, because collagen solidifies into a stiff gel at room temperature. In our experiments, alginate is advantageous because it maintains cell number without proliferation, which allows improved observation of cell death. In our future work, we will incorporate a cooling system into our bioprinting device to print collagen gels. We will then be able to study cell viability and proliferation after cells printed with nanoparticles have attached to the scaffold. By bioprinting nanoparticles in the biopolymer, bioactive factors such as growth factors, antibodies, drugs, and genes conjugated to the magnetic nanoparticles can be precisely patterned within a 3D scaffold. Chemical coupling via amide or ester bonds has been used by others to conjugate bioactive factors to iron oxide nanoparticles. Linker molecules, including 1-ethyl-3-(3-dimethylaminopropyl) carbodi-imide hydrochloride (EDCI), N-succinimidyl 3-(2-pyridyldithio) propionate, or N-hydroxysuccinimide were used to attach targeting ligands and proteins to nanoparticle surfaces.16 Insulin, lactoferrin, and ceruloplasmin were successfully conjugated to superparamagnetic iron oxide nanoparticles using EDCI. These nanoparticles were then targeted to cell surface receptors, thereby avoiding endocytosis, to achieve tissue and cell-specific drug targeting.26,27 Magnetic nanoparticles have also been dually conjugated, both with a ligand specific for a target cancer cell receptor and a cancer therapy drug. For instance, a radiolabeled anti-vascular endothelial growth factor monoclonal antibody was conjugated to magnetic nanoparticles to both target and deliver radiation therapy to liver cancer.28 By chemically conjugating bioactive factors to the nanoparticles, it would be possible to initially pattern the bioactive factors in the scaffold and then move them during tissue growth. This technique could, for example, provide endothelial cells with a changing growth factor gradient to promote angiogenesis. In this study, we observed decreased viability for cells exposed to both low and high nanoparticle concentrations in the alginate, independent of bioprinting. However, in our previous studies, endothelial cell viability was largely preserved in 2D culture up to an iron oxide nanoparticle concentration of 0.5 mg=mL.29 This apparent contradiction might be related to differences in 2D and 3D endothelial cell culture. In 2D cultures, endothelial cells form a stable, confluent monolayer, whereas the same cells form tubes when grown in a 3D matrix.30 The 3D angiogenic endothelial cell phenotype may be more susceptible to nanoparticle toxicity than the 2D monolayer. Cells may be able to uptake more nanoparticles in the 3D alginate structure because they are exposed to nanoparticles on all sides. In 2D culture, cells interact with nanoparticles only at their apical surface. The effect could also be specific to the alginate scaffold because cells do not attach to the alginate. These attachment-dependent cells may experience changes in critical cell functions when encapsulated, which coupled with functional changes caused by nanoparticle uptake may enhance nanoparticle cell toxicity.31 Interestingly, long-term tests showed that cell viabil- ity is stable after 72 h, perhaps because all available nanoparticles in the vicinity of a cell have already been taken up. The printing process itself did not impact endothelial cell viability when nanoparticles were mixed with the alginate. We hypothesize that cell viability is preserved because nanoparticles are free to move within the alginate without damaging cells during printing. In direct contrast, when nanoparticles were loaded inside cells, there were fewer viable printed cells than control cells but this decrease in viability was found to be related to the printing pressure. The system operates at printing pressures ranging from 1 to 40 psi. The majority of our experiments were conducted at a printing pressure of 5 psi, which was shown in previous studies to maintain cell viability. Only at pressures greater than 20 psi did cell viability decrease.23 As dispensing pressure was lowered to 2 psi, the forces imposed on the cell as it moved through the nozzle decreased, which likely increased cell viability. However, the printing process at 2 psi dispensing pressure took more than twice as long, and so, for automated mass production, higher printing pressures such as 5 psi might still be preferred with the known loss in cell viability. At both pressures, cells without magnetic nanoparticles recovered in time from the mechanical perturbation and eventually reached a steady-state condition. When cells uptake nanoparticles, they form vacuoles which disrupt the cell cytoskeleton.26 The cells with a disrupted cytoskeleton may experience increased damage due to printing-induced forces. Alternatively, the nanoparticles inside cells may be more likely to damage the cell membrane through direct shear effects during the printing process. As the cells are pushed through the nozzle, the nanoparticles may break through the cell membrane or the nucleus, causing irreversible damage. It is also possible that the cells loaded with nanoparticles are already in a state of internal stress, perhaps because of reactive oxygen species. Intracellular reactive oxygen species generation is hypothesized to increase with nanoparticle uptake, leading to protein, DNA, and tissue injury and eventually cell death.32 The addition of mechanical stress due to the bioprinting process may be more toxic when cells are already biochemically stressed by the internalized nanoparticles prior to the printing process. Both printing parameters and nanoparticle conditions will need to be optimized to minimize cell death. Cell toxicity could alternatively be decreased while still allowing cell tracking, by attaching bioconjugated nanoparticles to the cell membrane. However, it might be difficult to manipulate cells with externally attached nanoparticles. The nanoparticles may detach during printing because of mechanical forces as the cell moves through the nozzle, or the nanoparticles may detach during cell movement through the scaffold because of viscous drag. Further, attaching nanoparticles to cells via a surface receptor could activate unwanted intracellular signaling, or prevent the cell from using that receptor to perform a particular function. Keeping these challenges in mind, in our future work, we will investigate whether printing and moving cells with membrane-attached nanoparticles is feasible and maintains cell viability. Our studies revealed that cell viability decreased for samples printed in 2% (w=v) alginate when compared with 1.0% (w=v) alginate with and without any nanoparticles. The high solution viscosity may have exposed cells to higher printing forces during the cell-dispensing process. Cell membranes are highly fragile to mechanical loads, and excessive membrane perturbation can lead to cell death.33 The initial cell viability change observed from 0 to 12 h in samples with nanoparticles in 2% alginate suggests that nanoparticles further increased the biopolymer viscosity. At later times (36 and 60 h after printing), the presence of nanoparticles inside the alginate overcame this initial cell viability decrease in the 2% alginate, and there was no difference among the different alginate samples. Even the cells without nanoparticles demonstrated decreased viability in the long term because of the 2% alginate, which was in contrast to the recovery process for cells in 1% alginate. This suggests that endothelial cell health is compromised in the stiffer gel; however, additional studies are needed for confirmation. Although the cells in our experiments showed lower viability in stiffer gels, other studies have shown that cells prefer stiffer substrates. In 2D culture, cells form large, stable focal adhesions on stiff substrates, whereas cells form irregularly shaped, dynamic adhesions on softer substrates.34 However, for 3D cell studies, cell migration speed and viability may depend not only on the substrate stiffness but also substrate adhesivity or cell–matrix adhesion availability.35 Peyton and Putnam36 found that when cell adhesiveness was reduced using an integrin-blocking antibody, the maximum cell migration speed shifted from stiffer to softer Matrigel substrates. Our printed cells are in 3D alginate gels to which they do not attach, and therefore, it is possible that unattached cells are more viable in softer alginate gels. There are rich opportunities for future studies to address mechanisms underlying these distinct differences in cell viability and migration in 2D and 3D environments and the relationship between substrate stiffness and cell–matrix adhesion. The hybrid nanobioprinting system allows initial nanoparticle and cell patterning by computer-controlled printing, after which the nanoparticles and cells can be moved to a new location either in the initial pattern or in a new pattern defined by the magnetic field. We now also showed that nanoparticle movement depends on alginate viscosity. The nanoparticles moved toward the NdFeB magnet in 1% and 2%, but not 3% alginate. Movement of a single spherical magnetic nanoparticle at steady state in an external magnetic field is driven by the force due to the magnetic field gradient and opposed by the force due to viscous drag,37,38 which are given as follows: where m, B, Z, d, and v are the nanoparticle net magnetic moment, magnetic field, suspending medium viscosity, nanoparticle diameter, and instantaneous nanoparticle velocity, respectively. Considering a one-dimensional problem along the centerline of the magnet (x axis) at steady state, a force balance between equations (1) and (2) leads to a velocity given as Magneticallylabeling cells allows nondestructive imaging by microCT in the 3D tissue-engineered scaffold. The intracellular nanoparticle loading should be optimized to obtain maximum signal intensity while at the same time protecting cell viability. Our studies showed that it may be possible to qualitatively determine the nanoparticle or magnetically labeled cell density at a given location by microCT signal intensity. A sixfold increase in signal intensity was observed for 100,000 cells loaded with 1.0 mg=mL iron oxide nanoparticles when compared with the same cell number loaded with 0.1 mg=mL nanoparticles. However, excess nanoparticles that were not taken up by cells could also increase the microCT signal intensity. Nanoparticle loading parameters should be selected to minimize free nanoparticles, and any excess nanoparticles should be thoroughly washed away prior to cell printing. Magnetically labeled dead cells will light up on the microCT scans, which may decrease live cell tracking efficacy. An additional assay, such as Alamar blue or live=dead, may be needed to differentiate live from dead cells. Live magnetically labeled cells would also respond to biochemical signals by proliferating, moving, and forming 3D structures, which should help distinguish them from dead cells in microCT images. Although cell viability was decreased with nanoparticle loading and printing, there is still potential by optimizing nanoparticle concentration and printing parameters to bioprint magnetically labeled cells. In tissue engineering, the development of complex 3D tissues requires various cell types, such as smooth muscle and endothelial cells for vascular systems or hepatocytes and sinusoidal endothelial cells for liver. However, cell–cell interactions are difficult to manipulate in coculture systems, even in 2D cultures. Ito et al.39 used magnetic force to place magnetically labeled cells onto target cells and promote heterotypic cell–cell adhesion. The solid freeform cell writing system could enable assembly of 3D patterned tissue engineering constructs with various magnetically labeled cell types. Through microCT, both bioactive factors and cells could be noninvasively imaged within the tissue-engineered scaffold, which would allow longitudinal studies of tissue development. This hybrid nanobioprinting technique, which uses a combination of bioprinting and active magnetic patterning, could dramatically impact our ability to understand and recreate complex tissue development. We have shown that a solid freeform fabrication system can be used to create magnetically functionalized 3D tissue scaffolds, which can be manipulated after printing using a magnetic field. This system could be used to add bioactive factors or specific cell types into a precisely patterned tissueengineered scaffold, and later to move or remove these agents. Additionally, the bioactive agents or cells could be imaged within the tissue by MRI or CT. In our future work, we will improve printed magnetically labeled cell viability by exploring different nanoparticle sizes and shapes, as well as attaching nanoparticles to the cell membrane. The nanoparticles will be conjugated with bioactive factors, and cells will be printed in collagen gels with a cooled printing system. Finally, complex magnetic arrangements will be used to control cell movement and patterning after printing in the tissue-engineered construct. This work was partially supported by the Mechanical Engineering and Mechanics Department of Drexel University. The authors thank Wonjin Jo for her assistance with the solid freeform fabrication cell writing system. No competing financial interests exist. as a surface ligand to prevent endocytosis. IEEE Trans Nanobiosci 2, 255, 2003. Address correspondence to: Alisa Morss Clyne, Ph.D. MEM Office 3141 Chestnut St. Philadelphia, PA 19104 E-mail: Received: August 23, 2009 Accepted: September 18, 2009 Online Publication Date: November 3, 2009 </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 12.0, \"mgL\": null, \"viab\": 125.0}, {\"h\": 36.0, \"mgL\": null, \"viab\": 104.96454}, {\"h\": 60.0, \"mgL\": null, \"viab\": 96.4539}, {\"h\": 12.0, \"mgL\": null, \"viab\": 97.87234}, {\"h\": 36.0, \"mgL\": null, \"viab\": 81.56029}, {\"h\": 60.0, \"mgL\": null, \"viab\": 74.11347}, {\"h\": 24.0, \"mgL\": null, \"viab\": 84.69324}, {\"h\": 48.0, \"mgL\": null, \"viab\": 74.03898}, {\"h\": 72.0, \"mgL\": null, \"viab\": 67.576324}, {\"h\": 96.0, \"mgL\": null, \"viab\": 68.59861}, {\"h\": 144.0, \"mgL\": null, \"viab\": 64.980354}]}", "role": "assistant" } ]	142.json
[ { "content": "Extract sample context and measurement list. <doc> International Journal of Molecular Sciences ISSN 1422-0067 www.mdpi.com/journal/ijms OPEN ACCESS Article Masatoshi Watanabe 1,, Misao Yoneda 2 , Ayaka Morohashi 1 , Yasuki Hori 1 , Daiki Okamoto 1 , Akiko Sato 1 , Daisuke Kurioka 1 , Tadashi Nittami 1 , Yoshifumi Hirokawa 2 , Taizo Shiraishi 2 , Kazuaki Kawai 3 , Hiroshi Kasai 3 and Yukari Totsuka 4 Received: 8 June 2013; in revised form: 8 July 2013 / Accepted: 18 July 2013 / Published: 25 July 2013 CD44+ fraction of A549 cells. MgNPs-Fe3O4 induced the expression of heme oxygenase-1 at a concentration of 1 μg/mL, and in a dose-dependent manner. Despite these effects, MgNPs-Fe3O4 had minimal effect on cell viability and elicited only a small increase in the number of cells undergoing apoptosis. Together, these data suggest that MgNPs-Fe3O4 exert little or no cytotoxicity until a high exposure level (100 μg/mL) is reached. This dissociation between elevated indices of cell damage and a small effect on cell viability warrants further study. Keywords: magnetic nanoparticles; cytotoxicity; genotoxicity; A549; CD44 Nanotechnology—the manipulation and production of matter sized between 1 and 100 nm—has grown markedly with the promise of substantial benefits and applicability to such diverse areas as clothing, electronics, engineering, and healthcare [1]. The principal goal of nanotechnology is to develop new materials in the nanometer scale, including nanoparticles, defined as particulate materials with at least one dimension of less than 100 nm. The design and development of nanomaterials have been of fundamental importance to the industry, given their novelty and the benefits conferred by their physicochemical properties. Studies regarding the toxicological impact of MgNPs-Fe3O4 have yielded disparate results, depending on the cell type, surface modification, cell medium composition, protein-MgNP interaction, and oxidation state of iron [7,14]. We evaluated the cytotoxic effects of MgNPs-Fe3O4 in A549 cells. We report that MgNPs-Fe3O4 caused LDH leakage only at a concentration of 100 μg/mL; increased ROS production and 8-OH-dG content, and decreased glutathione (GSH) levels were found with 10 μg/mL MgNPs-Fe3O4. Despite these responses, MgNPs-Fe3O4 caused only a small decrease and increase in cell viability and apoptosis, respectively. Together, these data suggest that MgNPs-Fe3O4 agglomerate at a high concentration. The presence of FBS appeared to enhance the stability of MgNPs-Fe3O4 in suspension. These data are consistent with a previous report showing that MgNPs show increased stability against aggregation in the RPMI-1640 with an increasing amount of FBS [15]. Therefore, the influence by the sedimentation rate of the secondary nanoparticles (NPs) and rations of protein to NPs could be taken into consideration in the in vitro toxicity of NPs. These results shows the hydrodynamic sizes of secondary nanoparticles in Ham's F-12 medium with 10% FBS used in this study. A representative micrograph shows that after 24 h, MgNPs-Fe3O4 aggregate within intracellular vesicles in A549 cells (Figure 2a). Figure 2b shows the flow cytometric light scatter histograms of the cells treated with the 0, 1, 10, or 100 μg/mL MgNPs-Fe3O4. The forward-scattered (FS) intensity (reflective of cell size) did not change; conversely, side-scattered (SS) intensity (reflective cellular uptake) increased in a dose-dependent manner. That is, the cells, which took up higher doses of MgNPs showed higher intensities of SS. Treatment with MgNPs-Fe3O4 for 24 h did not affect cell viability as assessed by the Alamar Blue assay. However, treatment with 100 μg/mL MgNPs-Fe3O4 for 72 h caused a significant reduction in cell viability (Figure 3). Significant LDH leakage was detected following treatment with 100 μg/mL MgNPs-Fe3O4; lower concentrations had no effect (Figure 4). As shown in Figure 5a, treatment with 100 μg/mL MgNPs-Fe3O4 for 24 h caused a small but significant increase in the percentage Annexin V-staining cells; however, these values were greatly below that caused by H2O2 (Figure 5b). Figure 5. Effect of Fe3O4 magnetic nanoparticles (MgNPs-Fe3O4) on apoptosis in A549 cells. A549 cells were treated with 0 (control), 1, 10 or 100 μg/mL MgNPs-Fe3O4 for 24 h; cells were treated with 5 mM H2O2 for 24 h as a positive control. Apoptosis of A549 cells treated with MgNPs-Fe3O4 or H2O2 was determined by flow cytometry based on propidium iodide/Annexin V staining patterns; (a) Representative flow cytometry of one set of triplicate experiments; (b) Percentages of apoptotic cells from flow cytometry analysis. Apoptotic cells include early apoptotic cells (AnnexinV+/PI−) and late apoptotic or necrotic cells (AnnexinV+/PI+). Data are presented as the mean ± SD of three independent experiments. p < 0.05 vs. control. As shown in Figure 9, the transcript level of the HO-1 was induced in a dose-dependent manner after 12 and 24 h of MgNPs-Fe3O4 exposure, however its transcription level at 100 mg/mL exposure after 24 h was reduced compared to after 12 h. Oxidative stress is caused by an imbalance in the level of ROS and a biological system's ability to detoxify the reactive intermediates [16]. Cells possess both enzymatic and non-enzymatic mechanisms to counterbalance the cytotoxicity and genotoxicity caused by ROS [16]. In the lungs, the major enzymatic antioxidants are superoxide dismutases (SODs), catalase, and glutathione peroxidase (GSH-Px); others include those examined in this study, HO-1, thioredoxin (TR), and glutaredoxin (GLRX). HO-1 is involved in playing a major role in degradation of heme to biliverdin, but has recognized potent anti-inflammatory and anti-apoptotic effects [17,19]. HO-1 is induced mainly at the transcriptional level by oxidative stress, pro-inflammatory mediators, and some growth factors [18]. HO-1 mRNA expression is known to mediate antioxidant and cytoprotective effects and has been considered useful as a marker for particle-induced oxidative stress. Park et al. [20] showed that treatment of a human bronchial epithelial cell line with TiO2-MgNPs for four hours caused dose-dependent increases in mRNA expression of HOG-1, glutathione-S-transferase, and catalase; mRNA expression level of HO-1 had returned to baseline by 24 h [20]. Napierska et al. [21] also showed a marked induction of HO-1 mRNA in the endothelial cell at six hours after treatment of SiO2-NPs, but reduction of HO-1 mRNA at 24 h. Our results appear to be same as these two studies. MgNPs-Fe3O4 caused a dose-dependent reduction in the CD44+ subpopulation (Figure 10). CD44 is a cell surface glycoprotein that mediates cellular adhesion to the extracellular matrix and is involved in multiple processes, including inflammation, cell migration, signaling, and tumor metastasis [13,22]. CD44 is up-regulated in the damaged epithelium of asthma patients, and is believed to be involved in tissue repair by localizing chemokines and growth factors to the disrupted epithelium [23]. CD44 is also a marker of certain cancer stem cells [24], in which it functions to defend cancer cells against oxidative stress by increasing GSH synthesis [25]. CD44 has also been reported to be involved in the protective effect of hyaluronate on constitutive DNA damage by ROS in A549 cells [26]. Consistent with the previously noted reduction in GSH and increase in 8-OH-dG levels, we found that MgNPs-Fe3O4 markedly decreased the CD44+ cell fraction of A549 cells. Thus, these results highlight another mechanism by which MgNPs-Fe3O4 impair redox control and damage DNA in A549 cells. Our results also offer the possibility that CD44 may be a marker MgNP-Fe3O4-induced cytotoxicity; however, further study is warranted. MgNPs-Fe3O4 were sterilized by ultraviolet (UV) irradiation and suspended in phosphate-buffered saline (PBS), Ham's F-12 alone, and Ham's F-12 medium containing 10% fetal bovine serum (FBS) and 100 U/mL penicillin-streptomycin to yield a concentration of 1, 10 or 100 μg/mL. Suspensions were sonicated at 30 W for 10 min using an Ultrasonic HomogenizerVP-050 (TAITAEC, Koshigaya, Saitama, Japan). A549 human lung epithelial cells were purchased from American Tissue Type Culture Collection (Manassas, VA, USA). Cells were incubated in Ham's F-12 Medium with 10% fetal bovine serum (FBS) and 100 U/mL penicillin–streptomycin in 5% CO2 at 37 °C. Cells were maintained at a density of 60%–70% confluence in 100 mm dishes, and used in log-phase of growth. The cellular uptake of MgNPs-Fe3O4 in A549 cells was analyzed as follows. A549 cells were fixed with 3% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.3) at 4 °C for 4 h. Samples were post-fixed with 2% osmium tetraoxide at 4 °C for 2 h, dehydrated, and embedded in epoxy resin. Ultrathin sections (80 nm) were then stained with uranyl acetate and lead citrate, and observed by TEM. bottom 96-well culture plate containing 100 μL reaction mixture, and incubated for 30 min at room temperature. Formazan absorbance—an index of the number of lysed cells—was measured by a microplate reader at 500 nm (Viento XS, DS Pharma Biomedical, Osaka, Japan). A549 cells (1.0 × 106 cells) were cultured on 100-mm culture dishes, and treated with 0, 1, 10 or 100 μg/mL MgNPs-Fe3O4 for 24 h at 37 °C. Cells were harvested, washed gently with PBS, collected by centrifugation, and then stained using an Annexin V-FITC Kit (Beckman Coulter, Marseille, France) following the manufacturer's instructions. Cells were stained with Annexin V and propidium iodide (PI, Sigma-Aldrich, St. Louis, MO, USA), and analyzed by flow cytometry (Becton Dickinson, Franklin Lakes, NJ, USA) within 1 h of staining using the FL1 (FITC) and FL3 (PI) lines. ROS were measured using the CM-H2DCFDA assay (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. Cells (1.0 × 104 cells/well) in 24 well-plates were treated with 0, 1, 10 or 100 μg/mL MGNPs-Fe3O4 for 24 h at 37 °C. A fresh stock solution of 5 mM CM-H2DCFDA was prepared in DMSO and diluted to a final concentration of 1 μM in PBS. Cells were washed with PBS, followed by incubation with 50 μL of working solution of the fluorochrome marker CM-H2 DCFDA for 30 min. Fluorescent images were obtained using an IX2N-FL-1 microscope (Olympus, Tokyo, Japan), and analyzed using imaging soft (Photoshop Elements 8, Adobe systems, Tokyo, Japan). The data were expressed as percentage of the unexposed control. Intracellular GSH level was determined using a GSH-Glo Glutathione assay kit (Promega, Madison, WI, USA) according to the manufacturer's instructions. Briefly, cells were seeded in 96-well plates and treated with 0, 1, 10 or 100 μg/mL MgNPs-Fe3O4 for 24 h at 37 °C. The cells were washed with DPBS, and the GSH-Glo reagent was added to each well for 30 min at room temperature to allow the cells to convert a luciferin derivative into luciferin. Reconstituted luciferin detection reagent was then added to each well for 15 min, and the luminescent signal was measured with a Glomax multi detection system (Promega, Madison, WI, USA). A549 cells were treated with 0, 1, 10, or 100 μg/mL MgNPs-Fe3O4 for 24 h at 37 °C. Total RNA was isolated using ISOGEN (Nippon Gene, Tokyo, Japan), and cDNA was produced using a mixture containing Superscript RNase H Reverse Transcriptase (Invitrogen, Carlsbad, CA, USA), oligo dT primer, and 2.5 mmol/L dNTP. Quantitative real-time PCR was conducted using the LINE GENE real-time PCR detection system (BioFlux, Tokyo, Japan) with the SYBR Premix Ex Taq Perfect Real Time Kit (Takara Bio. Inc., Otsu, Japan). The PCR reaction consisted of initial thermal activation at 95 °C for 10 s and 40 cycles. Each cycle was as follows: 95 °C for 5 s; 60 °C for 26 s. PCR producs were verified by analysis of heat-dissociation curves and amplification plots. Quantitative values were acquired from linear regression of the PCR standard curve. The primer sequences of the amplified genes are as follows [28,29]; Heme oxygenase-1, forward 5'-GGTGATAGAAGAGGCCAAGAC-3' and reverse 5'-GCAGAATCTTGCACTTTGTTG-3', β-actin, forward 5'-GGATGCAGAAGG AGATCACTG-3' and reverse 5'-CGATCCACACGGAGTACTTG-3'. A549 cells were treated with 0, 1, 10 or 100 μg/mL MgNPs-Fe3O4 for 24 h at 37 °C. Cells were then labeled in a PBS solution with a mouse anti-human CD44 monoclonal antibody conjugated with fluorescein isothiocyanate (clone SFF-2, Millipore, Billerica, MA, USA) for 1 h at room temperature. A mouse IgG immunoglobulin and dye conjugate IgG was used as control for non-specific binding. Flow cytometric analysis was performed with a Guava-EasyCyte*HT using Guava Express Pro software (Millipore, Billerica, MA, USA) gating for CD44+ cells. A minimum of 10,000 cells was measured per sample. Data are presented as the mean ± standard deviation (SD). Differences between treated and untreated control cells were determined using one-way ANOVA followed by Dunnett's test. Differences were considered statistically significant at p < 0.05. MgNPs-Fe3O4 up to a concentration of 100 μg/mL exerted minimal effect on viability of A549 cells, despite causing a significant reduction in antioxidant capacity and an increase in oxidative damage to DNA. Increased expression of an oxidative stress-related gene was not sufficient to prevent the decrease in GSH content. The decrease in the CD44+ cell fraction was consistent with the observed drop in GSH concentration and increase in 8-OH-dG level. This research was supported in part by a Grant-in-Aid for the Global COE Program from the Ministry of Education, Culture, Sports, Science and Technology of Japan, a Grant-in-Aid for Research on Risk of Chemical Substances from the Ministry of Health, Labour and Welfare of Japan, and a Research Grand-in-Aid from Magnetic Health Science Foundation. The authors report no conflict of interest. The authors are responsible for the content and writing of the paper. 2013 by the authors; licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution license (http://creativecommons.org/licenses/by/3.0/). </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 96.346596}, {\"h\": 72.0, \"mgL\": null, \"viab\": 95.28805}, {\"h\": 24.0, \"mgL\": null, \"viab\": 95.04239}, {\"h\": 72.0, \"mgL\": null, \"viab\": 91.270515}, {\"h\": 24.0, \"mgL\": null, \"viab\": 93.7379}, {\"h\": 72.0, \"mgL\": null, \"viab\": 88.29703}, {\"h\": 24.0, \"mgL\": null, \"viab\": 97.32284}, {\"h\": 24.0, \"mgL\": null, \"viab\": 91.02362}, {\"h\": 24.0, \"mgL\": null, \"viab\": 99.49775452}, {\"h\": 24.0, \"mgL\": null, \"viab\": 99.48388338}, {\"h\": 24.0, \"mgL\": null, \"viab\": 98.21391782}]}", "role": "assistant" } ]	140.json
[ { "content": "Extract sample context and measurement list. <doc> J ournal of Nanomedicine & Praveen Sonkusre, Ravikanth Nanduri, Pawan Gupta and Swaranjit Singh Cameotra* Institute of Microbial Technology, Sector 39 A, Chandigarh 160036, India Selenium is reported to have excellent anticancer activity at higher doses. However, due to its low therapeutic index, it is desirable to have a selenium form with lesser toxicity even at high therapeutic dose; alternatively it would be progressive to attempt efficacious preparation at very low concentration. In this study, selenium nanoparticles were synthesized biologically by Bacillus licheniformis JS2, and a method was developed for extraction and purification of intracellular nanoparticles. Characterization of extracted nanoparticles for shape, size, and purity indicated that the particles were spherical with diameters ranging between 40 and 180 nm, composed of selenium and capped with a few functional groups that provide steric stability to them. These neutral charged, non-agglomerating selenium nanoparticles at a concentration as low as 2 μg Se/mL were efficacious in inhibiting proliferation and inducing caspase independent necrosis to human prostate adenocarcinoma cells (PC3) without causing any significant toxicity to human peripheral blood mononuclear cells. Keywords: Bacillus licheniformis JS2; Steric stability; Selenium nanoparticles; Necrosis; PC3; Human peripheral blood mononuclear cells Selenium is an essential trace element that plays several crucial roles in mammalian physiology [1]. As an integral part of glutathione peroxidase and thioredoxin reductase enzyme, it works as a potent antioxidant agent and has been found to be involved in thyroid hormone metabolism and fertility improvement. Evidence suggests that selenium also plays a role in the prevention of several diseases including cardiovascular disease, arthritis, muscular dystrophy, and cystic fibrosis; in addition it is known to have anticancer activity [2]. Due to low therapeutic index of organic and inorganic selenium, the use of selenium in nanoparticle (NP) form is gaining a great deal of attention [3,4]. Chemically synthesized selenium nanoparticles (SeNPs) have been studied as a potential cancer therapeutic agent and drug carrier [5,6] with excellent antioxidant activity [7,8]. They appear to have physiological properties similar to those of other selenium compounds, such as sodium selenite, Semethylselenocysteine, and selenomethionine (SeMet) [2,3,7], although other physiological activities need to be studied further. It has been also reported that SeNPs have better bioavailability and less toxicity than other organic and inorganic selenocompounds [5,7,9]. Because of key advantages over chemical and physical synthesis, biological synthesis of NPs is rapidly becoming an important field for study. Biological synthesis is clean, non-toxic, and eco-friendly; additionally it utilizes less energy than chemical and physical synthesis with an efficient use of material without generation of harmful by-products [10]. In addition, there are two other major advantages of biologically synthesized NPs over those chemically synthesized: increased biocompatibility and much greater stability [11]. Many reports have demonstrated the biosynthesis of various metal and metalloid NP by bacteria, fungus, and other microorganisms, under metal or metalloid salt stress, respectively. Bacteria are known to produce metal, metalloid, and metal oxide NPs with various structures compositions, sizes, and densities. Many of these organisms synthesize NPs intracellularly but few extracellularly [11-16]. If fully developed, these biological systems could be utilized commercially for nanomaterial synthesis [17]. Unfortunately, the purity of the particles from this eco-friendly synthesis process can be questionable. Since these particles are synthesized by microorganisms, there is a high possibility that the NPs produced are associated with the microorganism itself, various microbial cellular components, or both [15,16,18]. To overcome this problem, it is paramount to investigate improved extraction protocols. Recent efforts are focused on bacteria that synthesize NPs extracellularly or have used bacterial supernatants containing reductase enzymes and probably some other factors responsible for metal reduction and NP synthesis so that particles can be easily extract and are less likely to be associated with microbial components [15,19,20]. Many bacterial species have been reported to synthesize NPs very q u i c k l y and efficiently but intracellularly. However, because of the current unavailability of an efficient method for intracellular NP recovery and associated agglomeration issues [21], researchers cannot utilize the potential of these highly efficient bacteria and are more focused on extracellular NP-synthesizing microorganisms. Here, we report a novel method for the extraction and purification of intracellular selenium NPs from the Gram positive bacteria Bacillus licheniformis JS2. We have developed a new procedure that utilizes lysozyme and a French press for complete bacterial cell lysis. This protocol is very efficient and allows the recovery of pure and clean intracellular SeNPs. Additionally, we studied the effect of these biogenic SeNPs on human prostate adenocarcinoma cell line PC3 and observed excellent necrosis inducing ability at very low selenium concentration without a significant effect on human peripheral blood mononuclear cell (hPBMC) viability. Corresponding author: Swaranjit Singh Cameotra, Institute of Microbial Technology, Sector 39 A, Chandigarh 160036, India, Tel: 0172-6665223; Fax : 0172-2690632/585; E-mail: Received March 13, 2014; Accepted April 09, 2014; Published April 15, 2014 Citation: Sonkusre P, Nanduri R, Gupta P, Cameotra SS (2014) Improved Extraction of Intracellular Biogenic Selenium Nanoparticles and their Specificity for Cancer Chemoprevention. J Nanomed Nanotechnol 5: 194. doi:- 7439.1000194 : 2014 Sonkusre P, et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited. Bacterial strain Bacillus licheniformis JS2, isolated from seleniferous agricultural soil from Jainpur Village located in the Nawahshahr district (latitude 31°07' N and longitude 76°08' E) of Punjab, India, was used for SeNP synthesis under aerobic conditions. Bacteria was routinely cultivated and maintained on tryptic soya broth and agar plates at 37°C [22]. Tryptic soya broth and agar were purchased from Hi-Media. Lysozyme, lauryl-sulfate (sodium dodecylsulfate, SDS), absolute ethanol, Histopaque-1077, and menadione were obtained from Sigma. Tris-buffer, HCl, 1-octanol, and chloroform were purchased from Merck. A silver staining kit for protein detection was purchased from Pierce Thermo Scientific, and the manufacturer's protocol was followed. Type II Millipore water was used in all experiments. RPMI medium 1640, F12K nutrient mixture medium, fetal bovine serum (FBS), and penicillin-streptomycin solution (Pen-Strep) were procured from Invitrogen for mammalian cell growth, maintenance, and experimental setup. All plastic ware for cell culture was purchased from Nunc. XTT sodium salt for the cell viability assay was purchased from Santa Cruz Biotechnology. L-selenomethionine and an AnnexinV-FITC apoptosis detection kit were purchased from Calbiochem, and the manufacturer's protocol was used. ApoTox-Glo Triplex assay kit was purchased from Promega and the manufacturer's protocol was followed. Bacillus licheniformis JS2 was used to synthesize SeNPs aerobically under 1.8 mM sodium selenite stress. A bacterial culture at 0.02 OD600 was allowed to grow for 15 h at 200 RPM at 37°C in a 2 liters culture flask containing 600 mL of TSB medium. Cells were collected by centrifugation at 6000 × g for 10 min and then transferred to a 1 liter culture flask containing 300 mL of TSB medium supplemented with 1.8 mM sodium selenite. Bacteria were allowed to grow for 5 h at 200 RPM at 37°C. Cells were collected by centrifugation at 12500 × g for 10 min and transferred to a 50 mL centrifuge tube. The pellet was washed and resuspended in 20 mL of sterile water. To this 120 μl of 100 mg/mL (12 mg) lysozyme solution was added, and the tube was kept at 37°C for 3 h. The culture was then crushed under high pressure (1500 Psi) in a French press. The resultant slurry containing both cell debris and NPs was washed four times at 15000 × g for 10 min with 1. 5 M Tris-HCl (pH 8.3) containing 1% SDS. The resultant pellet containing NPs and the insoluble cell wall fraction was washed and resuspended in 16 mL of sterile water. Aliquots of 4 mL were transferred to four high tensilestrength glass tubes (Schott GL18), and 2 mL of 1-octanol were added. The solution was mixed vigorously on a vortex mixture for few min and centrifuged at 2000 x g for 5 min at 4°C. The tubes were then kept undisturbed at 4°C for 24 h. The upper phase and interface containing the insoluble cell fraction were discarded, and the bottom water phase was transferred to a clean 50 mL centrifuge tube. This was washed sequentially with chloroform, absolute ethanol, 70 % ethanol, and water at 16000 × g. Collected NPs were resuspended in water and stored at 4°C. NPs could also be resuspended and stored in 50 mM Tris-HCl (pH 7.4). 150 μg of these extracted and purified SeNPs were spread on tryptic soya agar plates to observe the cell lysis efficiency of the process. SeNPs were characterized for shape, size, and purity by using various spectrophotometric [dynamic light scattering (DLS), energy dispersive X-ray spectroscopy (EDX), Fourier transformed infrared spectroscopy (FTIR)], microscopic [scanning electron microscopy (SEM), transmission electron microscopy (TEM)], and electrophoresis (SDS-PAGE) techniques. Samples for TEM were prepared by placing a drop of NP suspension on carbon-coated copper grids. The images were visualized at 200 kV on a JEOL JEM 2100 TEM microscope. SEM samples were prepared by a standard fixation procedure used for bacterial visualization [20]. Images of nanoparticles were obtained on a Zeiss EVO 40 SEM microscope. EDX analysis of air-dried NP samples was performed on a Bruker AXS X-Flash Detector 4010 Energy Dispersive X-Ray Microanalyser to determine the composition and purity of the particles. The size distribution and surface charge on the SeNPs were measured in a suspension on a Zetasizer Nano ZS particle analyzer (Malvern). FTIR (Thermo Scientific NICOLET iS10) studies in the range of 800 cm-1 to 3500 cm-1 were performed to determine the functional groups on the NPs. Extracted SeNPs were digested overnight with 3:1 nitric/ perchloric acid. Digested samples were analyzed in an atomic absorption spectrophotometer (AA-6800, Shimadzu) to determine the concentration of selenium present. An air-acetylene (oxidizing) flame and selenium cathode lamp were used and a wavelength of 196 nm was selected to measure absorption of incident light. The NP sample was denatured at 95°C for 10 min in gel loading buffer then electrophoresed on 15 % SDS-PAGE at 12 mA. Silver staining was performed to analyze the proteins associated with the NPs. NPs were tested for agglomerate formation at physiological temperature and pH. Extracted NPs were kept undisturbed at 37°C for 5 h in various concentrations of bicarbonate buffer, 0. 1 M, 0. 01 M, and 0. 001 M, pH 7.4. Accumulation of NPs at the bottom of microcentrifuge tube was used as a sign of agglomerate formation. A human prostate adenocarcinoma cell line (PC3) was obtained from the National Centre for Cell Science, Pune, India. The cells were grown in F12K media supplemented with penicillin (100 units/mL), streptomycin (50 units/mL), and fetal bovine serum (10 %), at 37°C in a humidified incubator with 5% CO2 . Human peripheral blood mononuclear cells (hPBMC) were isolated from healthy volunteers by Ficoll-Hypaque (Histopaque-1077) density gradient centrifugation. Cells were maintained in RPMI 1640 medium supplemented with 10 mM HEPES, penicillin (100 units/mL), streptomycin (50 units/mL), and fetal calf serum (10%), at 37°C in a humidified incubator with 5 % CO2 . Cell viability was determined by quantifying the ability of cells to convert water-soluble XTT sodium salt [2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide] to a water-soluble, orange-colored formazan product. PC3 cells were seeded with F12K medium in 96-well tissue culture plates at a density of 3. 5 X 103 cells per well. After 24 h of resting period at 37°C, cells were treated with SeNPs or SeMet at various concentrations, 1, 2, 4, and 6 μg Se/mL, and cultured for 24 h at 37°C. After the addition of 100 μl of freshly prepared XTT solution (0.5 mg/mL) supplemented with menadione (final concentration 1 μM) to each well, cells were incubated for 3 h at 37°C. Plates were then analyzed at 490 nm on a BioTek Power Wave Microplate reader. Production of an orange color in this assay corresponds to cell viability. Respective morphological responses to selenium treatment were visualized at 40 X under bright field in Olympus IX71 microscope. PC3 cells were seeded in 12-well plates at a density of 2×105 cells per well in F12K medium supplemented with 10% FBS and kept at rest for 24 h at 37°C. After the rest period, hPBMC isolated from the blood of healthy volunteers were seeded in RPMI medium supplemented with 10 mM HEPES and 10 % FCS in 12-well tissue culture plates at a density of 5×105 cells per well. Both the cell types were treated with SeNPs or SeMet at a concentration of 2 μg Se/mL, and incubated for 24 h. Cells were harvested, and an AnnexinV-FITC Apoptosis Detection Kit was used to stain cells with annexin V and propidium iodide according to the manufacturer's instructions. Cells were acquired in a BD AccuriC6 Flow Cytometer (BD Biosciences). The data analysis was performed with FlowJo software. Caspase-3/7 activity was determined by providing a luminogenic caspase-3/7 substrate, containing the tetra peptide sequence DEVD, to the NP treated cells. PC3 cells were seeded in 96-well opaque walled plate at a density of 8×103 cells per well in F12K medium supplemented with 10 % FBS and kept at rest for 24 h at 37°C. Fresh hPBMC isolated from the blood of healthy volunteers were seeded in RPMI medium supplemented with 10 mM HEPES and 10% FCS at a density of 20×103 cells per well. Both the cell types were treated with SeNPs at a concentration of 2 μg Se/mL, and incubated for 12 h at 37°C. ApoTox-Glo Triplex assay kit was used according to manufacturer's instructions. Glow-type luminescent signal proportional to the amount of caspase activity was measured by GloMax® -96 Microplate Luminometer. hPBMC were obtained and used strictly according to the ethical guidelines for biomedical research on human subjects by the Central Ethics Committee on Human Research, Indian Council of Medical Research-2000 as well as the standards in the Declaration of Helsinki. Written information about the study and the written consent were in the native language of the subject (English, Hindi, and Punjabi). The data represented here are the mean ± SEM from at least three independent experiments. Statistical significances were examined using two-tailed Student's t-test. SeNPs were synthesized by the intracellular conversion of toxic selenite ions (Se+4) into nontoxic elemental SeNPs (Se0 ) under aerobic conditions by the bacterium Bacillus licheniformis JS2 [22]. A method reported by Shakibaie and colleagues was applied for recovery of intracellular SeNPs. However, we found that the NPs obtained by this method were associated with the bacteria and formed agglomerates after recovery (Supplementary Figure 1). We improved the cell lysis procedure and recovered intracellular SeNPs by bacterial cell lysis using lysozyme and French press, cleaned by successive washes with Tris-HCl buffer and finally separated from insoluble debris by two-phase water-octanol extraction (Figure 1a). Spreading of purified and cleaned SeNPs on TSA plate showed no bacterial (Bacillus) growth indicating the cell lysis process is highly efficient (Figure 1b). DLS data indicate the size distribution of the SeNPs ranged from 40 to 180 nm (Figure 2a). No charge was observed on the surface of the SeNPs by a Zeta potential measurement (Figure 2b). The particles were found to be stable at physiological temperature and pH. When kept at 37°C for 5 h in various concentration of bicarbonate buffer, the particles with no charge on the surface did not form agglomerates, whereas the negatively charged particles (-29 mV) agglomerated and settled to the bottom of the tube (Figure 2c). Electron microscopy was performed to determine the exact structure and size of the SeNPs. SEM and TEM images (Figure 3a and 3b) showed that the particles are spherical in shape with an average diameter of 120 nm and did not forming aggregates. EDX spectra of the SeNPs indicated that the particles are composed of only selenium (Figure 3c). SDS-PAGE of SeNPs (50 μg Se) followed by silver staining suggests there are few bacterial proteins associated with the NPs. Protein bands of 70 KDa and ~50 KDa were visible although the quantity of protein was very low (Figure 4a). FTIR analysis showed that the particles have some functional groups attached to their surfaces. Most of the peaks represent protein or peptide groups: the peak at 3405 cm-1 corresponds to the N-H stretch; the peaks at 2917 and 2849 cm-1 represent the C-H stretch; the Page 4 of 9 peaks at 1633, 1462, and 1384 cm-1 were assigned to the amide bond; and the peak at 1050 cm-1 was assigned to the C-O stretch (Figure 4b). The effect of SeNPs on PC3 cell viability was studied by performing an XTT assay on cells cultured in the presence of various concentrations of the SeNPs. We found that the SeNPs reduced the viability of the prostate cancer cell line PC3 at all the concentrations tested, 1, 2, 4, and 6 μg Se/mL. Only 45 % cell viability was observed at 6 μg Se/mL. Because SeMet is the major dietary supplement form of selenium, it was used as a control and caused no significant decrease in cell viability (Figure 5a). In support to XTT results, bright field micrograph of selenium treated PC3 cells confirms the cellular damage on SeNP treatment. Most of the cells lost their typical morphology, became shrunken. Detachment of cells from the plate surface has been observed after SeNP treatment. Decrease in the adherent cell number was observed with the increase in SeNP concentration, whereas, no change was observed in SeMet treated cells (Figure 5b). To determine if the reduction in cell viability of PC3 cells treated with SeNPs was caused by induction of apoptosis or necrosis, we performed AnnexinV-FITC and propidium iodide (PI) staining after treating the PC3 cells for 24 h with SeNPs or SeMet at a concentration of 2 μg Se/mL. hPBMC were used as a non-cancerous control. Fluorescence-activated cell sorting (FACS) analysis suggested that at a very low concentration, 2 μg Se/mL, the SeNPs induced late apoptosis and/or necrosis in PC3 cells (Figure 6a). At this concentration of SeNP no significant apoptosis or necrosis was seen in hPBMCs (Figure 6b). To find the mechanism of cell death ApoTox-Glo triplex assay was performed on NP treated PC3 cells and hPBMC to measure the caspase-3/7 activity. In support to the FACS result, decreased luminescence signals was observed in PC3 cells treated with 2 μg Se/mL SeNP for 12 h, suggests low caspase activity in these cells in comparison to untreated cells. While no change in the luminescence was observed in hPBMC. Results implicate that SeNPs are able to induce cytotoxicity in cancerous cells through caspase independent necrotic pathway with no cytotoxicity to non-cancerous hPBMC. Commercial physical and chemical processes for NP synthesis are not ecofriendly and thus create a burden on our environment. Therefore, there is a growing need to replace these processes with a green synthesis method. In this study, we used the bacterial strain Bacillus licheniformis JS2 for the biological synthesis of SeNPs. In this bacterium the SeNPs are synthesized intracellularly. We developed an improved method for the recovery of intracellular NPs and compared the toxicity of these biogenic SeNPs on human prostrate adenocarcinoma cell line PC3 and hPBMC. Biologically synthesized SeNPs were toxic to PC3 cells by inducing necrosis; however, they appeared to be without significant toxicity to hPBMC. Extraction of intracellular NPs is challenging because of the difficulty in separating the particles from the cell components. Consequently, no efficient method for this extraction has been reported yet. Shakibaie et al. [21] have reported a method for intracellular NP recovery that uses liquid nitrogen and sonication for bacterial cell lysis. However, we found this method was not effective in lysing all bacterial cells and resulted in association of microbes, both dead and alive, with the extracted NPs. In addition, the extracted particles formed agglomerates. We performed cell lysis using lysozyme and a French press at 1500 psi. This method is highly effective for obtaining complete lysis. After removal of all soluble cytosolic fractions by Tris-HCl washes, a twophase extraction system of water-octanol was used for NP recovery. Water-octanol has already been reported as a good partitioning system for the removal of cell wall debris, denatured proteins, and most polysaccharides found in living systems [23]. This procedure resulted in very pure SeNPs with no associated bacteria and only a minute quantity of protein, as confirmed by SEM, EDX, TEM, SDS-PAGE, and FTIR analyses. This appears to be the most successful method for the recovery of intracellular NPs reported thus far. DLS, SEM, and TEM analyses indicate these NPs do not form agglomerates. We observed an interesting result from Zeta potential analysis that showed a neutral charge on the NP surface. Even while Page 6 of 9 carrying the neutral charge, these particles were stable, not forming agglomerates at physiological temperature, pH and buffer condition for 5 h, compared to negatively charged particles. As these NPs were more stable at physiological conditions, we hypothesized that the SeNPs may be sterically stabilized. In support to our hypothesis, SDS- PAGE and silver staining results showed that the particles were associated with some protein, although the quantity of protein was very low (Figure 4a). In addition, FTIR analysis showed that the SeNPs have some functional groups attached to the surface (Figure 4b). These findings indicate that our NPs have a polymer and/or protein coating on their surface which provides steric stability to them, as reported previously [24]. Selenium has been shown to be an anticancer agent especially for prostate, colon, and lung cancers. However, the anticancer effect is seen at high doses that adversely affect cell homeostasis and cause severe toxicity [2,3]. As the difference between the effective and toxic concentrations of selenium is very marginal, it is vital to maintain the correct amount of selenium to treat the cancer but not cause toxicity. Reports on nano-sized selenium suggest it has similar anticancer activity with excellent bioavailability and lower toxicity compared to other organic and inorganic selenium compounds [3,4,7,25]. In this study we show that a concentration of 2 μg Se/mL of biogenic SeNPs is very effective in inducing necrosis in PC3 cells via caspase-3/7 independent necrotic pathway. This is significantly different from other reports that suggest that SeNPs are able to induce cytotoxicity to the same extent at 100 μg/mL concentration [21]. The SeNPs in those reports had a surface charge and formed aggregates, which may have led to less availability and efficacy resulting in the requirement for a high concentration to induce apoptosis or necrosis in cancerous cells. The biogenic SeNPs in this report are sterically stabilized and do not form aggregates at physiological conditions, which appears to lead to improved effectiveness. These results suggest that sterically stabilized SeNPs could be a potential candidate for cancer chemotherapy and chemoprevention. Further studies on the pathway behind the induction of necrosis are needed to investigate this potential. Use of lysozyme and a French press for bacterial cell lysis followed by an organic-aqueous extraction system is a more successful method for recovery of intracellular NPs then previously used techniques. By using this extraction procedure we obtained pure and clean, sterically Page 7 of 9 μ Figure 6: Biogenic SeNPs induced necrosis in PC3 cells but failed to induce apoptosis or necrosis in hPBMC. (a) PC3 cells (b) and hPBMC were cultured in the presence of SeNPs or SeMet at a concentration of 2 μg Se/mL for 24 h, while the control cells were not treated. The percent of apoptosis/necrosis was measured by Annexin-V and PI staining. Data presented are representative of flowcytometric experiments conducted in triplicate. PC3 cells (c) and hPBMC (d) were cultured in the presence of SeNPs at a concentration of 2 μg Se/mL for 12 h. Caspase-3/7 activity was measured by using ApoTox-Glo assay kit containing a luminogenic caspase-3/7 substrate. Cleavage of this substrate by caspases produced a \"glow- type\" luminescent signal. The amount of luminescence is proportional to the activity of caspases in the cells. Experiment was conducted in triplicate. p<0. 01 represents significant difference in the level of caspase-3/7 activity between SeNP treated and untreated PC3 cells PC3 comp-FL2-A:: Pl-A hPBMC comp-FL2-A:: Pl-A 250000 200000 150000 100000 Luminescence 50000 0 μ Page 8 of 9 stabilized SeNPs from Bacillus licheniformis JS2. Compared to the commercially available selenium supplement SeMet, these NPs are very effective in inducing necrosis via caspase independent necrotic pathway in human prostate adenocarcinoma cells without affecting the viability of hPBMC. To the best of our knowledge this is the first report of uncharged, sterically stabilized biogenic SeNPs that do not form aggregate at physiological conditions and are highly efficacious and specific against prostate cancer at concentration as low as 2 μg Se/mL. We are grateful to the Director, Institute of Microbial Technology - a constituent laboratory of Council of Scientific and Industrial Research, for providing excellent infrastructure and facility. This study was financially supported by Council of Scientific and Industrial Research (CSIR project head OLP0064); CSIR (fellowship to PS); and the University Grants Commission (fellowship to R.N.). 10. Bhattacharya D, Gupta RK (2005) Nanotechnology and potential of microorganisms. Crit Rev Biotechnol 25: 199-204. Page 9 of 9 </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 91.09037}, {\"h\": 24.0, \"mgL\": null, \"viab\": 77.62113}, {\"h\": 24.0, \"mgL\": null, \"viab\": 54.2135}, {\"h\": 24.0, \"mgL\": null, \"viab\": 44.062363}]}", "role": "assistant" } ]	128.json
[ { "content": "Extract sample context and measurement list. <doc> Fullerenes, Nanotubes and Carbon Nanostructures ISSN: 1536-383X (Print) 1536-4046 (Online) Journal homepage: https://www.tandfonline.com/loi/lfnn20 Svitlana V. Prylutska, Anna G. Grebinyk, Oksana V. Lynchak, Iryna V. Byelinska, Vsevolod V. Cherepanov, Eric Tauscher, Olga P. Matyshevska, Yuriy I. Prylutskyy, Volodymyr K. Rybalchenko, Uwe Ritter & Marcus Frohme To cite this article: Svitlana V. Prylutska, Anna G. Grebinyk, Oksana V. Lynchak, Iryna V. Byelinska, Vsevolod V. Cherepanov, Eric Tauscher, Olga P. Matyshevska, Yuriy I. Prylutskyy, Volodymyr K. Rybalchenko, Uwe Ritter & Marcus Frohme (2019): Invitro and invivo toxicity of pristine C60 fullerene aqueous colloid solution, Fullerenes, Nanotubes and Carbon Nanostructures, DOI: To link to this article: https://doi.org/ Accepted author version posted online: 18 Jun 2019. Published online: 29 Jun 2019. Submit your article to this journal Article views: 5 View Crossmark data Svitlana V. Prylutskaa , Anna G. Grebinykb,c , Oksana V. Lynchakb , Iryna V. Byelinskab , Vsevolod V. Cherepanovd , Eric Tauschere , Olga P. Matyshevskaf , Yuriy I. Prylutskyyb , Volodymyr K. Rybalchenkog , Uwe Rittere , and Marcus Frohmec a Faculty of Chemistry, Taras Shevchenko National University of Kyiv, Kyiv, Ukraine; b Educational and Scientific Center \"Institute of Biology and Medicine\", Taras Shevchenko National University of Kyiv, Kyiv, Ukraine; c Division Molecular Biotechnology and Functional Genomics, Technical University of Applied Sciences Wildau, Wildau, Germany; d Department of Physical Electronics, Institute of Physics of the National Academy of Sciences of Ukraine, Kyiv, Ukraine; e Institute of Chemistry and Biotechnology, Technical University Ilmenau, Ilmenau, Germany; f Department of Scientific and Technical Information, Palladin Institute of Biochemistry of the National Academy of Sciences of Ukraine, Kyiv, Ukraine; g Institute of High Technologies, Taras Shevchenko National University of Kyiv, Kyiv, Ukraine Toxicity of the C60 fullerene aqueous colloid solution in vitro and in vivo experiments was studied. C60 fullerene in the concentration range of 3.6–144 mg/mL demonstrated low toxicity against human embryonic kidney (HEK293) cells with 383.4 lg/mL IC50 value. The acute toxicity of C60 fullerene in a dose progression (75, 150, 300, 600, 1200 and 1800 mg/kg) was investigated at 14th day after single intraperitoneal (i.p.) administration into mice. No toxic effect of 60 fullerene in the dose range of 75–150 mg/kg was observed; the calculated LD50 value was 721 mg/kg. The toxic effect of C60 fullerene was observed at a concentration of 300 mg/kg and higher, it was accompanied with mice behavior disturbance, hematotoxicity and pathomorphological changes in spleen, hepatic and kidney tissues. In mice treated with 600 mg/kg of C60 fullerene indices of inflammatory process were detected, whereas at 1200 mg/kg dose leukopoies and thrombocytopoies were found to be suppressed. Thus, C60 fullerene aqueous colloid solution in a dose range of 75–150 mg/kg is nontoxic and could be used for biomedical purposes. Received 12 June 2019 Accepted 16 June 2019 C60 fullerene; DLS and AFM analysis; fluorescent microscopy; IC50 and LD50 values; hematological and histological analysis Rapid development of nanotechnologies is accompanied by using of nanocarbon materials in various industries. C60 fullerene and its derivatives are used in microelectronics and optics for solar and electrical batteries,[1] as well as in biology and medicine for targeted drug delivery, theranostics etc.[2–9] fullerene from the respiratory routes occurs through socalled mucociliary escalator of ciliated epithelial cells and alveolar macrophages phagocytosis.[48] C60 fullerene dispersed in squalane easily penetrates into the human epidermis through stratum corneum, but does not pass through the basement membrane and does not get into the dermal layer.[49] After peroral (intragastrical) administration C60 fullerene was detected mainly in the stomach and small intestines, liver, kidneys and spleen of animals and was excreted with feces.[50] Although the pristine C60 fullerene and its derivatives are widely used in various biomedical research,[56–58] C60 fullerene toxic effects remain poorly investigated. Therefore, our goal was to evaluate the toxicity of the pristine C60 fullerene aqueous colloid solution (C60FAS) both in vitro and in vivo experiments. For this purpose, we determined IC50 value of C60 fullerene in human embryonic kidney (HEK293) cells and LD50 value of C60 fullerene after i.p. injection into mice, as well as hematological parameters and histological changes in murine tissues. Phenyl-C61-butyric acid chloride, 4-Hydroxythiazol, 4-Dimethylaminopyridine, Toluene (VWR corporation, USA). Dulbecco's modified Eagle's medium (DMEM), Fetal calf serum (FCS), Penicillin, Streptomycin, L-gluthamine, Dimethylsulfoxide (DMSO), MTT [3-(4,5-dimethylthiazol-2 yl)-2,5-diphenyl tetrazolium bromide], NaCl, Hoechst33342 (Sigma-Aldrich Co, Ltd, USA). Kit for hemoglobin evaluation (Filicit, Ukraine), Acetic acid, Ammonium oxalate (Chemlaborreactiv, Ukraine), May-Grunwald and Romanowsky stain (Macrochem, Ukraine). Formalin, chloroform, ethanol (Inter Synthesis, Ukaine), Hematoxylin, Eosin (Leica Biosystems, USA), Paraplast plus (McCormick Scientific, USA). A highly stable C60FAS (initial concentration 2.6 mg/ml, purity >99.5%) was synthesized in Technical University of The state of C60 fullerene particles in aqueous solution was monitored using atomic force microscopy (AFM). AFM measurements were performed on the \"Solver Pro M\" system (NT-MDT, Russia) in a semicontact (tapping) mode using RTESPA-300 (Bruker) probes. The selection of sites for measurements was carried out by use of an integrated optical microscope. Layers deposited from aqueous solution droplets onto a freshly cleaved atomic smooth surface of mica (V1 grade, SPI Supplies) were investigated. All measurements were performed after complete evaporation of the water. Solvents were dried according to standard procedures. All reactions were carried out by the use of Schlenk-technique under an atmosphere of argon or nitrogen. 1 and 3 samples (Figure 1) were prepared according to protocols.[62–64] Nuclear magnetic resonance spectra (1 H, 13C, DEPT 135) were performed on a Bruker Spectrospin 300 MHz instrument. MALDI spectrometry was performed on a Bruker microflex LT. The high-performance liquid chromatography (HPCL) was performed on a JASCO system using a COSMOSIL \"bucky prep\" waters column, solvent and retention time are given in the procedure. HEK293 cells were kindly supplied by the Bank of Cell Cultures and Transplantable Experimental Tumors of R.E. Kavetsky Institute of Experimental Pathology, Oncology and Radiobiology of the National Academy of Sciences of Ukraine (Kyiv, Ukraine). Cells were maintained in DMEM supplemented with 10% FCS, 50 U/ml penicillin and 100 lg/ ml streptomycin at 37 C in a humidified atmosphere with 5% CO2. Cells were incubated at different concentrations with or without C60 fullerene. Curve fitting and calculation of the half-maximal inhibitory concentration (IC50 value) were done using GraphPad Prism 7 (GraphPad Software Inc., USA). Briefly, individual concentration-effect curves were generated by fitting the logarithm of the tested compound concentration versus corresponding normalized percent of cell viability values using nonlinear regression. The study was conducted on 42 adult male mice of inbreeding line 2–2.5 months old with the weight 21 ± 2 g. Animals were kept under standard conditions in the vivarium of the ESC \"Institute of Biology and Medicine\", Taras Shevchenko National University of Kyiv. Mice had been bred under ordinary conditions for research aims. Animals were housed in plastic cages with stainless steel mesh lids in a ventilated room, with maintained temperature (23 ± 2 C) on a reversed 12 h light/dark cycle, with free access to food and water. All experiments in this study were performed according to the Bio-Ethics Committee of the abovementioned institution. The experiment was simulated in accordance with the requirements for determining the acute toxicity of the compound.[66,67] Animals were randomly divided into seven groups. C60FAS in a C60 fullerene graded doses (75, 150, 300, 600, 1200 and 1800 mg/kg) and the same volume for obtaining the desired dose were i.p. injected into animals. Animals in the control group received 0.9% NaCl solution (saline) i.p. at same volume. Animals dying during the period of experiment and those that survived up to the end of the 14 days period after euthanasia with CO2 were inspected for organ weights, hematological and histopathological findings. Animals were continuously observed in their cages for 1 h, intermittently for 3, 6 and 12 h and then daily for signs of toxicity. Body weight was recorded on 0, 7th and 14th days. The blood for cytological analysis was collected from the femoral vein into the EDTA-treated tube. Quantitative blood indices such as RBC count, hemoglobin concentration, hematocrit, mean corpuscular volume, values of mean corpuscular hemoglobin, mean corpuscular hemoglobin concentration, white blood cell and platelet counts were determined using common methods.[68] The differential analysis of leukograms was conducted using blood smears, stained according to Pappenheim, counting 100 white blood cells, including basophil, eosinophil and neutrophil granulocytes, lymphocytes and monocytes. After blood collection, all mice were dissected and observed macroscopically. The heart, liver, spleen and kidneys were removed and weigted. The stomach, small and large intestine as well as the above organs were fixed in 10% neutralbuffered formal saline, and dehydrated in different ethanol concentrations and paraffin blocks were prepared. Paraffin sections were stained with hematoxylin-eosin (HE) for microscopic examination. The number of mice died within the period of the study was registered in each group and the LD50 value was calculated by probit analysis.[69] Values of IC50 and LD50, body and organ weight were represented as M ± SD of more than four independent experiments. Statistical analysis was performed using two-way ANOVA followed by post Bonferroni tests. A value of p < 0.05 was considered statistically significant. Data processing and plotting were performed by IBM PC using specialized applications GraphPad Prism 7 (GraphPad Software Inc., USA). The statistical analysis of the hematology data was performed using SPSS 16.0 software package for Windows. The Gaussian distribution of the data was estimated using Shapiro-Wilk test. Kruskal-Wallis criterion for nonstandard distribution (the difference was considered statistically significant at p < 0.05) was used for multiple comparisons with further application of Mann-Whitney test (the difference was considered significant at p < 0.05) for non-parametric data. The median, 25 and 75 percentiles were calculated.[70] The DLS measurement testified that the main fraction of light scattering C60 fullerene particles in aqueous solution at a concentration 2.6 mg/ml has the hydrodynamic diameter up to 125 nm. C60 fullerene particles demonstrated a high negative average surface charge (zeta potential f 1⁄4 -25.3 mV), indicating a very low tendency for aggregation in aqueous solution (i.e., a high solute stabilization). The polydispersity index (PDI) of C60FAS was shown to be 0.25 indicating on a moderately dispersed distribution of particles. The aqueous colloid solution of pristine C60 fullerene was stable for 9 months at room temperature indicating on possibility of its usage in biological studies. well as on the route, dose and type of administration to the organism and the number of exposures (single or multiple).[6,82–84] The cytotoxic activity of pristine 60 fullerene in a concentration range of 3.6–144 mg/mL against HEK293 cells at 24 h of incubation was investigated using MTT assay. The viability of HEK293 cells incubated without C60 fullerene was taken as 100% (control). The absence of C60 fullerene toxic effect against HEK293 cells is in accordance with the data about the lack of C60 fullerene toxicity against normal cells in a low concentration range: no toxic effects of pristine C60 fullerene in concentration of 1.5 mg/mL against human keratinocytes,[87] 1-10 lg/mL against nondifferentiated rat phencocytoma PC-12 neuronallike cells,[14] 10–200 lg/mL against normal human mammary epithelial MCF10A cells,[88] as well as of 50–300 lg/mL against baby hamster kidney BHK-21 cells[89] were observed. The toxicity of pristine C60 fullerene at high 320 mg/mL concentration against BHK-21 cells[89] was demonstrated, that is in a good agreement with our data. As the intensity of C60 fullerene fluorescence is very low,[90] the monitoring of its intracellular accumulation with fluorescent microscopy is complicated and requires additional labeling of this nanostructure with the fluorescent dye (sample 4; Figure 1). No unspecific green fluorescence was observed in the control untreated cells (Figure 4A). When HEK293 cells were incubated with labeled C60 fullerene (14.4 mg/mL) for 6 h, the green fluorescent signal was detected. The merged images of phase contrast and fluorescent microscopy confirmed accumulation of C60 fullerene inside HEK293 cells. HEK293 cells were also analyzed with flow cytometry to asses the value of the fluorescence intensity signal (Figure 4B). Autofluorescence of the untreated cells was used as a negative control. On the FACS histagramm the fluorescence distribution of the HEK293 cells treated with labeled C60 fullerene was shifted to the right detecting relatively slight accumulation of the studied nanoparticles in the normal cells. The i.p. injection has several advantages over other methods of drug administration, namely higher doses and accumulation in the organs of the peritoneal cavity as compared with intravenous administration; drugs are not exposed to enzymes and juices of of the gastrointestinal tract organs as compared with enteral administration. C60 fullerene administration in 75 and 150 mg/kg doses did not cause mice death during the experiment, while one (16.7%) and two animals (33.3%) died at 300 and 600 mg/kg doses of C60 fullerene, respectively (Table 1). After i.p. injection of 1200 mg/kg of C60 fullerene only two animals were alive at the 14th day of the experiment, while after injection of 1800 mg/kg all mice died (Table 1). The death of the first mice was registered at 30 min after i.p. administration of 60 fullerene at a dose of 1800 mg/kg. At 2 h after i.p. administration of 60 fullerene at a dose of 1200 mg/kg, the death of one mice was registred. The last death of animals in different groups was recorded up to 7th day of the experiment. Next task of this study was to evaluate the physiological state of mice after C60FAS i.p. administration. No statistically significant difference in water and dietary consumption between control groups of animals and those treated with C60 fullerenes was observed. As well as no noticeable changes in the body (Table 2) and organs (data are not shown) weights of mice after i.p. administration of C60 fullerene in different doses compared with the control group were detected. The blood has direct contact with tissues and assures their homeostasis, therefore hematological parameters are the universal clinical and diagnostic indicators of the organism physiological state both in normal condition and after C60 fullerene i.p. administration. The state of mice erythrocytes on 14th day after i.p. administration of C60 fullerene in increasing doses of 75, 150, 300, 600, 1200 mg/kg was evaluated with the use of such indicators as erythrocytes count, blood hemoglobin concentration (Hgb), hematocrit (HCT), mean corpuscular volume (MCV), mean corpuscular hemoglobin (MCH) and mean corpuscular hemoglobin concentration (MCHC) (Figure 5). Thus, C60 fullerene in high doses affected the state of RBC, as evidenced by the increase in Hgb, HCT, MCV and MCH levels and reticulocytes count in blood of mice. The increase of reticulocytes number, which is followed by the erythrocytes average volume (MCV) enlargement and hematocrit growth, indicated on erythropoiesis activation in hemopoietic organs in response to C60 fullerene high doses, while the increase of hemoglobin amount per erythrocyte (MCH) testified that no inhibition of hemoglobin synthesis by C60 fullerene occurred. No effects of 60 fullerene at doses 75–300 mg/kg at 14th day after single i.p. administration on the total count and composition of leukocytes were observed. When C60FAS dose was raised to 600 mg/kg, the increase of leukocytes count in blood by 82%, lymphocytes by 95%, monocytes by 94%, neutrophilic granulocytes by 62% and bands by 83% was detected (Table 3), that indicated on inflammation development. The neutrophilia and change in the morphology of neutrophils, elevated contents of band and hypersegmented neutrophils are the features of inflammatory processes, as well as of some of neoplastic changes in the hematopoietic system.[96] 60 fullerene in the highest 1200 mg/kg dose was shown to suppress leukopoiesis and thrombocytopoiesis by decreasing the total white blood cells and platelets counts by 63% and 34%, respectively, in comparison with control (Figure 6B, Table 3). detected at high doses (more than 300 mg/kg) and is accompanied by changes in the number and morphology of blood cells. The ability of C60 molecules to accumulate in the body[53] leads to its prolonged action, which is accompanied by more significant changes in the state of erythrocytes at high doses of this carbon nanostructure. Macroscopically no substantial abnormality was observed in organs of mice that received low or moderate doses of C60 fullerene. No changes in the size, shape and color of the organs or tissues hyperplasia were detected. C60 fullerene, i.p. injected to mice in the doses of 1200 and 1800 mg/kg, p < 0.05 compared to control group; ND – not defined. accumulated as brown pigmented inclusions in the mesentery, liver and spleen (data are partially presented on Figure 8). Autopsies revealed the dose dependent changes in the liver and spleen starting from the 150 mg/kg dose, and in kidneys at above 300 mg/kg dose. 60 fullerene at 14th day after single i.p. administration in doses 300 mg/kg and higher caused the fibrotic and distrophic changes in murine kidneys, liver and spleen tissues. At the highest doses (1200 and 1800 mg/kg) C60 fullerenes accumulated in the kidneys and liver of animals and caused the nephritis, fibros and distrophia of hepatocytes (Figure 8). The histopathological changes in the kidneys, liver and spleen testify the development of the inflammatory processes in mice treated with 1200 mg/kg of C60FAS (Figure 8) as confirmed by infiltration of white blood cells in organs. No abnormality was defined in other organs (heart, stomach, small and large intestine) of animals injected with C60 fullerenes (data are not presented). The pathomorphological changes in the spleen (the increase of red pulp area as well as the decrease of white pulp area) in mice treated with C60 fullerene (600 mg/kg and higher doses) correlated with hematological effects. Reticulocytosis in blood, increased red pulp area in the spleen and strong eosinophilia are assumed to be the result of RBC accumulation after the action of C60FAS in high doses. The data obtained demonstrated a low C60FAS toxicity in vitro and in vivo experiments, confirmed by calculated IC50 and LD50 values as well as by hematological and histological analyzes. The lethal C60 fullerene doses were shown to impact the behavioral reactions, hematological and histopathological indices, but to have no effect on the food and water consumptions, body and organs weight. The toxicity of the pristine C60 fullerene aqueous colloid solution (C60FAS) in vitro and in vivo experiments was estimated. Using the dynamic light scattering and atomic force microscopy techniques it was shown that C60FAS (2.6 mg/ ml) is a highly stable polydisperse system containing mainly single C60 molecules, their nano- and microaggregates up to 125 nm. The labeled C60 fullerene was taken up by the human embryonic kidney (HEK293) cells and localized in cytoplasm. C60 fullerenes demonstrated a low toxicity against HEK293 cells at 24 h of incubation, IC50 value was shown to be 383.4 lg/mL. Acute toxicity was determined after intraperitoneal (i.p.) injection of C60 fullerenes to mice, LD50 value was 721 mg/kg. No noticeable changes in the body and organs weights of mice were observed after C60FAS i.p. administration at investigated doses of 75, 150, 300, 600, 1200 and 1800 mg/kg. At 1200 and 1800 mg/kg doses C60 fullerene caused the decrease of mice activity, rapid respiration, twitches and tremor. No toxic effect of 60 fullerene in the range of 75–150 mg/kg was observed. After administration of 300 mg/kg of C60 fullerene hematologic indices were disturbed. At 600 and 1200 mg/kg doses of C60 fullerene the increased hemoglobin and hematocrit level, red blood cells and reticulocyte number as well as the erythropoiesis activation in the hematopoietic organs of mice were observed. At 600 mg/kg dose of C60 fullerene the development of inflammatory process in animals was detected and confirmed by the increased leukocytes, neutrophil granulocytes, lymphocytes and monocytes total amount. At 1200 mg/kg dose C60 fullerene suppressed the leuko- and thrombocytopoies, as confirmed by the decreased number of the nucleated cells and megakaryocytes in the bone marrow as well as leukocytes and platelets in the blood as compared with control. The pathomorphological changes in the spleen (increased red pulp area and decreased white pulp area) of mice treated with 600 mg/kg and higher dose of C60 fullerene correlated with hematological effects. Thus, the results obtained allowed to determine the nontoxic or low toxic C60FAS concentrations, recommended for application in modern nanobiotechnologies. This study was partially supported by the STCU project N 6256. SVP and AGG thank the DAAD for support. Svitlana V. Prylutska http://orcid.org/ Anna G. Grebinyk http://orcid.org/ Oksana V. Lynchak http://orcid.org/ Iryna V. Byelinska http://orcid.org/ Vsevolod V. Cherepanov http://orcid.org/ Eric Tauscher http://orcid.org/ Olga P. Matyshevska http://orcid.org/ Yuriy I. Prylutskyy http://orcid.org/ Volodymyr K. Rybalchenko http://orcid.org/ Uwe Ritter https://orcid.org/ Marcus Frohme https://orcid.org/ Carbon Nanotubes in Thymus Cells. Mat-Wiss. u Werkstofftech. 2009, 40, 238–241. DOI: . possible Involvement of the Nrf/ARE-Antioxidant Pathway. Oxid. Med. Cell. Longev. 2018, 2018, 2518676. DOI: 10.1155/ 2018/2518676. Accumulation in Human Leukemic Cells and Perspectives of LED-Mediated Photodynamic Therapy. Free Radic. Biol. Med. 2018, 124, 319–327. DOI: . of Water-Soluble Polyalkylsulfonated C60 in Rats. Toxicol. Pathol. 1998, 26, 143–151. DOI: . [56] Moussa, F. [60]Fullerene and derivatives for biomedical applications. Nanobiomaterials. 2018, 113–136. Intestinal Smooth Muscles. Cell Signal 2018, 43, 40–46. DOI: . Fullerenes in Vitro. Fullerenes. Nanotub. Carbon Nanostruct. 2008, 6, 698–705. DOI: . </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 3.6, \"viab\": 100.0}, {\"h\": 24.0, \"mgL\": 7.2, \"viab\": 96.117}, {\"h\": 24.0, \"mgL\": 18.0, \"viab\": 90.049}, {\"h\": 24.0, \"mgL\": 36.0, \"viab\": 79.854}, {\"h\": 24.0, \"mgL\": 72.0, \"viab\": 77.913}, {\"h\": 24.0, \"mgL\": 144.0, \"viab\": 73.058}]}", "role": "assistant" } ]	103.json
[ { "content": "Extract sample context and measurement list. <doc> dental mat e r ial s x x x ( 2 0 1 5 ) xxx–xxx Available online at www.sciencedirect.com j ou rnal homepa ge: www.intl.elsevierhealth.com/journals/dema Department of Biomaterials Science, Osaka University, Graduate School of Dentistry, 1-8 Yamadaoka, Suita, Osaka 565-0871, Japan Article history: Received 14 April 2014 Received in revised form 27 August 2014 Accepted 10 February 2015 Available online xxx Keywords: Oxide-nanoparticle Aluminum Silica Macrophage Cytotoxicity Genotoxicity Comet assay SEM TEM Confocal microscopy Objective. Although aluminum oxide and silicon oxide nanoparticles are currently available as dental materials, there is a lack of basic information concerning their biocompatibility. This study evaluates the biological responses of cultured macrophages (RAW264) to aluminum oxide (Al2O3NPs) and silicon oxide nanoparticles (SiO2NPs) by analyzing cytotoxicity and genotoxicity. Methods. The nanoparticles are amorphous and spherical, with diameters of 13nm for the Al2O3NPs and 12nm for the SiO2NPs. The cultured RAW264 are exposed to the nanoparticles (NPs) and examined for cytotoxicity using the WST-8 cell viability and Hoechst/PI apoptosis assay, for genotoxicity by micronucleus analysis, for changes in nuclear shape (deformed nuclei) and for comet assay using confocal microscopy, and micromorphological analysis is done using scanning and transmission electron microscopes. Results. Nuclei and DNA damage because of exposure to both types of NPs is observed by inmunostaining genotoxicity testing. The cytotoxicity and genotoxicity are well correlated in this study. Numerous NPs are observed as large aggregates in vesicles, butless or nonexistent NP internalization is seen in the nucleus or cytoplasm. These morphological results suggest that a primary cause of cell disruption is the chemical changes of the NPs in the low pH of vesicles (i.e., ionization of Al2O3 or SiO2) for both types of oxide NPs. Significance. Although further research on the elution of NP concentrations on cell or tissue activity under simulated clinical conditions is required, NP concentrations over 200g/mL are large enough to induce cytotoxic and genotoxic effects to cells. 2015 Academy of Dental Materials. Published by Elsevier Ltd. All rights reserved. Nanoscale particles are one of the fastest growing sectors in materials research and are currently being used in a wide range of fields, such as industrial, military, and biomedical applications, owing to their unique size-dependent properties. The main characteristic properties of the nanoparticle, namely quantum size effects and surface-induced effects, result from its small size. A large number of nanoparticle products are currently in development and under investigation, and some are already commercially available. For ∗ Corresponding author. Tel.: +81 6 6879 2917. dental applications, numerous new dental products containing nanoparticles have been developed and are available for implants, pulp capping agents, restorative materials, toothpaste, and oral hygiene products [1]. Titanium dioxide, silicon dioxide, aluminum oxide, and zinc oxide are commonly available for a variety of applications in dental materials. Nano-sized silicon dioxide is one of the most popular nanomaterials for restorative materials such as resin composites or bonding resins [2–5]. In addition, nano-sized aluminum oxide is used in orthopedic materials [6]. Recently, these two materials have been used as pulp capping agents and as a surface coating for dental implants that come in direct contact with pulp tissues and periodontium [3,6]. Although dental applications require nanoparticles (NPs) with a high degree of biocompatibility to protect human health, there is a serious lack of basic information concerning NP toxicity, and especially genotoxicity. A rapid approach for risk assessment is needed because of the rapid development of dental products using various particles with different chemical and physiological properties. Toxicological studies of the interaction of NPs with biological systems, including living cells and organs, have spurred a growing awareness of the need to predict the possible hazard of NPs via collaborative research in science and industry. Nanoparticle sizes are similar to that of cellular components, such as nuclei, mitochondria, and lysosomes, and they may be able to bypass cellular barriers, thereby leading to unexpected adverse tissue reactions. Therefore, an important topic in NP research is basic morphological information on the location and internalization of NPs; namely, where and how NPs are incorporated into cellular components. Because aluminum oxide and silicon oxide NPs are commonly used in dental materials, the purpose of this study is to evaluate the effects of aluminum oxide NPs (13nm diameter) and silicon oxide NPs (12nm diameter) upon cultured RAW264 using cytotoxicity (WST-8 assay), immunostaining genotoxicity (deformed nuclei, micronuclei, Hoechst/PI apoptosis, and comet assay) and micromorphological analysis of NP internalization using scanning and transmission electron microscopes. The null hypothesis in this work is that there exist no cytotoxic and genotoxic effects from aluminum oxide and silicon oxide NPs upon RAW264 cells. Two types of Al2O3 NPs (Al2O3NPs) and SiO2 NPs (SiO2NPs) (Sigma–Aldrich, St. Louis, MO, USA) were used in this study. The transmission electron microscope (TEM) samples were prepared by evaporating a droplet of the aqueous dispersion of NPs onto a carbon-coated copper grid. The crystallinity and surface charge of the NPs were characterized by an Xray diffractometer (XRD; Rint 2100; Rigaku, Tokyo, Japan) and a zeta potential analyzer (ZetaPALS, Brookhaven Instruments, NY, USA), respectively. The RAW264 murine macrophage cell line (Riken Biosource Center Cell Bank, Tsukuba, Japan) was cultured at 37 ◦C in an -minimal essential medium (Wako Chemical, Tokyo, Japan) containing 10% fetal bovine serum (Biowest, Nuaillé, France) and 1% antibiotic–antimycotic (Anti-Anti, Gibco, NY, USA) in a humidified atmosphere of 5% CO2 in air. Cells were grown in a cell culture dish and checked using a microscopic control that was free of any contamination. Confluent cells were detached using a cell scraper and aliquots of separated cells were subcultured at 1:4 splits every 3 days. Cells were seeded in Falcon 96-well plates (Corning, NY, USA) at a density of 1 × 104 cells per well with 200-L of medium. After 72h of culturing, the medium was removed and 200-L of medium containing two different NP concentrations (200 and 400g/mL) was added to the 72h cultured cells. Cells that had not been treated using the NPs were used as a control. The final particle dispersions were prepared fresh before use by serial dilution of the stock suspension (400g/mL) in medium. The stock solution was mixed using a vortex, and then sonicated for 5min to reduce agglomeration of the nanoparticles. The NP-exposed cells were subsequently incubated for 24h. The cell viability was measured using a WST-8 assay (CCK-8, Dojindo, Kumamoto, Japan). The cells were incubated in medium in the 96-well plates (1 × 104 cells/200-L medium/well) for 72h, and subsequently for 24h after being exposed to the NPs. The medium was then removed via aspiration and the cells were washed twice using phosphatebuffered saline (PBS). The medium and WST-8 test solution was then added, and the plate was incubated at 37 ◦C for 1h, after which the cell supernatant was placed in new 96-well plates and the absorbance at 450nm was measured using a microplate reader (Model 680, Bio-Rad, Hercules, CA, USA). The test medium was carefully transferred to the new 96-well plate to avoid NP contamination in the medium. Blank and medium controls (medium containing NPs without cells) were treated identically. Nuclear morphology and distribution were investigated by immunocytochemistry. An established biomarker for genomic instability was carried out to evaluate the susceptibility of the cells to NP-induced chromosomal damage endpoints (micronuclei and deformed nuclei). Following exposure to the NPs, the cells were incubated with Hoechst 33342 (Dojindo) for 30min. The number of cells exhibiting micronuclei and deformed nuclei was counted, with over 400 nuclei analyzed per treatment time point and all analyses performed in dental mat e r ial s x x x ( 2 0 1 5 ) xxx–xxx 3 a The NP size values are given by the supplier. The morphology (TEM), crystallinity (XRD) and surface charge (zeta potential analyzer) of the NPs are obtained in this study. triplicate. The cell numbers were scored under laser scanning confocal microscopy (TS1200-E; Nikon, Tokyo, Japan). After each incubation period under test conditions, the cells were rinsed twice with PBS and then incubated with Hoechst 33342 and propidium iodine (PI; Dojindo) as nuclear probes in PBS for 30min at 37 ◦C. Use of Hoechst 33342 and PI staining helped distinguish between apoptosis and necrosis owing to the difference in the permeability of the cell membranes of live and damaged cells. Subsequently, the cells were rinsed three times with PBS and analyzed using laser scanning confocal microscopy (TS1200-E) at 40× magnification via oil. The groups were categorized such that Hoechst (+)/PI (−) were living cells, and Hoechst (+)/PI (+) were late apoptosis cells. The cells stained with PI alone (Hoechst(−)/PI (+)) were represented as necrotic cells that were excluded from the data. The DNA damage in RAW264 cells was analyzed using the alkaline comet assay. Treated cells were rinsed with PBS and resuspended in Hank's balanced salt solution (Sigma–Aldrich) with 10% dimethyl sulfoxide (DMSO) and (ethylenedinitriol)tetraacetic acid (EDTA). The cells were embedded in agarose gel on comet slides (Trevigen, Gaithersburg, MD, USA) and lysed in a prechilled lysis solution (2.5 M NaCl, 0.1 M EDTA, 10mA Tris base, pH 10) with 1% Triton X for 1h at 4 ◦C. The cells were then subjected to denaturation in an alkaline buffer (0.3 M NaCl, pH 7.5) for 20min. Electrophoresis was performed at 21 V and 300mA for 30min in a cooled chamber. After electrophoresis, the slides were washed with water and dehydrated using 70% ethanol. The slides were subsequently stained with SYBR® green dye (Invitrogen, Carlsbad, CA, USA). Images of the samples were observed and captured using fluorescence microscopy (TS1200-E). For the scoring method, a total of 50 comets on each gel were classified as belonging to one of five categories according to the tail and head intensity. Each category was given a value between 0 and 4, and the total scores of the slide could be between 0 and 400 arbitrary units [7]. For SEM and TEM analysis, the cells incubated with and without NPs were washed using PBS, and fixed with 2.5% Fig. 2 – Cell viability of RAW264 cells cultured in a control medium and other media containing 200 and 400 g/mL of Al2O3NPs and SiO2NPs. The optical density values are calculated as a percentage of the control group value. The different lower case letters above each group denotes a statistically significant difference compared with the other group (n = 6, p < 0.05). 4 dental mat e r ial s x x x ( 2 0 1 5 ) xxx–xxx glutaraldehyde for 24h. The cells were then post-fixed in 1% osmium tetraoxide for 1h and subsequently washed and dehydrated in graded concentrations of ethanol (50, 60, 70, 80, 90, and 100%) and hexamethyl-disilazane (Wako). For SEM analysis, the specimens were sputter-coated with gold and observed using SEM (S-3500N, Hitachi, Tokyo, Japan) with an accelerating voltage of 15 kV in a vacuum. For TEM analysis, the cell specimens were infiltrated and embedded in Epoxy resin (EPON 812, TAAB, Berks, UK). Ultrathin sections (90nm) were subsequently cut using a diamond knife on a microtome (Sorvall MT-5000, Du Pont, Wilmington, DE, USA). The sections were collected on copper grids and stained with saturated aqueous uranyl acetate, counter stained with 4% lead citrate, and observed with the TEM (H-7100; Hitachi, Tokyo, Japan). Three independent experiments were conducted, and the results were expressed as the mean ± standard deviation. The Mann–Whitney U test was applied to compare the mean values for the control and treatment groups of cell viability (WST-8 assay) and genotoxicity (frequency of deformed nuclei, micronuclei, Hoechst/PI apoptosis and DNA damage). Statistically, the treatment groups were significantly different from the control groups (p < 0.05 for each). The spherical shapes of the NPs are seen in the TEM images (Fig. 1). The crystallinity is found to be amorphous for both types of NPs using XRD (not shown), and the zeta potential of the Al2O3NPs and SiO2NPs is measured as −16.8 and −9.9mV, respectively (Table 1). The effects the nanoparticles have upon cell viability (WST-8 assay) are illustrated in Fig. 2. The cell viability is significantly reduced after NP exposure (p < 0.05) for both types of NPs, though the cell viability of the Al2O3NP group is significantly greater than that of the SiO2NP group (p < 0.05). dental mat e r ial s x x x ( 2 0 1 5 ) xxx–xxx 5 Fig. 4 – Genotoxicity of 200 and 400 g/mL of Al2O3NPs or SiO2NPs plotted as a function of (a) the frequency of deformed nuclei, (b) the frequency of micro-nuclei, (c) the frequency of Hoechst 33342 and PI staining, and (d) the comet assay for DNA damage. Different lower case letters denote a statistically significant difference compared with the other group (n = 6, p < 0.05). Fig. 3 displays typical confocal microscopic images of the cells. Round nuclei are found in the normal control cells (Fig. 3a), but after exposure to NPs we observe half-moon-shaped nuclei in deformed damaged cells (Fig. 3b). Images of a micronuclei and apoptotic cell are shown in Fig. 3c and d, respectively. Fig. 4 plots the frequency of the occurrence of deformed nuclei (Fig. 4a), micro-nuclei (Fig. 4b), Hoechst/PI apoptosis (Fig. 4c), and DNA damage in the comet assay (Fig. 4d). The frequency of deformed nuclei and DNA damage from the Al2O3NP group is significantly lower than that of the SiO2NPs (p < 0.05). However, no significant differences were found in the frequency of micro-nuclei occurrence (p > 0.05). Fig. 5 shows the SEM images of RAW264 cells in the control group (Fig. 5a) and the groups treated with Al2O3NPs (Fig. 5b) and SiO2NPs (Fig. 5c). Particles and aggregates are clearly observed in the cell surface of the Al2O3NP group. However, the amount of SiO2NP aggregates (Fig. 5c) is less than that of Al2O3NPs aggregates (Fig. 5b). Fig. 6 shows TEM images of RAW264 cells in the control group (Fig. 6a), and cells in groups treated with Al2O3NPs exposure (Fig. 6b–d) and SiO2NP exposure (Fig. 6e and f). The internalized NPs are mainly observed in the cell vesicles for both types of NPs, though NP internalization is also frequently observed in the nuclei (Fig. 6c and f). However, the sizes of the particles in the nucleus are larger, than those seen in the vesicles. The number of NPs observed in the plasma is less than the number observed in the lysosomes. There is a lack of basic toxicological data available for the dental field, despite the fact that a large number of nanoparticle are being used in practical applications. Although some studies have shown that Si and Al NPs are relatively safe materials, our aim in this study is to observe the cytotoxical or genotoxical changes of cells under exposure to Si and Al NPs. The NP concentrations in this study (200 and 400g/mL) are relatively large, but this makes it possible to easily obtain the cytotoxic and genotoxic effects upon and micromorphological changes of the cells. In addition, macrophages are highly specialized in phagocytosis of NPs. To better understand the morphology of the internalization of NPs in cells and the process of engulfing the NPs via the cell membrane, the RAW264 macrophages are selected for this study because of their suitability for this type of research. However, further research is required to measure the effect of NPs upon cells at low concentrations, such as below 100g/mL,to evaluate the potential genotoxic effects at non- or low-toxic concentrations of the tested NPs. The current explanation of the toxic mechanism for solubility of nanoparticles is that particles are digested and dissolved by the acidity of the lysosomes (pH = 5.5), resulting in a release of toxic metal ions from the core metals [16,17]. Finally, the intercellular release of free metal ions generates radicals in the cells that harm the activity of mitochondria and nuclei [18]. The distribution of NPs in the cells is observed using TEM. In Fig. 6, the NPs are clearly visible in the vesicles as large aggregates for both Al2O3NPs (Fig. 6b and c) and SiO2NPs (Fig. 6e). Most of the internalized particles are seen in the vesicles, less are seen in the nuclei, and little or none are seen in the cytoplasm. Because aggregated NPs in vesicles are the typical morphological phase seen in this test, these TEM observations are in line with the above intercellular dissolution mechanism. Therefore, we have to reject the null hypothesis. dental mat e r ial s x x x ( 2 0 1 5 ) xxx–xxx 7 Fig. 6 – (a–d) TEM micrographs of RAW264 cells incubated with Al2O3NPs concentrations of (a) 0 g/mL (control) and (b–d) 400 g/mL. The location of the particle aggregates are observed in the vesicles in (b) and (c) and in the nuclei in (c). The beginning of an endocytosis process seems to be occurring in (d). (e, f) TEM micrographs of RAW264 incubated with 200 g/mL of SiO2NPs. The nanoparticles tend to agglomerate in micron-sized clusters inside the vesicles, which make the lysosomes larger, as seen in (e). Several particles aggregates are seen in nuclei in (f), though the NPs found in vesicles are most likely in lysosomes. Black arrows indicate NPs, and the nuclei are labeled with an N. internalization is greater for Al2O3NPs. However, controversial cytotoxic results are obtained in this study, indicating that the intercellular dissolution rate of SiO2NPs may be greater than that of Al2O3NPs. Hoechst/PI apoptosis frequency, and DNA damages of the comet assay) are observed at NP exposure concentrations of 200 and 400g/mL. Many NPs are seen to aggregate in vesicles, with volumes on the order of microns. In contrast, large aggregates cannot be found in the nuclei, but careful observation show that the internalized NP size in the nuclei is larger than the NP prior to exposure, suggesting that some NPs exist in tight aggregates in the nuclei. These morphological appearances and the degree of aggregation are different than those exhibited by NPs in the vesicles. Although we have speculated that the NPs can penetrate the nucleus and then directly bind and interact with the DNA, the extent of NP penetration into the nuclei is a relatively minor morphological phase for both 8 dental mat e r ial s x x x ( 2 0 1 5 ) xxx–xxx types of NPs used in this study. Although NP internalization is a minor phenomenon in the TEM images, deformed nuclei or micro-nuclei are easily observed in the confocal microscope images. Based on these results, the direct contact of NPs to induce cell damage is minor compared to the secondary effects from released ions in the cells to other cell organelles. Moreover,the cytotoxicity and genotoxicity are well correlated atthe concentrations used in this study. The existence of a discrepancy such as cytotoxicity (−) and genotoxicity (+) implies important biological and clinical informations. Therefore, further research is required to examine and compare the various tests at lower NP concentrations. This study finds that the difference in the core material (aluminum oxide or silica oxide) of NPs is the main reason for differences in their cytotoxicity or genotoxicity. Because the internalization of NPs into the vesicles is the typical morphological phase, the ionization of the NPs in the vesicles and the subsequent toxic intercellular response of the generated ions greatly affect the activity of the cells. It is difficult to judge if these cytotoxic and genotoxic responses are related to the accumulation of NPs in the nuclei, because the internalization of NPs into the nucleus is a sporadic phenomenon, as evidenced by TEM examination. Typically, the amount of NPs release from dental materials is thought to be low in clinical situations because NPs are immobilized into the mass of the biomaterials. Silicon dioxide NPs, for example, are silanized before being incorporated into the resin composites and, therefore, the rate of release is expected to be quite low. Further research is therefore required to determine the extent of NP cytotoxicity or genotoxicity upon cell,tissue or organ biocompatibility for conditions that are similar to those in clinical environments. This work was supported in part by Grants-in-Aid for Scientific Research (C) (Nos. 26462953 and 24659846) from the Japan Society for the Promotion of Science (JSPS). The authors wish to express appreciation for laboratory support from Dr. H. Toshima and Prof. K. Endo (Health Sciences University of Hokkaido, Japan). intracellular solubility: comparison of stabilized copper metal and degradable copper oxide nanoparticles. Toxicol Lett 2010;1907:169–74. nanoparticle and carbon nanotube toxicity and intracellular accumulation in A549 human pneumocytes. Toxicology 2008;253:137–46. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 200.0, \"viab\": 54.026}, {\"h\": 24.0, \"mgL\": 400.0, \"viab\": 54.319}]}", "role": "assistant" } ]	97.json
[ { "content": "Extract sample context and measurement list. <doc> 1 Pharmacology and Toxicology Division, Defence Research and Development Establishment, Jhansi Road, Gwalior 474002, India 2 Saveetha University, P.H. Road, Chennai 600077, India Correspondence should be addressed to Devashri Sahu; Received 12 September 2013; Revised 22 November 2013; Accepted 1 December 2013; Published 5 February 2014 Academic Editor: Irfan Rahman 2014 Devashri Sahu et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. Increased levels of particulate air pollution are associated with increased respiratory and cardiovascular mortality and morbidity. Some epidemiologic and toxicological researches suggest ultrafine particles (<100 nm) to be more harmful per unit mass than larger particles. In the present study, the effect of particle size (nano and micro) of carbon black (CB) particle on viability, phagocytosis, cytokine induction, and DNA damage in human monocytes, THP-1 cells, was analysed. The cells were incubated with nanosize (∼50 nm) and micron (∼500 nm) size of CB particles in a concentration range of 50–800 g/mL. The parameters like MTT assay, phagocytosis assay, ELISA, gene expression, and DNA analysis were studied. Exposure to nano- and micron-sized CB particles showed size- and concentration dependent decrease in cell viability and significant increase in proinflammatory cytokines IL-1, TNF- and IL-6 as well as chemokine IL-8 release. Gene expression study showed upregulation of monocyte chemoattractant protein-1 gene while cyclooxygenase-2 gene remained unaffected. Nano CB particles altered the phagocytic capacity of monocytes although micron CB had no significant effect. CB particles did not show any significant effect on DNA of monocytes. The investigations indicate that CB particles in nanosize exhibit higher propensity of inducing cytotoxicity, inflammation, and altered phagocytosis in human monocytes than their micron size. The nanoparticle industry has expanded substantially in recent years leading to exposure of various nanomaterials to human and environment. Particle size plays an important role in determining the particular biological behavior of nanomaterials. Due to their extreme small size, nanoparticles possess specific large surface area, which makes the number of surface atoms or molecules increasing exponentially. Hence, particles at nanorange exhibit much higher chemical and biological reactivity than fine particles [1]. The risks associated with nanoparticles exposure require investigation due to evidence that these particles can be more inflammogenic and toxic than larger particles comprising of the same material [2]. In recent years, size-dependent toxicity between micro- and nanoscale particles has been demonstrated [3–5]. Carbon black (CB) has wide industrial applications, and used as a reinforcement agent in rubber products, black pigment in printing inks and lithography, electrode for batteries in electrical conductors and during the finishing process of leather goods production. Additionally, carbonaceous nanoparticles are present as an environmental contaminant. Combustion processes are a significant source of carbon nanoparticles. Elemental carbon-based nanoparticles with a diameter of less than 100 nm are a major part of diesel exhaust and ambient pollution. After deposition in the lungs, larger particles are phagocytized by alveolar and airway macrophages [6, 7], but the fine and ultrafine carbon particles remain in the lungs for a longer period of time [8]. Ultrafine particles are phagocytized to a minor extend but they can still enter macrophages and epithelial cells and even penetrate into the circulation. Thus, ultrafine particles not only trigger local inflammatory reactions in the lungs but also cause systemic extrapulmonary effects [9]. Ultrafine particles also have the capacity to inhibit phagocytosis by alveolar macrophages [10]. Macrophages and their monocyte progenitors are major elements of the inflammatory response. In addition to performing phagocytosis, they can release inflammatory mediators such as cytokines and chemokines, crucially involved in destruction of microbes and particles using various enzymatic systems [11]. CB nanoparticles are reported to cause cytotoxic injury, increase levels of proinflammatory chemokines, and inhibit cell growth [12]. Epidemiological as well as experimental studies have confirmed the role of CB nanoparticles in aggravating pulmonary disorders such as asthma, lung cancer, pulmonary fibrosis, and systemic cardiovascular disorders [13]. In this study, CB particles were chosen considering their production in huge quantities posing high environmental risk compromising health of general population [13, 14]. Because of the sporadic information on in vitro size-dependent effect of CB particles, the present study was conducted to determine the effect of their nano- and micron-sized particles on viability, phagocytosis, and cytokine induction in human monocytes, THP-1 cells. These undifferentiated cells express many of the properties of monocytes and represent a model of innate immune system [15]. These cells are an essential link between the adaptive and innate immune responses because they develop into various forms of antigen-presenting cells (macrophages and dendritic cells). They are often used as a model to study human inflammatory responses, which allow for the possibility of elucidating the interactions of nanoparticles with innate immune cells [16, 17]. 2.1. Particle Preparation and Characterization. Carbon nanopowder <50 nm and carbon powder ∼500 nm were purchased from Sigma-Aldrich. Physicochemical properties of particles were analyzed using transmission electron microscopy (TEM), dynamic light scattering (DLS), and zeta potential analyzer. The morphology and size of particles in the stock dispersion were determined by TEM. Dry powder of particles was suspended in cell culture medium at a concentration of 1 mg/mL and then sonicated at room temperature for 10 minutes to form a homogeneous suspension. After sonication and stabilization, the TEM samples were prepared by drop coating of the stock suspension on carbon-coated copper grids. The films on the grids were allowed to dry prior to measurement. TEM measurements were performed at an accelerating voltage of 120 kV (Model 1200EX, JEOL Ltd., Tokyo, Japan). ZetaPALS (Brookhaven Instruments Corporation, Holtsville, NY) was used to determine the hydrodynamic size and zeta potential of particle suspension in cell culture medium. 2.2. Cell Culture. The human monocytic cell line, THP-1 was obtained from National Centre for Cell Sciences, Pune, India. They were maintained in RPMI 1640 medium supplemented with 10% heat inactivated fetal bovine serum (FBS), Lglutamine (2 mM), streptomycin (100 g/mL), and penicillin (100 U/mL). Cells were cultured at 37∘ C in a humidified atmosphere containing 5% CO2. THP-1, cells were seeded into 24-well plates at 1 × 106 cells/mL and exposed to particles in the concentration range of 50 to 800 g/mL for 24 h. Based on the results of screening study (data not shown) done with different concentrations of particles and previous reports [18, 19], a concentration range was selected. The cells were exposed within 10 min of preparation of nano- and micro-CB suspensions. Cell-free controls were included in order to assess the interference of particles with each assay. 2.3. Assessment of Cytotoxicity. THP-1 cells were incubated with CB particles (nano and micro) in a concentration range of 50, 100, 200, 400, and 800 g/mL for 24 h. Following this incubation period, cytotoxicity of particles was assessed using MTT assay [20]. Briefly, MTT (20 L per well of 5 mg/mL stock) was added and incubated for 4 h. Supernatants were removed by centrifugation and then 300 L of DMSO was added. After thorough mixing, optical density at 570 nm was detected by microplate reader (BioTek, USA). Control values (without stimuli) were set at 100% viable and all values were expressed as a percentage of the control and respective TC-50 (particle concentration inducing 50% cell mortality) concentrations were calculated using GraphPad Prism software. 2.4. Phagocytosis Assay. The phagocytic ability of THP-1 cells after 4 h exposure to varying concentrations of CB particles (namely, 1/2TC,50, TC50 and 2TC50) was assessed by measuring their ability to phagocytose 1 m latex beads (1 m Latex beads, carboxylate modified polystyrene, and fluorescent yellow-green). The method used was modified method of Schroeder and Kinden [21]. After exposure, cells were washed two times with PBS to remove excess particles. Culture medium containing latex beads at a bead-to-cell ratio of 10 : 1 was transferred to the culture wells. Monocyte and bead suspensions were then incubated for 1 h to allow phagocytosis. Beads not phagocytosed were removed by centrifugation at 225 g for 5 min; the cell pellet was then resuspended in phosphate buffer saline. The process was repeated three times and finally the cells were vortexed for 10 s and fluorescence of the cells was determined at an excitation and emission wavelengths set at 440 and 485 nm, respectively. Cell viability during phagocytosis assay was monitored by trypan blue exclusion. Viability was 95 ± 5% throughout the assay. Microscopic images of phagocytosed latex beads, only at one concentration, that is, TC-50 concentration of test particles, were taken. After washing, the cells were seen under microscope. DIC images or paired DIC and fluorescence images of phagocytosed beads by monocytes were acquired using fluorescent microscope (Zeiss, Germany) with 40x dry and 100x (oil immersion) objectives. 2.5. Cytokine Analysis. To investigate the effect of CB particles on cytokine production, an enzyme linked immunosorbent assay (ELISA) was performed. For determination of IL-1, IL-6, TNF-, and IL-8, monocytes were cultured at 1 × 106 cells/mL and were exposed to TC-50 concentration of CB particles (nano carbon: 591.4 g/mL and micro carbon 687.1 g/mL) for 6, 18, 24, and 48 h. After particle exposure, cell-free supernatants were harvested via successive 10 min centrifugations (2,000 rpm, 7,000 rpm, and 13,000 rpm) and a According to the manufacturer Sigma-Aldrich. b Using transmission electron microscopy. c,d, and eUsing ZetaPALS. PDI: polydispersity index. stored at −80∘ C until analysis. ELISA was performed according to the manufacturer's protocol (Abcam ELISA Kit) and absorbance values were measured using microplate reader (BioTek Instruments, USA). A minimum of three independent experiments were performed and concentrations calculated from the linear regression equation were derived from a set of standard absorbance values. 2.6. Gene Expression Analysis. Gene expression analysis of COX-2 and MCP-1 was evaluated by exposing cells to TC-50 concentration of CB particles (nano carbon: 591.4 g/mL and micro carbon 687.1 g/mL) for 24 h. Total RNA were isolated using RNeasy Mini Kit (Qiagen, USA).The concentration and integrity of RNA was measured using multimode microplate reader (BioTek, USA) prior to the experiment. The Enhanced Avian HS RT-PCR kit (Sigma, USA) was used for the amplification of COX-2, MCP-1, and 18 sRNA gene, according to the manufacturer's instructions. Amplified cDNA products were separated on 1.2% agarose gel by electrophoresis. The primer sequences of amplified genes were shown in Table 1. 2.7. Detection of DNA Damage. THP-1 cells exposed to nano- and microparticles of CB (0, 50, 100, 200, 400, and 800 g/mL) for 24 h were collected into tubes and washed with PBS. The cells were incubated for 3 h in lysis buffer (20 mM Tris-HCl, pH 8.0, 5 mM EDTA, 0.1 M NaCl, 0.5% SDS, and 100 g/mL RNase) at 37∘ C. After incubation, phenol : chloroform (1 : 1) mixture was used to extract DNA. By adding an equal volume of ice-cold absolute isopropanol, DNA was precipitated. DNA was dissolved in 50 L of 1X TE (10 mM Tris, 1 mM EDTA, and pH 8.0) buffer. 20 g of DNA was loaded onto 1.2% agarose gel and electrophoresis was carried out at 60 V for 2 h with TBE as the running buffer. DNA in the gel was visualized under UV light [22]. 2.8. Statistical Analysis. Statistical analysis was carried out with GraphPad Prism 4 statistical software (Graphpad Software Inc., CA, USA). One-way analysis of variance (ANOVA) with Tukey's method for multiple comparisons was used to evaluate the various responses induced by different concentrations of particles and statistical comparisons between particle sizes were performed with two-way ANOVA, followed by a Bonferroni posttest. Differences were considered statistically significant when the value was less than 0.05. 3.1. Particle Characterization. TEM analysis was performed to determine the morphology and size of the particle and micrographs are shown in Figure 1. The particles were found nearly spherical and cubical in shape. The primary sizes of the particles estimated from TEM images (Figure 1) were presented in Table 2. Since nanoparticles often form agglomerates in a solution, the hydrodynamic sizes of the dispersed particles and their agglomerates in cell culture medium were estimated using ZetaPALS. These values were found larger than the per se particle size measured by TEM (Table 2). 3.2. Cell Viability. After 24 h exposure of THP-1 cells to varying doses (50, 100, 200, 400, and 800 g/mL) of nano- and micro-CB particles, cellular metabolic activity was detected by MTT assay. The cell viability decreased in a concentrationand size-dependent manner following exposure to CB particle (Figure 2) and a significant decrease was observed at concentration from 100 to 800 g/mL for nano carbon particles and at 400 and 800 g/mL for micro carbon particles. The nano carbon particles decreased the percentage of cell viability from 82% to 41% while micro carbon particles decreased it from 69% to 56%. The significant size selective difference ( < 0.01) between nano- and micro carbon was found only at highest concentration of 800 g/mL. The TC-50 value (particle concentration causing 50% cell mortality) calculated for nano- and micro carbon was found to be 591.4 g/mL and 687.1 g/mL, respectively (Table 3). 3.3. Phagocytic Capacity of THP-1 Cells. The phagocytic ability of the monocytes was measured after the uptake of test particles. The assay was carried out by taking 3 different concentrations of test particle, namely, 1/2 TC-50, TC-50, and 2TC-50 for each particle. A significant reduction in the phagocytosis of indicator latex beads occurred after exposure to all concentrations of nano-CB particles ( < 0.01), while no significant reduction was observed with their microsize (Figure 3). Fluorescent images represent monocytes phagocytosing latex beads (Figure 4). Qualitatively control cells were found to phagocytose more beads as compared to nano-CB particle exposed monocytes whereas the phagocytic capacity of monocytes was not affected by the presence of micro-CB particles (Figure 5). 3.5. MCP-1 and COX-2 mRNA Expression. A significant upregulation of MCP-1 gene expression was observed with both sizes of CB particles (Figure 7(a)). Micron size of CB particles exhibited more expression of MCP-1 gene than nanosize (Figure 7(c)). However, no significant change was observed in COX-2 gene expression by both sizes of CB particles (Figures 7(a) and 7(b)) as compared to control. 3.6. DNA Analysis of THP-1 Cells. DNA damage was studied by observing smearing pattern on agarose gel under UV light. Exposure of THP-1 cells to both sizes of CB particles for 24 h did not exhibit any smearing pattern on agarose gel (Figure 8). The inconsistencies of the effects of different CB samples in relation to carcinogenicity and toxicity were reported [23, 24]. These inconsistencies are likely to be related to variation in the particle sizes between the different samples. Therefore, the present in vitro study compared the potential differences in the ability of nano- and micro-CB particles to produce toxicity on exposure to human monocytes, THP-1. The physicochemical characteristics of nano- and micron size of CB particles were extensively characterized. The average size of the particles was in agreement with the size provided by the supplier. DLS analysis revealed a hydrodynamic size of dispersed particles in cultured medium indicating agglomeration in aqueous media which was further corroborated by zeta potential measurement. Needless to surprise, the mean particle sizes and size distribution of particles (measured by DLS) were found enhanced when measured in aqueous media compared to measurements in the dry phase (measured by TEM). To minimize the effects of particles aggregation and sedimentation, suspension of particles was always freshly prepared and sonicated before each experiment. In this study, the cytotoxicity results demonstrated that nanoparticles of CB were more potent than its micron size in causing toxicity to THP-1 cells as revealed by MTT assay. Both Microcarbon sizes of particles followed concentration-dependent toxicity in monocytes. The effect of CB particle on phagocytic ability of monocytes was also demonstrated. It was found that only nano-CB particle impaired the phagocytic capacity of monocytes. Our findings of impaired phagocytosis in nano-CB particle exposed cells support the findings of Renwick et al. who demonstrated that ultrafine (CB and TiO2) particles impaired macrophage phagocytosis to a greater extent than fine particles compared on a mass basis [25]. The study of inflammatory responses of THP-1 cells on exposure to nano- and micro particles of CB revealed the induction of proinflammatory cytokines IL-1, IL-6, and TNF-. Many cytokines including IL-1, IL-6, and TNF activate functions of inflammatory cells during acute inflammatory responses. These cytokines increase the vascular permeability and thus cause swelling and redness associated with inflammation. IL-1 and IL-6 are responsible for fever reactions while TNF- stimulates endothelial cells and is responsible for hypotension [26]. The nanoform of CB particle showed increase in release of IL-1 and IL-6 with increasing time of exposure up to 24 h and then declined thereafter distinctly from its micron form. TNF secretion was induced at early time point, that is, at 6 h, which later on decreased with time. It was observed that the secretion of IL-1, IL-6, and TNF- cytokines was more profound with nanosize than with micron size. This suggests that the response of particles to proinflammatory cytokine release was size dependent. Chemokines are secondary proinflammatory mediators; that is, they are typically induced by primary proinflammatory mediators such as IL-1 or TNF. In the present study, gene expression of MCP-1, a CC chemokine, and release of IL-8, a member of the CXC chemokine subfamily, were studied. MCP-1 stimulates both chemotaxis of monocytes and several cellular events associated with chemotaxis. The result of the present study demonstrated the stimulation of chemotaxis as revealed by upregulation of MCP-1 gene and increased release of IL-8 on exposure to CB particles. Niwa et al. [27] have also shown the upregulation of IL-6 and MCP-1 in rats after inhalation exposure of CB and substantiate the findings. IL-8 is a chemokine which plays a key role in the activation of neutrophils and their recruitment to the site of inflammation [28]. Results demonstrated increase in the release of IL-8 with time after exposure to both sizes of CB particles. Kim et al. [29] reported increased expression of IL-8 mRNA and protein on exposure of ultrafine carbon particles in normal human bronchial epithelial cells and supported the findings. Noticeably, the MCP-1 gene expression was more upregulated by micron size of CB than nanosize. Similar results were observed with another chemokine, IL-8, where micron size of CB was able to induce more secretion of IL-8 at later time of exposure. Another inflammatory marker, COX-2, was also studied. COX-2, an inducible isoform of cyclooxygenase, is induced by several mitogenic and proinflammatory stimuli including LPS, interleukin-1 (IL-1 and IL-1), and TNF- [30, 31]. In the present study, CB particles did not show any change in the COX-2 expression. Overall, in accordance with the results of Ferin et al. [32] and Li et al. [33], it was found that nano-CB induced a greater toxicity and inflammatory response than micro-CB in human monocytes. In the present study, the effect of CB particles on DNA damage of human monocytes was also studied. In assessing toxicity, DNA damage to macrophages is an important outcome since (i) these cells remove inhaled NPs [34] and (ii) DNA damage is considered to be an important initial event in various diseases including carcinogenesis [35]. The results showed that there was no smearing pattern observed on agarose gel indicating that both sizes of CB particles did not cause any DNA damage. In conclusion, a comparative toxicity study between nano- and micro-CB particle resulted in size-dependent cytotoxicity and increased inflammatory responses in human monocytes, THP-1 cells. Nano-CB particles altered the phagocytic capacity of monocytes although micro-CB had no significant effect on phagocytic capacity of monocytes. However, both sizes of CB particle did not have any effect on DNA of THP-1 cells. Further studies are required to elucidate the exact pathway of inflammatory response induced by CB particles in immune cells. The authors declare that there is no conflict of interests. The authors are thankful to the Director of Defence Research and Development Establishment, Gwalior, India, for providing facility. They acknowledge Advanced Instrumentation Research Facility (AIRF), Jawaharlal Nehru University (JNU), New Delhi, India, for nanoparticle characterization by TEM. The hydrodynamic size and zeta potential measurements of particles were performed using facilities at CeNSE, funded by Department of Information Technology, Government, of India and Indian Institute of Science, Bangalore, India. the immune system and its potential effects on nanoparticle biodistribution,\" Molecular Pharmaceutics, vol. 5, no. 4, pp. 487– 495, 2008. of International Journal of Inflammation is the property of Hindawi Publishing Corporation and its content may not be copied or emailed to multiple sites or posted to a listserv without the holder's express written permission. However, users may print, download, or email articles for individual use. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 235.5, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 50.0, \"viab\": 87.918}, {\"h\": 24.0, \"mgL\": 100.0, \"viab\": 81.877}, {\"h\": 24.0, \"mgL\": 200.0, \"viab\": 77.763}, {\"h\": 24.0, \"mgL\": 400.0, \"viab\": 56.684}, {\"h\": 24.0, \"mgL\": 800.0, \"viab\": 40.617}]}", "role": "assistant" } ]	89.json
[ { "content": "Extract sample context and measurement list. <doc> Lucie Armand, Adeline Tarantini, David Beal, Mathilde Biola-Clier, Laure Bobyk, Sephanie Sorieul, Karin Pernet-Gallay, Caroline Desvergne, Iseult Lynch, Nathalie Herlin-Boime, Marie Carriere Doi: Titanium dioxide nanoparticles (TiO2 -NPs) are one of the most produced NPs in the world. Their toxicity has been studied for a decade using acute exposure scenarios, i.e. high exposure concentrations and short exposure times. In the present study, we evaluated their genotoxic impact using long-term and low concentration exposure conditions. A549 alveolar epithelial cells were continuously exposed to 1-50 μg/mL TiO2 -NPs, 86% anatase/14% rutile, 24±6 nm average primary diameter, for up to two months. Their cytotoxicity, oxidative potential and intracellular accumulation were evaluated using MTT assay and reactive oxygen species measurement, transmission electron microscopy observation, micro-particle-induced X-ray emission and inductively-coupled plasma mass spectroscopy. Genotoxic impact was assessed using alkaline and Fpg-modified comet assay, immunostaining of 53BP1 foci and the cytokinesis-blocked micronucleus assay. Finally, we evaluated the impact of a subsequent exposure of these cells to the alkylating agent methyl methanesulfonate. We demonstrate that long-term exposure to TiO2 -NPs does not affect cell viability but causes DNA damage, particularly oxidative damage to DNA and increased 53BP1 foci counts, correlated with increased intracellular accumulation of NPs. In addition, exposure over 2 months causes cellular responses suggestive of adaptation, characterized by decreased proliferation rate and stabilization of TiO2 -NP intracellular accumulation, as well as sensitization to MMS. Taken together, these data underline the genotoxic impact and sensitization effect of long-term exposure of lung alveolar epithelial cells to low levels of TiO2 -NPs. 2016 Taylor & Francis. 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. Lucie Armand1,2 , Adeline Tarantini1,2 , David Beal1,2, Mathilde Biola-Clier1,2 , Laure Bobyk1,2 , Sephanie Sorieul 3 , Karin Pernet-Gallay4,5 , Caroline Desvergne6 , Iseult Lynch7 , Nathalie Herlin-Boime8 , Marie Carriere1,2,§ 1Université Grenoble-Alpes, INAC-LCIB, Laboratoire Lésions des Acides Nucléiques, 17 rue des Martyrs, F-38000 Grenoble, France. 2CEA, INAC-SCIB, Laboratoire Lésions des Acides Nucléiques, 17 rue des Martyrs, F-38000 Grenoble, France. 3CENBG, Université Bordeaux 1, IN2P3, UMR5797, 33175 Gradignan Cedex, France. 4Université Grenoble Alpes, Grenoble Institut des Neurosciences, Grenoble F-38000, France 5 INSERM U 836, F-38000 Grenoble, France. 6Université Grenoble-Alpes, CEA, Nanosafety Platform, Medical Biology Laboratory (LBM), 17 rue des Martyrs, F-38054 Grenoble, France. 7 School of Geography, Earth and Environmental Sciences, University of Birmingham, Edgbaston, B15 2TT Birmingham, United Kingdom JUST ACCEPTED 8UMR3685 CEA-CNRS, NIMBE, LEDNA, CEA Saclay, F-91191 Gif sur Yvette, France. §Corresponding author: Marie Carrière, CEA Grenoble, LAN, Bât C5, pce 632, 38054 Grenoble Cedex 9, France. Phone: +33 4 38 78 03 28, fax: +33 4 38 78 50 90. Nanoparticle, TiO2, toxicity, genotoxicity, chronic Titanium dioxide nanoparticles (TiO2-NPs) are one of the most produced NPs in the world. Their toxicity has been studied for a decade using acute exposure scenarios, i.e. high exposure concentrations and short exposure times. In the present study, we evaluated their genotoxic impact using long-term and low concentration exposure conditions. A549 alveolar epithelial cells were continuously exposed to 1-50 μg/mL TiO2-NPs, 86% anatase / 14% rutile, 24±6 nm average primary diameter, for up to two months. Their cytotoxicity, oxidative potential and intracellular accumulation were evaluated using MTT assay and reactive oxygen species measurement, transmission electron microscopy observation, micro-particle-induced X-ray emission and inductively-coupled plasma mass spectroscopy. Genotoxic impact was assessed using alkaline and Fpg-modified comet assay, immunostaining of 53BP1 foci and the cytokinesis-blocked micronucleus assay. Finally, we evaluated the impact of a subsequent exposure of these cells to the alkylating agent methyl methanesulfonate. We demonstrate that long-term exposure to TiO2-NPs does not affect cell viability but causes DNA damage, particularly oxidative damage to DNA and increased 53BP1 foci counts, correlated with increased intracellular accumulation of NPs. In addition, exposure over 2 months causes cellular responses suggestive of adaptation, characterized by decreased proliferation rate and stabilization of TiO2-NP intracellular accumulation, as well as sensitization to MMS. Taken together, these data underline the genotoxic impact and sensitization effect of long-term exposure of lung alveolar epithelial cells to low levels of TiO2-NPs. JUST ACCEPTED models, since biological effects and related mechanisms may differ depending on exposure concentration and time. drastic reduction of cellular ability to repair damaged DNA (Jugan, 2012). Our previous work, using 2D-gel electrophoresis analysis together with functional validation to analyze the molecular response of A549 cells to 2 months, chronic exposure to these NPs, showed that this exposure scenario impacts the abundance of 22 gene products involved in glucose metabolism, trafficking, gene expression, mitochondrial function, proteasome activity and DNA damage response (Armand, 2015). This chronic exposure was also found to increase the intracellular content of the serine-threonine kinase receptorassociated protein (STRAP) whose role is to stabilize p53 coactivators p300 and JMY so that they activate p53, which are indicative of DNA damage. Moreover, p53 phosphorylation and acetylation were increased, leading to cell cycle slowdown (Armand, 2015). very detailed picture of key steps to be elucidated. A549 human lung carcinoma cells were purchased from ATCC (Manassas, VA, USA, reference CCL-185) and grown in DMEM 4.5 g/L glucose (Life Technologies, Carlsbad, CA, USA) containing 2 mM/L glutamine, penicillin/streptomycin (50 IU/mL and 50 mg/ml respectively) and 10% FBS. They were maintained at 37°C in a 5% CO2/air incubator. Over a period of up to 2 months, cells were subcultured in cell culture medium containing freshlydiluted NPs twice a week (Figure S1), using the following protocol: wells were rinsed with PBS, trypsinized with trypsin-EDTA 0.25% and counted after trypan blue staining. Cells were then seeded in a new dish (seeding details described in the specific experimental protocols below), in cell culture medium containing 1, 2.5, 5, 10 or 50 μg/mL TiO2-NPs, these concentrations being in line with those used in the previously published TiO2-NP chronic exposure studies (Huang, 2009; Kocbek, 2010; Vales, 2014; Wang, 2011). The 10 mg/mL NP stock suspension (prepared in water) was the same during the whole exposure time. The dispersion state of the suspension was checked by DLS Cytotoxicity was assessed using the 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyl tetrazolium bromide (MTT) assay (Sigma-Aldrich, St. Louis, MO, USA). Cells were seeded at sub-confluence in 96-well plates, in cell culture medium containing 1, 2.5, 5, 10 or 50 μg/mL TiO2-NPs (6 wells per condition). After 16-hours, to allow cell adhesion, the medium in each well was replaced by 0.5 mg/mL MTT diluted DMEM. After 1 h of incubation at 37°C, formazan crystals were dissolved in 100 μL DMSO. To limit interference NPs were then allowed to sediment for 1 h, after which the plate was centrifuged at 450 × g for 5 min. 50 μL per well was transferred to a new 96-well plate. Absorbance was measured at 570 nm and cell viability was determined as percentage of absorbance of negative control (unexposed cells). Trypan blue count (2 replicates per condition) was used to calculate the population doubling level (PDL) at each exposure concentration and at each cell passage, using the following formula: PDL = tlog2/(logNt−logN0) (N0 and Nt being cell counts immediately after seeding and t days after seeding, respectively). JUST ACCEPTED We also used propidium iodide staining to assess cell viability. After exposure to 0, 1, 2.5, 5, 10, 50 μg/mL TiO2-NP, cells were collected (106 cells per condition, 4 replicates) and rinsed with PBS containing 2 mM EDTA (PBS-EDTA). Cells were fixed in ice-cold 70% ethanol for 30 min then washed with PBS-EDTA. They were suspended in PBS-EDTA containing 25 μg/mL propidium iodide (Life Technologies) and 25 μg/mL RNAse A (Sigma-Aldrich). A minimum of 20 000 events per condition was measured by flow cytometry using a FacsCalibur analyzer (BD Biosciences, Franklin Lanes, NJ, USA) equipped with CXP software (Beckman Coulter Inc., Pasadena, CA, USA). Median fluorescence was calculated using Flowing Software 2.5.1 (http://www.flowingsoftware.com/). Fluorescence intensity median results were normalized to untreated control. NP accumulation and intracellular distribution were measured by μ-particle-induced X-ray emission (μPIXE) and Elastic Backscattering (EBS). Cells were seeded onto polycarbonate membranes and allowed to adhere for 16 h. They were then rinsed twice with PBS and cryofixed by immersion in isopentane chilled to -160°C in liquid nitrogen (Carriere, 2005). Samples were freeze-dried for 24 h at - 10°C, 0.37 mbar. Micro-PIXE and EBS spectra were recorded simultaneously on the microbeam line of the AIFIRA platform (CENBG, Bordeaux, France) (Barberet, 2009). The 3.5 MV Singletron accelerator (HVEE) was adjusted in order to deliver a focused beam (2.5 μm in diameter) of 3 MeV protons with a beam current of 1 nA. X-rays were detected with an 80 mm2 Si(Li) detector (Gresham) orientated at 135° with respect to the incident beam axis, and equipped with a 12 μm thick beryllium window. A funny filter (Al, thickness 200 μm, size hole = 1 mm) was used in order to limit the deadtime below 10%. Backscattered protons were recorded at 135° with a silicon PIPS detector (Canberra, 25 mm2 , thickness 100 μm, resolution: 17 keV). Four elemental maps of 100×100 μm2 were recorded on each sample, and drawn using the SupaVISIO software (http://biopixe.free.fr). For Ti intracellular content measurement, data were fitted using SIMNRA (EBS) (Mayer, 2002) and Gupix (X-ray spectra) (Campbell, 2010) as described previously and normalized by analysis of standards, kindly provided by Dr. R. Ortega. JUST ACCEPTED Intracellular distribution was observed by TEM. After exposure cells were rinsed twice with PBS, fixed with 2% glutaraldehyde and 1% osmium tetroxide. They were then gradually dehydrated with ethanol and embedded in Epon resin. Sections were cut by ultramicrotomy (80 nm) and observed on a JEOL 1200EX TEM operating at 80 kV (Grenoble Institut des Neurosciences, Grenoble, France). In cell samples exposed to NPs for 2 months, glutathione concentration, both total (GSH+GSSG) and reduced (GSH), as well as the activity of the 4 antioxidant enzymes glutathione reductase (GR), glutathione peroxidase (GPx), superoxide dismutase (SOD) and catalase (CAT) were monitored as previously described (Barillet, 2010), with 3 replicates per condition. For detailed protocols, see the Electronic Supplementary Information. NP-induced DNA strand breaks and alkali-labile sites were assessed through the alkaline version of the comet assay (Singh, 1988). Oxidatively damaged DNA was quantified by using the bacterial DNA repair enzyme formamidopyrimidine-DNA glycosylase (Fpg) in the comet assay (Gurr, 2005). This enzyme converts oxidized purines, including the major purine oxidation product 8-oxoguanine, into single-strand breaks (SSB, therefore detectable by the comet assay) through a base-excision process (Collins, 2009). At the end of the exposure time, cells (1 well per condition) were rinsed three times with PBS, trypsinized, then pelleted by centrifugation for 5 min at 250 g. Cell pellets were then resuspended in sucrose 85.5 g/L, DMSO 50 mL/L prepared in citrate buffer (11.8 g/L), pH 7.6, and immediately frozen at -80°C. At the end of the 2 months exposure period, samples were thawn and processed as described hereafter for the comet assay. Six microscope slides per condition were coated with 1% normal melting point agarose (NMA) and allowed to dry. 10000 cells per slide were mixed with 0.6% low melting point agarose (LMPA) and deposited over the NMA layer. The cell/LMPA mix was then allowed to solidify on ice for 10 min. Slides were immersed in cold lysis solution (2.5 M NaCl, 100 mM EDTA, 10 mM Tris, 10% DMSO, 1% Triton X-100) overnight at 4°C, before being rinsed in 0.4 M Tris pH 7.4. Then 3 slides were treated with 100 μL Fpg (5 U/slide, in enzyme buffer, Trevigen, Gaithersburg, MD, USA) and 3 slides were incubated with Fpg enzyme buffer for 45 min at 37 °C. DNA was then allowed to unwind for 30 min in alkaline electrophoresis solution (300 mM NaOH, 1 mM EDTA, pH > 13). Electrophoresis was performed in an electric field of 0.7 V/cm and 300 mA for 30 min. Slides were then neutralized in 0.4 M Tris pH 7.4 and were stained with 50 μL of 20 mg/ml ethidium bromide (Life Technologies). As a positive control for alkaline comet assay, we used 50 μM H2O2 directly deposited onto the agarose layer containing the cells, incubated for 5 min at room temperature. As a positive control for comet-Fpg we used A549 cells exposed for 20 min at 37 °C to 1 μM riboflavin, then exposed to 10 J/cm2 of UVA. At least 50 comets per slide were analyzed under a fluorescence microscope (Carl Zeiss, Oberkochen, Germany) connected to a charge-coupled device camera with a 350-390 nm excitation and 456 nm emission filter, at x20 magnification. Comets were measured and analyzed using Comet IV software (Perceptive Instruments, Suffolk, UK). JUST ACCEPTED To evaluate DNA double strand breaks (DSB) or replication fork blockade, p53 binding protein 1 (53BP1) foci were revealed and counted in cell nuclei. 53BP1 is a non-enzymatic protein which is recruited shortly after primary DSB detection. This protein is homogeneously distributed in the nuclei of unperturbed cells then it is recruited within 1-2 min to DSB sites (Bekker-Jensen, 2010); like gamma-H2AX it can therefore serve as a marker for DSB. As a positive control, we exposed cells to 25 μM etoposide (Sigma-Aldrich) for 24 h. Cells were fixed for 20 min in 3% paraformaldehyde (Sigma-Aldrich), stained using anti-53BP1 antibody (Novus Biologicals, Littleton, CO, USA, 1/500 vol./vol.) and slides were mounted with Fluoroshield (Sigma-Aldrich) containing DAPI (1 slides per condition). 53BP1 foci were visualized on an Axio ImageA1 microscope coupled to an Axiocam MRm camera (Carl Zeiss). At least 15 images per condition were captured; on each image we determined both total number of 53BP1 foci and total number of nuclei. Apoptotic cells and dividing cells were rejected. The micronucleus assay, probing aneuploidic and clastogenic events, was performed as described previously (Fenech, 2000). Cells exposed to TiO2-NPs were seeded on glass coverslips (3 coverslips per condition, i.e. 3 independent replicates). As a positive control, we used A549 cells exposed for 24 h to 100 μM methyl methanesulfonate (MMS, Sigma-Aldrich) (3 replicates). They were then cultured for another 28 h in culture medium (without NPs) containing 4 μg/ml cytochalasin B (Sigma-Aldrich) to block cytokinesis. Cells were then immediately fixed for 20 min in 3% paraformaldehyde and stained with acridine orange (5 mg/mL, Life Technologies) for 15 min. Coverslips were mounted on microscope slides with Fluoroshield mounting medium containing DAPI. At least 20 images per coverslip were captured with an Axio ImageA1 microscope (Carl Zeiss). On each image, total nr. of cells, binucleated cells and micronuclei were counted. For each assay, the number of biological replicates per experiment as well as the number of 2 months exposure experiment in which the assay was done is summarized in Table S1. Except for TEM, μPIXE and ICP-MS assays, data curation was carried out as follows: results from biological replicates of each independent experiment were averaged; results presented in figures are mean ± s.e.m. of these averages. For ICP-MS assay, the assay was performed only once, with 4 independent replicates. Data presented in Table S2 are thus mean ± s.e.m. of these 4 independent replicates. For μPIXE, 4 measurements were performed on each of the 2 independent samples (prepared during 2 independent 2 month exposure experiments); data presented in Figure 1 are the mean ± s.e.m. of the 8 values that were obtained. Statistical significance was assessed by Kruskal-Wallis non-parametric one-way analyses of variance by ranks, using the Statistica 8.0 software (Statsoft, Chicago, IL, USA). When significance was demonstrated (p < 0.05), paired comparisons were run using Mann-Whitney tests. JUST ACCEPTED The TiO2-NPs used in the present study, P25 Aeroxide, have already been characterized in terms of morphology, primary size, surface specific area and isoelectric point (Jugan, 2012; Simon-Deckers, 2008). Briefly they are round-shaped (Figure S2) with an average primary particle diameter of 24±6 nm (as measured by TEM) and average specific surface area of 46±1 m2/g. Their crystalline phase is 86% anatase and 14% rutile. We dispersed these NPs in water, using high energy sonication as previously described (Carriere, 2014; Simon-Deckers, 2008). After this dispersion stage in water, their average hydrodynamic diameter, measured by DLS and expressed in number, was 44±25 nm (Figure S2). Their polydispersity index (PdI) was 0.146±0.009. This distribution did not evolve significantly during the 2 months of the experiment; distribution profiles were almost identical 24 h and 2 months after dispersion (not shown). Just before each cell passage, after vigorous vortexing this suspension was diluted in cell culture medium containing 10% FBS to reach NP concentrations of 1, 2.5, 5, 10 or 50 μg/mL. In this exposure medium their zeta potential was -11.2±0.8 mV; their hydrodynamic diameter increased to 342±15 nm and PdI increased to 0.236±0.048, indicating that NPs agglomerated somewhat (Figure S2). JUST ACCEPTED NP accumulation and distribution in exposed cells shows no statistically significant increase of intracellular Ti between 1 and 2 months of exposure (Table S1). At the sub-cellular level, TEM observation of cells exposed for 2 months to 2.5 and 50 μg/mL TiO2- NPs showed that NPs were accumulated inside the cells under both conditions, mostly in cytoplasmic vacuole-like compartments which may be lysosomes or autophagosomes. No NP was observed in the nucleus or mitochondria (Figure 2). No major accumulation of NPs was observed on cell membranes, confirming that the μPIXE and ICP-MS measurements reflected NP accumulation inside the cells rather than NPs adsorbed on cell membranes. The MTT assay, assessing cell mitochondrial activity, demonstrated that chronic exposure to NPs did not affect cell survival, even after 2 months of exposure and at concentrations as high as 50 μg/mL TiO2-NPs (Figure 3A). Interference of NPs with the assay was tested using a previously reported protocol (Brun, 2014) and turned out to be negative (data not shown). This result was confirmed by trypan blue staining and cell counting at each cell passage, that revealed no overt mortality (Table S2), as well as propidium iodide staining and flow cytometry analysis of live/dead cells (Figure 3B), that also did not reveal any overt mortality. JUST ACCEPTED However, the population doubling level (PDL), inversely correlated to cell proliferation, was moderately but significantly increased for the 10 and 50 μg/mL TiO2-NP exposure concentration and for the last passage (Figure 3C). This indicates that the highest exposure concentrations and the longest exposure time, even if non cytotoxic, significantly decreased cell proliferation. as suggested by Guadagnini et al (Guadagnini, 2015); it did not account for more than 10% of the total DCF fluorescence even at the highest concentration, i.e. interference was not significant. We did not observe any significant modulation of both total (GSH+GSSG) and reduced (GSSG) glutathione contents in cells continuously exposed to TiO2-NPs for 2 months (Table S3). Neither did we observe any modulation of the activity of glutathione peroxidase (GPx) and glutathione reductase (GR) which are responsible for glutathione cycling between its reduced and oxidized form. The activity of superoxide dismutase (SOD) and catalase (CAT), involved in hydrogen peroxide and oxygen superoxide radical dismutation, respectively, were also unchanged after 2 months of exposure (Table S3). Finally, we monitored mRNA expression of CAT, SOD1, SOD2, GPx1, GPx4 and GSR and observed no modulation of their expression (Table S4). μg/mL TiO2-NP than in cells exposed to any other concentration of NPs, confirming the concentrationdependence of DNA damage. We also measured the specific presence of DSB or DNA replication fork blockade by counting 53BP1 foci in the nuclei of cells chronically exposed to TiO2-NPs. We observed higher 53BP1 foci counts in cells exposed to 2.5 and 5 μg/mL NPs during 1 month, and in cells exposed to 10 and 50 μg/mL NPs for 1 and 2 months (Figure 5C). Finally, in order to evaluate clastogenic and aneugenic effects of NPs we used the cytokinesis-blocked micronucleus assay. Due to the possibility that cytochalasin B blocked endocytosis and consequently NP cellular uptake, we used the modified version of the assay where cells are incubated with NPs before being treated with cytochalasin B (Magdolenova, 2012). The percentage of binucleated cells, after 28 h of exposure to cytochalasin B of cells chronically exposed to TiO2-NPs, ranged between 55 and 65% and was not statistically different in control cells, as compared to TiO2-NP-exposed cells. In binucleated cells, micronucleus counting did not reveal any chromosomal damage (Table S5). Moreover our positive control (100 μM of MMS for 24 h) significantly increased the number of micronuclei per binucleated cell (30.9±0.1 micronuclei per 1000 cells in cells exposed to MMS vs. 7.3±2.5 micronuclei per 1000 cells in unexposed cells, see Table S5). This proves that the mutagenicity and clastogenicity of TiO2-NPs, under this exposure condition (i.e. chronic low level exposure, 1-50 μg/mL over 2 months), was insignificant. JUST ACCEPTED We previously showed that acute exposure to TiO2-NPs impaired cellular ability to repair DNA lesions via NER and BER pathways (Jugan, 2012). Chronic exposure to TiO2-NPs may thus sensitize cells to other genotoxic agents, particularly those which induce lesions that are classically repaired by the NER and the BER. We thus evaluated the impact of post-exposure to the alkylating agent methanemethyl sulfonate (MMS), which generates DNA lesions repaired by the BER, on cells continuously exposed to TiO2-NPs for 2 months. First, these cells were exposed to a non-lethal concentration of MMS, i.e. 100 μM for 24 h. The MTT assay then demonstrated that cell mortality was higher in cells continuously exposed to 5 to 50 μg/mL NPs and subsequently acutely exposed to MMS, as compared to cells exposed to MMS without previous chronic exposure to NPs (Figure 6A). In order to confirm that this higher cell mortality was related to higher DNA damage, we quantified DNA strand break in cells chronically exposed to TiO2-NPs and then acutely exposed to 50 μM or 100 μM of MMS. Postexposure to 50 μM MMS did not increase the % tail DNA in alkaline comet assay (not shown). Conversely % tail DNA was increased by 14% in cells exposed to 50 μg/mL TiO2-NPs for 2 months then post-exposed to 100 μM of MMS for 24 h, as compared to cells that were only exposed for 24 h to MMS (Figure 6B, grey bars). As a comparison, 2 months of exposure to 50 μg/mL TiO2-NPs increased % tail DNA by 4%, as compared to unexposed cells (Figure 6B, white bars). In addition, 53BP1 foci were counted in cells exposed to TiO2-NPs for 2 months, then post-exposed to 50 μM or 100 μM of MMS. No statistically significant difference was observed in this assay (Figure S3). Finally we analyzed the mRNA expression of genes encoding proteins involved in DNA repair processes, especially via the BER pathway, i.e. APE1, PARP1, PCNA, XRCC1, POLB, LIG3, as well as genes encoding TP53 and the proteins involved in cell cycle checkpoint ATM and ATR, in cells chronically exposed to TiO2-NPs. Their expression was not modified in cells exposed for 2 weeks, 1 month or 2 months to 2.5, 10 or 50 μg/mL TiO2-NPs (Table S6). This indicates that if cells are sensitized to MMS through reduction of their DNA repair capacities, as suggested by our previous results obtained after acute exposure to these TiO2-NPs, this does not rely on the inhibition of expression of genes encoding DNA repair proteins. In the present article, we report the impact of long-term exposure of A549 lung cells to TiO2-NPs, particularly focusing on the genome instability that it may provide. Our previously published results show that this exposure scenario induces DNA damage response via activation of the p53 pathway, through increased levels of STRAP which is a stabilizer of the coactivators of p53, p300 and JMY (Armand, 2015). We also showed that p53 acetylation and phosphorylation levels are increased (Armand, 2015). In the present article we complement these data by showing that this exposure scenario causes the accumulation of reactive oxygen species, oxidative damage to DNA, as well as the formation of foci of 53BP1 on the DNA which suggestss the presence of double-strand breaks and/or DNA repair processes by excision/synthesis and/or replication fork blockade. These lesions to DNA may be the cause of the p53 pathway activation that we previously reported. In this previous study, we also report that chronic exposure induced mitochondrial damage and perturbation of JUST ACCEPTED autophagy (Armand, 2015), which may be an explanation for the accumulation of ROS that we observe here. Indeed mitochondria and cellular respiration are the main source of endogenous ROS in mammalian cells. If these ROS are not appropriately scavenged, or if, as a result of mitochondrial damage, higher levels of ROS are produced, then these ROS would attack biomolecules including DNA. As discussed in our previous article (Jugan, 2012), oxidative damage to DNA may be created by weak oxidizing species such as hydroperoxides. These species may be produced in the cell cytoplasmic vacuole-like structures where TiO2-NP accumulate, then migrate to the nucleus where they would oxidize DNA. Conversely the lifetime of OH°, the only ROS which is able to attack DNA backbone and create strand breaks (Pogozelski, 1998), is very short. Consequently, its migration from the cytoplasm/vacuoles to the nucleus, leading to direct DNA backbone attack, is improbable. The accumulation of 53BP1 foci that we observe might thus reflect the presence of double-strand breaks that result from secondary, indirect production of OH° in the nucleus. Alternatively, and more probably, it may result from the repair of oxidative lesions by excision-synthesis processes, or replication fork blockade during mitosis due to damaged DNA (Bekker-Jensen, 2010). This hypothesis is further suggested by the low number of 53BP1 foci that we observe: in control cells we counted 0.2-0.3 53BP1 foci per cell nucleus while in cells exposed for 1 month to 2.5, 5 or 10 μg/mL TiO2-NPs we observe 0.5 to 0.7 foci per cell nucleus. If secondary production of OH° in the nucleus was the cause of DNA backbone attack and 53BP1 foci formation, then this number would certainly be higher. Moreover, since we did not detect any micronuclei, it appears that these DNA strand breaks are efficiently repaired. JUST ACCEPTED G1 phase due to activation of the p53 pathway (Armand, 2015). Such perturbation of the cell cycle has also previously been reported by others upon acute (Wu, 2010) or long-term (Wang, 2011) exposure to TiO2-NPs. We show that TiO2-NPs accumulate in cells where they distribute in cytoplasmic vesicles. Their accumulation kinetics suggest cellular adaptation to NP exposure, since Ti intracellular content after 2 months of continuous exposure is not statistically higher than after 1 month of exposure. This observation is in line with the results of Wang et al., who observed significant decrease of Ti content in 60-day exposed cells, as compared to 2-day exposed cells (Wang, 2011). This adaptation could either be expulsion of NPs from the cells, for instance by activation of the exocytosis of NP-containing vesicles, or exclusion, i.e. reduced accumulation while dividing cells also divide their Ti content between daughter cells. It was recently demonstrated that NPs are not exported out of cells, and that NP accumulation depends on the phase of the cell cycle, with more efficient uptake in G2/M phase than in S phase than in G0/G1 phase (Aberg, 2013; Kim, 2011). The cell cycle slowdown and cell accumulation in G1 phase thus supports the hypothesis of reduced intracellular accumulation rather than expulsion of TiO2-NP from chronically-exposed cells. Taken together, all these data can be summarized as follows: Chronic exposure to TiO2-NPs leads to their accumulation in cell cytoplasmic vesicles. Cells react to these exogenous compounds by activating autophagy, which fails to eliminate TiO2-NPs. Autophagosomes then accumulate in cells, but no longer fulfil their role and do not eliminate altered mitochondria. Altered mitochondria accumulate in the cell cytoplasm, leading to increased release of ROS which induces oxidatively-generated DNA lesions. These lesions accumulate, together with strand breaks due to excision/resynthesis DNA repair processes and the replication fork blockade that they cause. Moreover the p53 pathway is activated by this mild stress, and leads to cell cycle slowdown and consequently decreased cell proliferation and decreased intracellular accumulation of TiO2-NPs. JUST ACCEPTED Another important issue highlighted in this study is that cells chronically exposed to TiO2-NPs for 2 months are sensitized to the alkylating agent MMS, which is classically used as a positive control in genotoxicity experiments (Beranek, 1990). Indeed we observe MMS cytotoxicity when cells chronically exposed to TiO2-NPs are then incubated with a sublethal dose of this alkylating agent, as well as increased levels of DNA strand breaks compared to the chronic controls (not exposed to MMS). This sensitization might be explained by two hypotheses. First, the DNA damage that we observed in our model can enhance the MMS-induced DNA damage to saturate DNA repair activity, leading to cell death. Second, TiO2-NPs themselves can also impair cellular DNA repair activity, as we previously demonstrated upon acute exposure (Jugan, 2012). This observation underlines the need to deeply investigate the combined effects of NPs with other pollutants, particularly genotoxic compounds and alkylating agents. Finally, our results show more drastic impacts from P25 TiO2-NPs upon chronic exposure of A549 cells, particularly after 4 weeks of exposure, than those previously reported in the study of Vales et al. (Vales, 2014). This discrepancy may be due to the different cell lines that we used, BEAS-2B being bronchial cells while A549 are alveolar cells. Indeed alveolar cells, particularly A549 cells, produce and secrete pulmonary surfactant (Lieber, 1976), which changes the biochemical environment of the milieu around these cells and may be adsorbed on the surface of NPs, changing their properties. Another explanation lies in the fact that different NPs were used in these two studies: we used P25 NPs, which are mixed anatase-rutile crystal phase and which generate a highly stable suspension in water and in cell culture medium. Conversely, Vales et al. (2014) used NMP102 TiO2-NPs which are pure anatase and very hard to disperse in aqueous medium. For their experiments, these NPs were dispersed in BSA before being diluted in cell culture medium. This probably leads to the formation of a dense protein corona on the surface of these NPs. This corona may be different from the one that forms on the NPs used in our study, which were first dispersed in water then diluted in FBS-containing cell culture medium then exposed to the surfactant secreted by A549 cells. Since the protein corona on the surface of NPs plays a role in their cellular accumulation and impact (Monopoli, 2012), and since direct comparison shows that TiO2-NPs dispersed either in BSA or in water lead to non-genotoxic or genotoxic impact, respectively (Magdolenova, 2012), these different NPs and dispersion protocols may explain the different toxicological outcomes observed. JUST ACCEPTED We demonstrate here that long-term exposure to TiO2-NPs (AEROXIDE® P25, Evonik; received as NM105 from the JRC) induces DNA damage in A549 cells and sensitizes these cells, as evidenced by the increased amount of DNA strand breaks induced by the alkylating agent MMS. Although this longterm exposure model is still far from the reality of human inhalation exposure, some genotoxicity endpoints appear positive even at very low exposure concentrations (as low as 1 μg/mL). Chronic exposure causes more severe DNA damage than acute exposure, particularly those revealed by 53BP1 foci immunostaining, i.e. double-strand breaks or replication fork blockade. This underlines the necessity to use chronic exposure scenarios in addition to acute exposure models in order to investigate NP impact, since these distinct models lead to different adverse outcomes, with additional triggering events and more intense effects in long-term exposure scenarios. This work was funded by CEA through the Nanoscience and Toxicology research programs, and via the European Commission's 7th Framework Programme project NanoMILE (Contract No. NMP4-LA-2013-310451). It is a contribution to the Labex Serenade (n° ANR-11-LABX-0064) funded by the \"Investissements d'Avenir\" French Government program of the French National Research Agency (ANR) through the AMIDEX project (n° ANR-11-IDEX-0001-02). This research received support from the QualityNano Project http://www.qualitynano.eu which is financed by the European Community Research Infrastructures under the FP7 Capacities Programme (Grant No. INFRA-2010-262163), and its partner University of Birmingham. The authors would like to thank Veronique Collin-Faure for flow cytometry analyses. JUST ACCEPTED The authors declare that there are none. Figure 1. NP intracellular accumulation measured by μPIXE. Distribution maps of calcium and potassium (K/Ca, used as indicator of cell outlines) and titanium (Ti) together with their colour scale (counts): the hotter the color, the higher the concentration (A). Cells were analyzed after exposure to 0, 2.5 and 50 μg/mL NPs for 1 and 2 months. PIXE spectra displaying the regions of interest selected for mapping and quantification of Ti content in cells (B). Titanium concentration (ng/cm2) obtained by integration of PIXE spectra recorded on unexposed cells and cells exposed at 2.5 or 50 μg/mL for either 24 h, 1 month or 2 months (C). Results are presented as the mean of 8 points ± SEM. : p<0.05 vs control cells; #: p<0.05 vs 2.5 μg/mL NP exposed cells. Figure 2. TEM images of A549 cells exposed to TiO2-NPs. (A) Control cells (unexposed cells); (B) Cells exposed to 2.5 μg/mL NPs for 2 months; (C) and (D) Cells exposed to 50 μg/mL NP for 2 months. Figure 3. Cell viability after NP exposure. (A) Cellular metabolic activity, represented by the MTT assay; (B) cell viability, evaluated by propidium iodide staining; (C) Population doubling level (PDL), indicative of cell proliferation. Results are presented as the mean of 4 points ± SEM. : p<0.05 vs control. JUST ACCEPTED Figure 4. Intracellular ROS content after TiO2-NP exposure. DCF fluorescence was normalized with respect to protein content, reflecting cell number. The obtained data were then normalized to untreated control. Value for positive control: H2O2 50 μM, 30 min: 2.1±0.3. Results are presented as the mean of 4 points ± SEM. : p<0.05 vs control; #: p<0.05 vs lower dose for similar exposure period. Figure 5: TiO2-NP genotoxicity. (A) Alkaline comet assay, assessing single and double strand breaks as well as alkali-labile sites. Value for positive control: 50 μM H2O2 deposited on the slide 5 min before migration: 61.9%; (B) Fpg-modified comet assay, assessing single and double strand breaks, alkalilabile sites and oxidized bases. Value for positive control: riboflavin/UVA: 22.6%. Results are presented as the mean of % DNA in comet tail ± SEM (n=4). : p<0.05 vs control; #: p<0.05 vs alkaline comet assay equivalent condition; $: p<0.05 vs lower dose for equivalent exposure period. (C) 53BP1 foci counting, assessing double strand DNA breaks. Total 53BP1 foci per image was reported relative to total cell number, and normalized to control data. Results are presented as the mean of 15 images ± SEM. : p<0.05 vs control (unexposed cells). Value for controls: unexposed cells 0.27%, 25 μM etoposide: 1.95%. Figure 6: Cell sensitization to MMS due to long-term exposure to TiO2-NPs. (A) Viability and (B) DNA strand breaks in cells exposed to NPs for 2 months and then to 100 μM MMS for 24h were assessed by MTT and alkaline comet assay, respectively. (A) Data were normalized to the results obtained in control cells, i.e. cells that were not exposed to MMS. Results are presented as the average of 6 points ± SEM. : p<0.05 vs control (unexposed cells). (B) Results from alkaline comet assay, from cells exposed to TiO2-NPs for 2 months (white bars) or to TiO2-NPs for two months, then to 100 μM MMS for 24 h (grey bars). Data are expressed as % tail DNA and are the average of 3 comet slides (50 comets per slides) ± SEM. *: p<0.05 vs control (unexposed cells); the % indicates % increase as compared to control (unexposed) cells. Value for positive control: 50 μM H2O2 deposited on the slide 5 min before migration: 64.3%. JUST ACCEPTED for the assessment of nanoparticle toxicity in the lung-A dialog between aerosol science and biology. Journal of Aerosol Science 42(10): 668-692. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 342.0, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 1.0, \"viab\": 104.88}, {\"h\": 24.0, \"mgL\": 2.5, \"viab\": 101.8}, {\"h\": 24.0, \"mgL\": 5.0, \"viab\": 101.8}, {\"h\": 24.0, \"mgL\": 10.0, \"viab\": 105.66}, {\"h\": 24.0, \"mgL\": 50.0, \"viab\": 91.517}]}", "role": "assistant" } ]	78.json
[ { "content": "Extract sample context and measurement list. <doc> RESEARCH PAPER Zi-jin Zhang & Zhi-jie Tang & Zhen-yu Zhu & Zhao-ming Cao & Hong-juan Chen & Wei-juan Zheng & Xin Hu & Hong-zhen Lian & Li Mao Received: 5 April 2018 /Accepted: 17 August 2018 # Springer Nature B.V. 2018 Abstract Zinc oxide nanoparticle is one of the nanomaterials people engaged most in their life and its health effect has been taken into concern. In this work, A549 cell line was used as cell model, and the cytotoxicity of zinc oxide nanoparticles was revealed to be concentration-dependent. Through the measurement of cellular proteome, much more differentially expressed proteins were observed after the cells being treated for 9 h than 24 h. Also, most of these proteins expressed in the pattern which showed a significant decrease after exposure to zinc oxide nanoparticles and then an increase at 24 h. Intracellular reactive oxygen species and glutathione determination indicated that high level of oxidative stress was presented in cell after treatment Electronic supplementary material The online version of this article (https://doi.org/-018-4333-0) contains supplementary material, which is available to authorized users. Z.<j. Zhang : Z.<j. Tang : Z.<m. Cao : X. Hu : H.<z. Lian () State Key Laboratory of Analytical Chemistry for Life Science, Collaborative Innovation Center of Chemistry for Life Sciences, School of Chemistry and Chemical Engineering and Center of Materials Analysis, Nanjing University, Nanjing 210023, China e-mail: State Key Laboratory of Pharmaceutical Biotechnology, School of Life Sciences, Nanjing University, Nanjing 210023, China Ministry of Education (MOE) Key Laboratory of Modern Toxicology, School of Public Health, Nanjing Medical University, Nanjing 211166, China e-mail: with zinc oxide nanoparticles for 9 h. It can be observed from western blot analysis that the expression of NF-κB p65, PNPase, and HSP90 rose significantly after 9 h of exposure. Thus, a deduction was reached that toxicity of nanoparticles consists both of particle toxicity and ion toxicity, and a long-time treatment may conceal the toxicity induced by particles. The conclusion we made highlighted the importance of exposure time in the study of nanoparticle toxicity and would provide a new perspective for studying toxicity mechanism of nanoparticles. Proteomics.Toxicity.Time-dependent response .Health effects Recent innovations in mass production of nanoparticles have led to rapid commercialization of these kinds of materials with characteristic length less than 100 nm (Meyer, et al. 2009). As a result, nanoparticles are now widely used in industrial and commercial fields. Due to the rising application of nanoparticles, their health and environment effect has been taken into concern. A large number of studies have been taken to evaluate the toxicity of nanomaterials both in vivo and in vitro. It was reported that ultrafine carbon black induced lung injury in mice (Chang, et al. 2007). Titanium oxide nanoparticles were found to cause cell death in BEAS-2B cells (Park, et al. 2008) and reduced cell viability was showed in silica nanoparticle-treated A549 cells (Lin, et al. 2006). Zinc oxide nanoparticles are one kind of most concerned nanomaterials among them since its application in commodities such as paints, cosmetics, batteries, sunscreens, and electronic devices (The Project on Emerging Nanotechnologies Consumer Products Inventory, http://www.nanotechproject.org/inventories/consumer/). However, a great number of studies have shown its toxicity in vitro (Zhong, et al. 2017). For instance, LDH leakage and a great reduction in cell viability were caused by ZnO nanoparticles while using Neuro-2A mouse neuroblastoma cell line as cell model (Jeng and Swanson, 2006); ZnO nanoparticle-induced cytotoxicity was also observed in THP1 human monocytic cell line (Prach, et al. 2013); Primary human nasal mucosa cells showed DNA damage and inflammation after treatment with ZnO nanoparticles (Hackenberg, et al. 2011). On the other hand, few researchers have used proteomic method, a powerful tool for us to have an overall view of the effect induced by zinc oxide nanoparticles to study the toxicity mechanism of zinc oxide nanoparticles. Triboulet et al. used two-dimensional gel electrophoresis to investigate the proteome changes in RAW264 macrophage. They found a rather weak response in oxidative stress response pathway but strong responses both in central metabolism and in proteasome protein degradation pathway, while carbohydrate catabolism and proteasome are critical in response to zinc ions. By contrast, glutathione level and phagocytosis did not show significant changes at moderately toxic zinc concentration. It is concluded that zinc ions bound proteins and induced impairment of central metabolism and thus mainly contributed to the zinc oxide nanoparticle toxicity, and proteasome plays an important role in zinc resistance (Triboulet, et al. 2014). Considering the various results coming out before, our work chose A549 cells as subject, profiled the proteome after different period of treatment with zinc oxide nanoparticles, and analyzed the intracellular ROS level in order to verify the result. We aimed to study the toxicity mechanism of zinc oxide nanoparticles and tried to explain the divergence in previous works. Zinc oxide nanoparticles were purchased from Sigma-Aldrich (catalog 544906). Nanoparticles were dispersed in water as a suspension by sonication for 30 min. The concentration was determined after digestion with concentrated hydrochloric acid using an inductively coupled plasma atomic emission spectroscopy (ICP-AES, PerkinElmer SCIEX, Optima 5300). Elemental composition of zinc oxide nanoparticle was determined by an energy dispersive X-ray spectroscopy (EDX, HORIBA, EX250). The morphology of nanoparticles was observed by a scanning electron microscopy (SEM, Hitachi, S-3400N II) and a transmission electron microscopy (TEM, JEOL, JEM-200CX). The hydrodynamic diameter (Dh) of zinc oxide nanoparticles was determined by dynamic light scattering (DLS) experiment using a particle size analyzer (BIC, NanoBrook 90 Plus PALS). Cell culture and treatment with zinc oxide nanoparticles Human lung adenocarcinoma cell line A549 was purchased from Cobioer Biotech (Nanjing, China) and were maintained in the complete cell culture medium F12-K supplemented with 10% fetal bovine serum, 100 units/L penicillin, and 0.10 mg/mL streptomycin at 37 °C in a humidified 5% carbon dioxide (CO2) atmosphere. All cell samples were prepared using at least three replicates for an experiment. A stock solution of up to 1.22 g Zn/L was prepared and dispersed for 30 min by a sonicator to prevent aggregation, and diluted to the specified concentrations (1, 2.5, 5, 10, 25, 50 mg Zn/L) for treatment of cells. Cellular viability was determined using the CCK-8 assay and Annexin V-FITC/PI staining (Beyotime). Cells were seeded with equal density in each well of 96-well plates (5×103 cells per well), 100 μL of cell culture medium per well, and incubated for 24 h at 37 °C. Then, cells were treated in 96-well plates with 1 mg Zn/L, 2.5 mg Zn/L, 5 mg Zn/L, 10 mg Zn/L, 25 mg Zn/L, or 50 mg Zn/L of zinc oxide nanoparticles for 24 h at 37 °C. The medium was renewed with fresh culture medium before assay and 10 μL CCK-8 solution was added to each well, and the plate was incubated for 30 min at 37 °C. Subsequently, the absorbance was measured at 450 nm using a Bio-Rad 680 enzyme micro-plate reader. Cell viability was calculated by comparing absorbance value of cells treated with nanoparticles and untreated cells, and the cell viability of untreated cells was as 100%. Another group of nanoparticletreated cells were collected and resuspended using PBS. 105 cells were taken and centrifuged. The supernatants were discarded and 195 μL Annexin V buffer, 5 μL Annexin-FITC, and 10 μL PI staining solution were added to each sample and mixed. After being incubated for 20 min in a dark place at room temperature, the samples were measured on a flow cytometer (Becton Dickinson, FACSCalibur). Preparation of protein samples Cell lysis was applied using a cell lysis kit (Beyotime, Haimen, China). Briefly, cells were washed with icecold PBS after removing the cell culture medium. Then, 1 mL cell lysis buffer (20 mM Tris (pH 7.5), 150 mM NaCl, 1% Trition X-100, Inhibitor (sodium pyrophosphate, β-glycerophosphate, EDTA, Na3VO4, and leupeptin)) was added and cells were harvested after 1 min. Lysates were sonicated for 1 min at a 10% amplitude at interval settings of 2 s on and 3 s off with a sonicator. The heat generated by sonication was below 1000 J. The supernatants were clarified and recovered after centrifugation at 15000×g for 30 min at 4 °C. Protein samples were flash-frozen and stored at − 80 °C. The concentration of the protein extracts was determined using the BCA method. An aliquot (200 μg) of acetone-precipitated protein sample was reduced with 4 μL of reducing reagent (AB Sciex, Redwood City, CA, USA) at 60 °C for 1 h. 2 μL of cysteine-blocking reagent (AB Sciex, Redwood City, CA, USA) was then added. After reaction for 10 min, samples were centrifuged at 12000 rpm for 20 min and bottom solution was disposed. Protein for each sample was digested with trypsin overnight at 37 °C in a 1:50 trypsin-to-protein mass ratio. iTRAQ Reagent was then added and samples were dried by vacuum centrifugation after incubation for 2 h. The proteins were labeled with iTRAQ tags as follows: control group, Tag113; 9-h group, Tag114; 24-h group, Tag115. Labeled peptides were resuspended with loading buffer (20 mM ammonium formate, pH 10) and loaded onto a Durashell C18 column (4.6 mm × 150 mm, particle size 5 μm) at 0.8 ml/min. Peptides were eluted from the analytical column after a 65-min reverse phase solvent gradient from 5% B (A: 20 mM ammonium formate, B: 20 mM ammonium formate, 80% acetonitrile) to 90% B. Peptides were then resuspended and loaded onto a TripleTOF TM 5600 mass spectrometer (AB Sciex, Redwood City, CA, USA). TOF MS was performed under the following condition: m/z: 350–1250, accumulation time: 0.25 s. Raw data of iTRAQ-labeled proteins were processed for database searching using Mascot (Matrix Science, London, UK). The search parameters were set as follows: cysteine alkylation with IAM; trypsin digestion; maximum of two missed cleavage sites; 0.02 Da fragment mass tolerance. A decoy database search strategy was used to determine the false discovery rate (FDR) for peptide and protein identification. Peptide identifications were accepted if their FDR value was < 1.0%, while protein identifications contained at least 1 identified peptide. Protein quantification required a p value < 0.05, only fold-change ratios > 1.500 or < 0.667 were considered statistically significant. GO annotations for the identified proteins were assigned according to those reported in the UniProt database (http://www.uniprot. org). The differentially expressed proteins that we identified were categorized according to the DAVID Classification System (https://david.ncifcrf.gov/). Pathway analyses of identified proteins were performed using the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (http://www.genome. jp/kegg/). GO terms and KEGG pathways with corrected p value < 0.05 were considered significant. Thirty-five micrograms of proteins in whole cell extracts were fractionated on 12% acrylamide gels by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) according to Laemmli's method (Laemmli, 1970 ). Proteins were then electrotransferred on to PVDF membranes using a Mini P-4 electrotransfer apparatus (Cavoy, Beijing, China). PVDF membranes were activated by soaking in methanol for 1 min prior to blotting. The membranes were then equilibrated for 30 min in blotting buffer [48 mM Tris-base, 39 mM glycine, 20% (v/v) methanol, and 0.0375 (w/v) SDS]. A Bblotting sandwich^ was made according to the manufacturer's instructions. Blotting was carried out for 1 h on ice at a constant voltage of 300 mA. After transfer, the membrane was blocked in western blocking buffer for 1–2 h at room temperature. After three washing steps with PBST (PBS with Tween-20) for 10 min, the membrane was incubated with primary antibody overnight at 4 °C. The primary antibodies were diluted in primary antibody dilution buffer as ratio of 1:500 according to manufacturer's specifications. Then, the membrane was washed three times with PBST for 10 min and incubated for 1 h at room temperature in the presence of the appropriate horseradish peroxidase-conjugated secondary antibody. After several washes, the membrane was incubated with Tanon High-sig ECL Western Blotting Substrate (Tanon, Shanghai, China) and immune complexes were detected using the Tanon 6200 chemiluminescence workstation (Tanon, Shanghai, China). The intracellular ROS was determined using a wellcharacterized probe, 2′,7′-dichlorofluorescin diacetate (DCFH-DA). DCFH-DA passively enters the cell, and is hydrolyzed by esterases to DCFH. This nonfluorescent molecule is then oxidized to fluorescent compound dichlorofluorescein (DCF) by cellular oxidants. A DCFH-DA stock solution of 10 mM was diluted 1000 fold in the cell culture medium without serum or other additives to yield a 10 μM working solution. Cells were washed three times with PBS and then incubated with 1 mL DCFH-DA working solution for 30 min in the 37 °C incubator. Then, the cells were washed three times with cell culture medium without serum to eliminate DCFH-DA that did not enter the cells. Cells were harvested and the fluorescence was determined at 488 nm excitation and 525 nm emission using a fluorospectrophotometer (Shimadzu, RF-5301PC). Total glutathione and oxidized glutathione measurement Intracellular total glutathione and oxidized glutathione (GSSG) was detected using a total glutathione and oxidized glutathione assay kit (Beyotime, Haimen, China). Glutathione (GSH) can reduce 5,5′-dithiobis-(2-nitrobenzoic acid) (DTNB) to yellow 5-thio-2-nitrobenzoic acid (TNB) and be oxidized to GSSG, and addition of glutathione reductase can reduce GSSG to GSH, so that the reaction rate was limited to the amount of total glutathione (GSH + GSSG), which thus can be detected according to the absorbance. In addition, presence of tris(2-hydroxyethyl) amine and 2-vinylpyridine can abolish GSH and, as a result, the amount of GSSG can be observed. Briefly, cells were washed with PBS and then collected. Supernatants were discarded after centrifugation and 30 μL protein eliminating solution was added. The samples were then flash-frozen to − 80 °C and melt under 37 °C water bath, twice, and put in the 4 °C environment for 5 min. The samples were further centrifuged at 10000×g for 10 min at 4 °C, and the supernatants were collected. Subsequently, the absorbance was measured at 412 nm. Intracellular total glutathione and oxidized glutathione was calculated by comparing absorbance value of cells treated with nanoparticles and untreated cells. All measurements were repeated at least three times and data were expressed as mean ± SEM. Statistical significance for comparison of two groups was assessed using one-way ANOVA with the Tukey–Kramer multiple comparison post hoc test. Differences that were considered to be statistically significant are indicated as follows: , p < 0.05 vs. untreated controls. SEM and TEM micrographs showed that zinc oxide nanoparticles were spherical and had a diameter about 30 nm (Fig. 1). The EDX analysis (Fig. S1 and Table S1, see Supplementary Material) showed that the major elemental composition (> 98%) of zinc oxide nanoparticles was zinc and oxygen with minor amount of aluminum. The DLS analysis (Table S2 and S3, see Supplementary Material) showed that zinc oxide nanoparticles are relatively stable in culture medium (F12K + 10% FBS). In order to verify that zinc oxide nanoparticles had not dissolved before they contacted the cells, the supernatants of stock solution (1.22 g Zn/L) were measured after being stored for 7 days and the concentration of zinc ions was only 7.58 mg/L. Cytotoxicity induced by zinc oxide nanoparticles was estimated as shown in Figs. 2 and 3. Treatment with zinc oxide nanoparticles at the concentration below 5 mg Zn/ L for 24 h failed to induce significant reduction of A549 cell viability and less than 5% cell death was observed. However, 10 mg Zn/L zinc oxide nanoparticles significantly decreased cell viability. And as the concentration of nanoparticles increased, the cell viability dropped continuously. The cell viability was about 60% at the dosage of 25 mg Zn/L and nearly half of cell death could be observed at the dosage of 50 mg Zn/L (Fig. 2). According to the result of flow cytometry, 96.5% cells were viable after being treated with 5 mg Zn/L for 24 h while only 76% cells were viable when the concentration rose to 25 mg Zn/L (Fig. 3). Accordingly, we chose 5 mg Zn/L zinc oxide nanoparticles, which is considered to be sub-cytotoxic and can reduce the cell impairment to a minimum in order to analyze the protein regulation for further research of toxic mechanism in the following experiments. Proteome expression patterns in response to zinc oxide nanoparticles iTRAQ assay was conducted in order to further evaluate the effect of zinc oxide nanoparticles on proteome of A549 cells. Five milligrams zinc per liter of zinc oxide nanoparticles was used to treat cells for 9 h and 24 h according to our previous work which revealed that the response to sub-cytotoxic zinc ions of A549 cells reached its maximum at about 8–10 h. On the other hand, this can help us compare the proteome differences induced by zinc oxide nanoparticles and zinc ions and further study the toxicity mechanism of zinc oxide nanoparticles (Zhao, et al. 2014; Zhao, et al. 2015). Four thousand thirty-two proteins were identified through the assay, in which 120 were differentially expressed proteins (p < 0.05, fold-change > 1.500 or < 0.667). Among them, 99 were presented in the 9-h group, in which 12 (12.1%) were up-regulated while rest 87 (87.9%) were down-regulated; 41 were presented in the 24-h group and 9 (22.0%) of them were up-regulated while rest 32 (78.0%) were down-regulated. Twenty proteins were identified differentially expressed both after 9 h and 24 h, in which 3 were up-regulated proteins that showed higher expression after 24 h of treatment than 9 h, while 2 proteins (11.8%) among the rest 17 down-regulated proteins showed lower expression after 24 h of treatment compared to 9 h, and the rest 15 proteins (88.2%) showed an up-regulation that took place at 24 h compared to 9 h, though still down-regulated compared to the control group. Detailed differentially expressed proteins are shown in the Supplementary Material, Table S4 and S5, and a clear view of the change of protein expression is showed in Fig. S2. Functional classification of differentially expressed proteins Multiple function analyses of identified differentially expressed proteins were performed. The differentially expressed proteins that we identified were categorized according to the DAVID Classification System (https://david.ncifcrf.gov/) and a great number of biological processes were changed significantly after treatment with zinc oxide nanoparticles, including cellcell adhesion (GO:0098609), glycolytic process (GO:0006096), gluconeogenesis (GO:0006094), rRNA processing (GO:0006364), response to unfolded protein (GO:0006986), cell redox homeostasis (GO:0045454), and many processes directly engaged in protein synthesis, for example, translation (GO:0006412), protein folding (GO:0006457), etc. Fig. S3 shows some main biological processes involved in response to zinc oxide nanoparticles. In addition, molecular functions of differentially expressed proteins were also studied, including protein binding (GO:0005515), RNA binding (GO:0003723), oxidoreductase activity, acting on the CH-OH group of donors, NAD or NADP as acceptor (GO:0016616), ion channel binding (GO:0044325), kinase binding (GO:0019900), etc. Detailed information is shown in Supplementary Material, Table S6-S9. Besides, KEGG System (http://www.genome.jp/kegg/) was used to investigate the pathway these differentially expressed proteins participated in, which includes glycolysis/gluconeogenesis (hsa00010), pyruvate metabolism (hsa00620), RNA degradation (hsa03018), etc. Detailed lists of pathways can be found in the Supplementary Material (Table S10 and S11). Expression of HSP90, NF-κB p65, and PNPase The expression of heat shock protein 90, nuclear factorkappa B, and purine nucleoside phosphorylase were observed after A549 cells were treated with zinc oxide nanoparticles for 9 h and 24 h. The relative abundance of proteins treated with nanoparticles compared to the internal reference protein beta-actin was analyzed and normalized. These differential expression patterns are shown in Fig. 4. Exposure to zinc oxide nanoparticles Fig. 4 Expression of NF-κB p65, HSP90, and PNPase at protein level produced a peak value of Hsp90 after 9 h, which was about a twofold increase, and then decreased to the level of control group after 24 h of treatment. Statistical analysis revealed significant differences in abundance of Hsp90 after treatment for 9 h. The expression of NF-κB p65 and PNPase showed a similar pattern. It increased about 40% after treatment for 9 h with significant differences, and then dropped significantly to about only 60% of the control group level. Two proteins both showed a response to zinc oxide nanoparticle treatment in a time-dependent manner. Intracellular ROS levels increased about 10% after A549 cells being treated with zinc oxide nanoparticles for 9 h, though no significant differences were discovered (Fig. 5). After 24 h of treatment, the intracellular ROS level fell back, slightly below the untreated level. Intracellular total glutathione/oxidized glutathione level was investigated in order to further verify the oxidative stress in A549 cells. As shown in Fig. 6, cellular total glutathione level maintained stable in the 24-h period, while oxidized glutathione increased significantly after cells being treated by zinc oxide nanoparticles for 9 h and then fell back at 24 h, which means the glutathione decreased after 9 h of treatment and then increased to control group level after 24 h. A549 cell viability shows responses to treatment with zinc oxide nanoparticles in a concentration-dependent manner, and the nanoparticles show cytotoxicity similar to zinc sulfate according to our previous work (Zhao, et al. 2015), in which zinc sulfate at the concentration of 150 μM (corresponding to 9.8 mg Zn/L) caused significant cell viability reduction after treatment for 24 h, while 10 mg Zn/L zinc oxide nanoparticles also induced significant cytotoxicity. Both 100 μM zinc sulfate (corresponding to 6.5 mg Zn/L) and 5 mg Zn/L zinc oxide nanoparticles caused minor cell viability reduction in A549 cells after 24 h. In addition, differences in cytotoxicity were also observed. Five hundred micromolars of zinc sulfate (corresponding to 32.7 mg Zn/L) treatment resulted in nearly 80% cell death, while 50 mg Zn/L zinc oxide nanoparticles only caused about 40% cell viability loss, which indicated that zinc ions induced higher cytotoxicity than zinc oxide nanoparticles at high concentration. Besides, zinc oxide nanoparticles dissolved incompletely after 24 h, and this ensured the basis of differences in toxicity mechanism. On the other hand, cytotoxicity induced by 25 mg Zn/L and 50 mg Zn/L zinc oxide nanoparticles was similar. In such high concentration, nanoparticle aggregation is unavoidable unless aggregation inhibitor is involved, which means that the specific surface area will decrease and cause less oxidative stress to cells and thus less cell viability loss. According to the result, 5 mg Zn/L zinc oxide nanoparticles were chosen for the following experiments since it is sub-cytotoxic and can further reveal cell homeostasis regulation in response to exogenous stimulation. Also, 9 h and 24 h were chosen so that we can make direct comparison between toxicity of zinc oxide nanoparticles and zinc sulfate. One hundred twenty differentially expressed proteins were detected using iTRAQ assay and showed timedependent expression patterns. Most (79) of 99 differentially expressed proteins after treatment with zinc oxide nanoparticles for 9 h were not detected to be differentially expressed after 24 h, which means that cells have managed to maintain homeostasis after treatment for 24 h. Among 20 proteins which showed to be differentially expressed both in 9 h and 24 h, 15 proteins were down-regulated in 9 h but the regulation was then suppressed, and the suppression was supposed to continue, which led to the expression level same to the untreated group after longer period of time. This result correlates with our previous conclusion that protein expression changed largest after 9 h of treatment. Moreover, much more differentially expressed proteins were detected at 9 h than 24 h which also supports the conclusion and further reveals the importance of treatment time in mechanism of cellular response to zinc oxide nanoparticles. In addition, most of differentially expressed proteins were found to be down-regulated, and down-regulated proteins also outnumbered up-regulated proteins among 20 differentially expressed proteins after both 9 h and 24 h. By contrast, most of differentially expressed proteins in response to zinc sulfate were up-regulated (Zhao, et al. 2015), which showed a different toxicity mechanism. Also, differentially expressed proteins were involved in different biological processes to some extent, which is evidence that nanoparticle is crucial in the cytotoxicity even at low concentration. ROS plays an important role in nanoparticle toxicity (Nel, et al. 2006). In order to evaluate the cytotoxicity originating from ROS, DCFH-DA fluorescent probes were used to determine the intracellular ROS level after A549 cells being treated with zinc oxide nanoparticles for 9 h and 24 h. It is shown that the ROS level increased about 10% after treatment for 9 h though no significant difference could be found, and then decreased to the level similar to control group. We further investigate the level of intracellular total glutathione and oxidized glutathione. Total glutathione level maintained stable in the 24-h treatment period while oxidized glutathione level increased significantly after treatment for 9 h and then fell back after 24 h. These results indicated that ROS were generated after cells being treated with zinc oxide nanoparticles and cell would respond to the oxidative stress, and the ROS level was suppressed to normal level after 24 h. These results corresponded to our proteomic results, and it is clear that cells sustained the homeostasis and the exogenous interference was suppressed after 24 h treatment, which is reasonable since the concentration chosen was sub-cytotoxic and a totally different outcome can be predicted if higher concentration was taken in the treatment. Three proteins involved in related cell signaling pathways, NF-κB p65, PNPase, and HSP90 expression, were determined using western blot and they showed similar expression pattern, which corresponded to the conclusion that most proteins showed the greatest changes after 9 h of treatment. NF-κB is a multifunction transcription factor and presents in almost all types of cells. It participates in various biological processes including inflammation, immunity, apoptosis, proliferation, differentiation, and tumorigenesis and plays an important role in innate and acquired immune responses. NF-κB is a protein family, in which p65 is one of highest abundant proteins. Normally, p65 presents as a protein complex in cytoplasm and combined with NF-kappa-B inhibitor (IκB) family. After receiving an immune signal like TNF, IL-1, LPS, or other stimuli like virus and ultraviolet radiation, IκB kinase will promote the phosphorylation of IκB and cause it to degrade, and thus activate the NF-κB, which will transfer into cell nuclei and initiate the transcription process (Baldwin, 1996; Okamoto, et al. 1997; Karin and Ben-Neriah, 2000; Ghosh and Karin, 2002). Nanoparticles have been proved to induce the generation of ROS directly and ROS can activate various immunoreactions and lead to the activation of NF-κB p65 (Nel, et al. 2006; Yang, et al. 2013; Fooksman, et al. 2010). The main function of PNPase is to catalyze the phosphorolysis of purine nucleosides and deoxynucleosides to their respective purine bases and pentose-1-phosphates. Lack of PNPase will cause deficiency in purine catabolism and lead to immunological diseases. On the other hand, PNPase was also proved to be induced by ROS (Ealick, et al. 1990; Rao, et al. 1990). The significant increase of NF-κB p65 and PNPase indicates that the nanoparticle accounts for an important part in the toxicity of zinc oxide nanoparticles. HSP90 protein family shows a high-level expression in all types of cells. It is a chaperone protein that assists other proteins to fold properly and aids in degradation of misfolded proteins, so it is synthesized to deal with denatured proteins when cells encounter exogenous stimuli like high temperature, anoxia, and heavy metal (Csermely, et al. 1998). HSP90 increased significantly after cells being treated with 5 mg Zn/L zinc oxide nanoparticles for 9 h, and then declined to control group level after 24 h. However, when facing zinc sulfate stimuli in our previous work, HSP90α increased significantly after treatment for both 9 h and 24 h, and the expression at 24 h was even higher than 9 h. Considering HSP90 family responds to most of stimuli, combined with other results in our present work, we deducted that zinc oxide nanoparticles tend to have an Bacute^ effect on A549 cells while zinc ions will have a long-lasting effect on cells. This may be because more time is needed to regulate the concentration of intracellular zinc ions, while oxidative stress induced by ROS, which is originated from particles, can be abolished in a short period of time through regulation. This deduction reveals the importance of controlling the time of treatment in the study of metal oxide nanoparticle toxicity mechanism. A long-time treatment may lead to the conclusion that metal ion-induced toxicity is the main contributor to metal oxide nanoparticle toxicity since particle-induced toxicity, which largely comes from ROS, can be abolished rapidly. The dissolving nanoparticles will cause the reduction of nanoparticle toxicity and the increase of metal ion toxicity, and vice versa. However, if cells fail to eliminate the excess ROS due to various reasons, for instance, being treated with metal oxide nanoparticles at high concentration, accumulated ROS may emphasize the toxicity induced by particle. The deduction can explain zinc oxide nanoparticle toxicity mechanism in this work properly. Proteomes of A549 cells in response to zinc oxide nanoparticles stimulation after different period of time were profiled and the differentially expressed proteins are mainly involved in biological processes like transcription translation and protein folding. Although similar cytotoxicity was observed at low concentration, differences between cells in response to zinc oxide nanoparticles and zinc sulfate were investigated through cell viability assay and protein expression analysis. In addition, more differentially expressed proteins, more significant changes, and higher ROS level after treatment for 9 h than 24 h reveal the process of cell regulation in homeostasis maintenance. We further make a deduction that toxicity of nanoparticles consists both of particle toxicity and ion toxicity. When at low concentration, cells will suffer from particle and metal ion toxicity simultaneously for a short period of time, and then particle-induced toxicity will soon be relieved or eliminated through regulation, while dissolved metal ions may cost a long time to regulate. As a result, long-time treatment may conceal the toxicity induced by particles and in turn highlight the toxicity of metal ions. Still, some other causes which generate or suppress the elimination of ROS will lead to the ion toxicity and should be taken into consideration. This hypothesis tries to explain the controversies in explanations of zinc oxide nanoparticle toxicity mechanism and to reveal the significance of the exposure time in the study of metal oxide nanoparticle toxicity, which may provide a new perspective for studying toxicity mechanism of nanoparticles. Funding This work was supported by the National Natural Science Foundation of China (91643105, 21577057, 91543129, 81072712, and 90913012), the Natural Science Foundation of Jiangsu Province (BK20171335), and the National Basic Research Program of China (973 program, 2011CB911003). We thank Mr. LY Huang for assistance in the ICP-MS determination of zinc. Conflict of interest The authors declare that they have no conflict of interest. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 54.17, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 1.0, \"viab\": 96.943}]}", "role": "assistant" } ]	65.json
[ { "content": "Extract sample context and measurement list. <doc> Available online at www.sciencedirect.com CERAMICS INTERNATIONAL Ceramics International 39 (2013) 6917–6921 www.elsevier.com/locate/ceramint Majid Darroudia,n , Mohammad Hakimib , Mina Saranib,c , Reza Kazemi Oskueea,d , Ali Khorsand Zake , Leila Gholamia a Department of Modern Sciences and Technologies, School of Medicine, Mashhad University of Medical Sciences, Mashhad, Iran b Chemistry Department, Payame Noor University, 19395-4697 Tehran, Iran c Zabol Medicinal Plants Research Center, Zabol University of Medical Sciences, Zabol, Iran d Targeted Drug Delivery Research Center, Mashhad University of Medical Sciences, Mashhad, Iran Electroceramics and Materials Laboratory, Physics Department, Faculty of Science, Ferdowsi University of Mashhad, Mashhad, Iran Received 27 January 2013; received in revised form 5 February 2013; accepted 11 February 2013 Available online 17 February 2013 e Cerium oxide nanoparticles (CeO2-NPs) were synthesized via the sol–gel method in gelatin media. Long-chain gelatin compounds were utilized to terminate the growth of CeO2-NPs and stabilize them. The CeO2-NPs were characterized by a number of techniques, including X-ray diffraction analysis (XRD), UV–vis spectrophotometry, and high-magnification transmission electron microscopy (TEM). The CeO2-NPs calcined at different temperatures exhibited a cubic fluoride structure with sizes less than approximately 10 nm. The influence of the calcination temperature on the morphology of CeO2-NPs was also investigated. In vitro cytotoxicity studies on neuro2A cells showed a dose-dependent toxicity with non-toxic effect of concentration below 10 mg/mL. The results indicated that gelatin is an interesting material that can be used as a stabilizer in the sol–gel processes for preparing small CeO2-NPs. & 2013 Elsevier Ltd and Techna Group S.r.l. All rights reserved. Keywords: A. Sol–gel processes; B. Electron microscopy; D. CeO2 Gelatin, the protein from collagen, has a three-chain helical structure in which individual helical chains are stranded in a super-helix around the common molecular axis [13–15]. Gelatin contains positively and negatively charged as well as hydrophobic domains folded into a The cytotoxicity of nanoparticles was evaluated by the method using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay [23]. Briefly, neuro2A cells were seeded at a density of 1 104 cells per well in 96 well plates and incubated for 24 h. Thereafter, the cells were treated with various concentrations of nanoparticles in the presence of 10% FBS. The sample S3 was suspended in a stock solution at 5 mg/ml in a solution of dimethyl sulfoxide (DMSO)/DDW water. After 24 h of incubation, 20 ml of 5 mg/ml MTT in the PBS buffer was added to each well, and the cells were further incubated for 4 h at 37 1C. The medium containing unreacted dye was discarded, and 100 ml of DMSO was added to dissolve the formazan crystal formed by live cells. Optical absorbance was measured at 590 nm (reference wavelength 630 nm) using a microplate reader (Statfax–2100, Awareness Technology, USA), and cell viability was expressed as a percent relative to untreated control cells. Values of metabolic activity are presented as mean7SD of triplicates. This condition-dependent experiment and our previous research [25,26] demonstrated that the introduction of gelatin played double roles—stabilizing agent for initial ions and final products—in many preparations. Above all, gelatin greatly stabilized the as-formed Ce(OH)4 nuclei and the as-obtained CeO2-NPs through the bonding between the carbonyl groups and surface hydroxyls while simultaneously preventing them from agglomeration. Typical UV–vis absorption spectrum of S3 is shown in Fig. 1. The CeO2-NPs were dispersed in water, and the suspension was then used to perform the UV–vis measurement in the wavelength range of 200–800 nm. The spectrum revealed a characteristic absorption peak at wavelength of 312 nm for S3, which can be assigned to the intrinsic band-gap absorption of CeO2-NPs due to the electron transitions from the valence band to the conduction band. In other words, the band at approximately 300 nm is caused by the absorption of the charge-transfer transition from O 2p to Ce 4f in CeO2 [27,28]. As shown in Fig. 1, the maximum peak in the absorbance spectrum does not correspond to the true optical band gap of the CeO2-NPs. A common way to obtain the band gap of the CeO2 has a kind of typical calcium fluoride (CaF2) structure with space group Fm3 m. Fig. 2 shows the XRD patterns of the dried and calcined CeO2-NPs prepared in the gelatin media. The same crystalline structure for all conditions was observed. All of the detectable Bragg peaks with Miller indices (111), (200), (220), (311), (222), (400), (331), (420), and (422) can be indexed as fluorite cubic structures (JCPDS # 00-043-1002). The broadening of the peaks indicates that the crystallite sizes are below 10 nm, according to the literature [31]. This result indicates that the size of the obtained sample is small, as confirmed by the TEM image and its corresponding size distributions of S3 (Fig. 3). After the as-prepared CeO2-NPs was calcined The results of in vitro cytotoxicity studies after 24 h of incubation with different concentrations of nanoparticles, ranging from 0 to 175 mg/mL, are shown in Fig. 4. Concentrations below 10 mg/mL were not cytotoxic in the MTT assay. The level of cytotoxicity as a function of the concentration decreased as the concentration decreased. but also for applying and extending the green chemistry rules in preparation of nanoparticles. The other advantages of the method are simple synthesis, in a normal atmosphere, and low cost, giving a potential avenue for further practical scaleup of the production process and applications. It is expected that these nanoparticles can find potential applications in different fields such as catalysts, cosmetics, and optical or electrical devices as well as medicinal applications. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 5.46, \"viab\": 82.4}, {\"h\": 24.0, \"mgL\": 10.93, \"viab\": 84.667}, {\"h\": 24.0, \"mgL\": 21.87, \"viab\": 74.933}, {\"h\": 24.0, \"mgL\": 43.75, \"viab\": 75.467}, {\"h\": 24.0, \"mgL\": 87.5, \"viab\": 67.733}, {\"h\": 24.0, \"mgL\": 175.0, \"viab\": 66.8}]}", "role": "assistant" } ]	64.json
[ { "content": "Extract sample context and measurement list. <doc> Available online at www.sciencedirect.com CERAMICS INTERNATIONAL Ceramics International ] (]]]]) ]]]–]]] www.elsevier.com/locate/ceramint Hadi Kargara,n , Faezeh Ghasemib,c , Majid Darroudid,e,nn a Department of Chemistry, Payame Noor University, P.O. Box 19395-3697, Tehran, I.R. Iran b Neurogenic Inflammation Research Centre, School of Medicine, Mashhad University of Medical Sciences, Mashhad 9177948564, Iran Department of Medical Biotechnology, School of Medicine, Mashhad University of Medical Sciences, Mashhad, Iran d Nuclear Medicine Research Center, Mashhad University of Medical Sciences, Mashhad, Iran e Department of Modern Sciences and Technologies, School of Medicine, Mashhad University of Medical Sciences, Mashhad, Iran Received 11 September 2014; received in revised form 15 September 2014; accepted 16 September 2014 c A simple one-step, eco-friendly, and \"green\" approach for the preparation of monodispersed cerium oxide nanoparticles (CeO2-NPs) are described. This process uses nontoxic and renewable degraded agarose as a natural matrix and mild reaction conditions. The agarose acted as a stabilizing and/or capping agent for the CeO2-NPs. The CeO2-NPs were successfully grown at different calcination temperatures within the agarose matrices, and their crystallite structures were characterized using various methods, including FESEM, PXRD, FTIR, TGA/DTA and UV–vis spectroscopy techniques. This process was found to be comparable to those obtained from conventional preparation methods that use hazardous materials proving to be an excellent alternative for the preparation of CeO2-NPs, using bio-organic materials. In vitro cytotoxicity studies on L929 cells, a non-toxic effect in all concentration (up to 800 μg/ml) was illustrated and we believe that these samples will have viable applications in different fields in medicine. & 2014 Elsevier Ltd and Techna Group S.r.l. All rights reserved. Keywords: D. CeO2; Nanoparticles; Agarose; Bioorganic; Cytotoxicity E-mail addresses: (H. Kargar), (M. Darroudi). http://dx.doi.org/ 0272-8842/& 2014 Elsevier Ltd and Techna Group S.r.l. All rights reserved. sensors [7], ultraviolet absorbers [8], polishing materials [9], medicine [10], and other fields. A number of preparation methods have been used to synthesis of CeO2-NPs, including hydrothermal [11], sol–gel [12], co-precipitation [13], polymeric precursor [14], microwave-assisted heating [15], sonochemical [16], and other methods. Some of these routes have been used in solutions containing polymers and surfactants in order to improve or control the size and shape of CeO2-NPs. In fact, natural polymers as a category of macromolecules can also be used as biotemplates in the green synthesis of CeO2-NPs. When these polymers are used as a capping/stabilizing agent, the diameter of nanoparticles can be logically controlled [17]. We have also reported the green synthesis of cerium oxide nanoparticles, stabilized with gum molecules via a sol–gel method [18]. The ease with which nearly monodisperse gum mediated CeO2-NPs were synthesized, provided an encouragement for the controlled synthesis of plant extract protected CeO2-NPs. Gel formation occurs when cooling the hot aqueous agarose solution to ambient temperature which is stable over a wide range of pH from 3 to 9. The gelation phenomena in agarose created by the presence of H-bonds can be destroyed by any factor lead to the destruction of H-bonds. It is because of this particular gelation properties combined with the considerable stability, shrink, and swell features, agarose gel has been widely used in different subjects such as pharmaceutical, medicine, textile, chemical industries, etc. [20–22]. In our recent studies, bioorganic polymers such as gelatin, starch, and honey as \"green\", ecofriendly degradable materials, and abundant in nature have been used as the capping and/or stabilizing agents for the synthesis of different nanomaterials [23–25]. Therefore, agarose was chosen as a new bio–based medium for the synthesis of water-soluble CeO2-NPs. Caruso et al. [26] used the agarose biopolymer matrix as a sacrificial media to prepare macroporous metal oxides. Agarose is a green matrix, easily available, eco-friendly and cost-effective as compared to other stabilizers available on the market. Indeed, a general and simple method to use of biopolymer as a template for preparing of metal oxide nanoparticles would be advantageous given their potential application in many areas. In this work, an improved sol–gel route was used for synthesis of CeO2-NPs. The CeO2-NPs were first synthesized with cerium nitrate hexahydrate and agarose was used as starting material at different heating treatments. All the materials used were of analytical grade and were used without any purification. Cerium (III) nitrate hexahydrate was purchased from Fluka (Germany) and agarose was purchased from Sigma-Aldrich (USA). All glasswares used The prepared CeO2-NPs were characterized by PXRD (Philips, X'pert, Cu Kα), FTIR (ST-IR\\ST-SIR spectrometer), TGA/DTA (Q600), UV–vis (Evolution 300s Thermo Fisher Scientific), and FESEM (Carl Zeiss Supra 55VP). L929 cells were obtained from Pasteur Institute (Tehran, Iran). Cells were maintained at 37 1C in a humidified atmosphere (90%) containing 5% CO2. Cells were cultured in highglucose Dulbecco's modified Eagle's medium (DMEM) (4.5 g/l) with 10% (v/v) fetal bovine serum, 100 units/ml penicillin and 100 mg/ml streptomycin. The cell viability was determined using a modified 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium (MTT) assay [27]. For MTT assay, 5000 cells were seeded in each well of a 96-microwell plate and treated with various concentrations of Seo agarose (0–800 mg/ml) for 24 h. MTT solution in phosphate-buffered saline (PBS, 5 mg/ml) was added to a final concentration of 0.05%. After 3 h, the formazan precipitate was dissolved in DMSO. The absorbance at 570 and 620 nm (background) was measured using a StatFAX303 plate reader. All experiments were carried out in triplicate; the percentage of viable cells was calculated as the mean7SD and as a percentage of non-treated control groups, which was assumed to be 100% and morphological deformations of the cells were also examined. Research on polymer–metal oxide nanoparticles received much attention in recent years owing to their application in wide and different fields. Among the wide variety of polymer matrices, biopolymers become the preferred choice as they are readily available, inexpensive, environmentally green and Fig. 1. Chemical structure of repeating unit of agarose. more amenable to scale up. Moreover, the oxygen rich functionalities of the biopolymers and their affinity towards metals and metal oxides make them ideal candidates for the stabilization of nanoparticles [28]. Agarose, which forms a gel, develops a network of narrow channels due to the interpenetrating H bonds between the sugar moieties with pore sizes 200 nm [29]. These nanochannels and nanocages provide pathways that CeO2-NPs can occupy. The thermogravimetric and derivative analysis (TGA–DTA) curves of the as-prepared aqueous agarose gel containing zinc precursor are shown in Fig. 2. The heating process was started at about 20 1C, and then increased up to 1000 1C along with a temperature rate of 10 1C/min. The first weight loss between 40 and 120 1C is an initial mass loss of 74% attributable to the loss of adsorbed water, dehydration of cerium hydroxide, and decomposition of chemically bound groups [30]. The second and third weight losses of 6% take place in the range of 240–360 1C, associated with further oxidation of cerium components [25]. The total mass loss of the sample is about 80%. 4 H. Kargar et al. / Ceramics International ] (]]]]) ]]]–]]] crystal was about 10.5 nm. Compared to the FESEM crystallite values the Scherrer formula gave lower crystallite sizes. This may be attributed to the nonuniformity in the interplanar spacing due to stress in the crystallites can also produce the broadening of the PXRD peaks [35]. Furthermore, welldefined Bragg's peaks in the PXRD diffraction confirm that the nature of CeO2-NPs was formed in good crystal quality [36]. The cytotoxicity effect of Seo agarose was evaluated on L929 cells. This cell line was treated for 24 h with different concentrations of Seo agarose (0–800 mg/ml). As shown in Fig. 7, no significant cytotoxic effect was observed in the L929 cell line in all concentrations. It is expected that these nanoparticles have the potential to improve in different applicable fields such as cosmetics and optical/electrical devices as well as medicinal applications. Statistical analysis was performed using ANOVA and posthoc Tukey test using SPSS software. Significant differences were established at po0.05. studies on L929 cells, non-toxic effect in all concentrations was shown. [22] H.H. Jeong, J.H. Lee, Y.M. Noh, Y.G. Kim, C.S. Lee, Macromol. Res. 21 (2013) 534–540. emulsions using response surface methodology, LWT – Food Sci. Technol. 48 (2012) 82–88. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 50.0, \"viab\": 105.86}, {\"h\": 24.0, \"mgL\": 100.0, \"viab\": 103.06}, {\"h\": 24.0, \"mgL\": 200.0, \"viab\": 104.97}, {\"h\": 24.0, \"mgL\": 400.0, \"viab\": 100.0}, {\"h\": 24.0, \"mgL\": 800.0, \"viab\": 96.943}]}", "role": "assistant" } ]	63.json
[ { "content": "Extract sample context and measurement list. <doc> http://tih.sagepub.com/ Toxicol Ind Health published online 5 December 2013 Ahamed Mohammad Javed Akhtar, Sudhir Kumar, Hisham A Alhadlaq, Salman A Alrokayan, Khalid M Abu-Salah and Maqusood DOI: http://tih.sagepub.com/content/early/2013/12/03/0748233713511512 The online version of this article can be found at: Published by: http://www.sagepublications.com Additional services and information for Toxicology and Industrial Health can be found at: Email Alerts: http://tih.sagepub.com/cgi/alerts Subscriptions: http://tih.sagepub.com/subscriptions Reprints: http://www.sagepub.com/journalsReprints.nav Permissions: http://www.sagepub.com/journalsPermissions.nav >> OnlineFirst Version of Record - Dec 5, 2013 What is This? Toxicology and Industrial Health 1–13 The Author(s) 2013 Reprints and permissions: sagepub.co.uk/journalsPermissions.nav DOI: tih.sagepub.com Mohammad Javed Akhtar1 , Sudhir Kumar2 , Hisham A Alhadlaq1,3, Salman A Alrokayan4 , Khalid M Abu-Salah1 and Maqusood Ahamed1 Copper oxide nanoparticles (CuO NPs) are of great interest in nanoscience and nanotechnology because of their broad industrial and commercial applications. Therefore, toxicity of CuO NPs needs to be thoroughly understood. The aim of this study was to investigate the cytotoxicity, genotoxicity, and oxidative stress induced by CuO NPs in human lung epithelial (A549) cells. CuO NPs were synthesized by solvothermal method and the size of NPs measured under transmission electron microscopy (TEM) was found to be around 23 nm. The 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide (MTT) and lactate dehydrogenase (LDH) assays showed that CuO NPs (5–15 mg/ml) exert cytotoxicity in A549 cells in a dose-dependent manner. Comet assay suggested concentration-dependent induction of DNA damage due to the exposure to CuO NPs. The comet tail moment was 27% at 15 mg/ml of CuO NPs, whereas it was 5% in control (p < 0.05). The flow cytometry data revealed that CuO NPs induced micronuclei (MN) in A549 cells dose dependently. The frequency of MN was 25/103 cells at 15 mg/ml of CuO NPs, whereas it was 2/103 cells for control. CuO NPs were also found to induce oxidative stress in a concentration-dependent manner, which was indicated by induction of reactive oxygen species (ROS) and lipid peroxidation along with glutathione depletion. Moreover, MN induction and DNA damage were significantly correlated with ROS (R2 1⁄4 0.937 for ROS vs. olive tail moment, and R2 1⁄4 0.944 for ROS vs. MN). Taken together, this study suggested that CuO NPs induce genotoxicity in A549 cells, which is likely to be mediated through ROS generation and oxidative stress. Nanotoxicity, nanoparticle, oxidative stress, apoptosis, DNA damage At present, metal oxide nanoparticles (NPs) are used in manufacturing of hundreds of commercial products, and their industrial applications are expected to expand during the next decade. Metal oxide NPs are of great interest in nanotechnology, in part because the family of metal oxides offers intriguing new properties for applications in future technologies. Copper oxide (CuO) NPs are being used in various applications such as antimicrobial preparations, heat transfer fluids, semiconductors, and intrauterine contraceptive devices (Aruoja et al., 2009; Chang et al., 2005; Zhou et al., 2006). Furthermore, CuO NPs, for example, can be used to dope other materials for photocatalysis (Song et al., 2008), can be added to fluids to create novel thermal properties as the so-called nanofluid (Karthikeyan et al., 2008; Khandekar et al., 2008), can aid in the synthesis of fullerenes (Liu et al., 2008), and can be used in energetic materials such as explosives and propellants (Shende et al., 2008). The suitability of NPs in biological and, specifically, biomedical applications must be supported by rigorous studies of their potential toxicity (Singh and Nalwa, 2007; Ye and Mahato, 2008). There is also potential for multiple adverse interactions such as oxidative stress and inflammatory responses (Gwinn and Vallyathan, 2006). Such cellular processes may lead to cell death via cell necrosis or apoptosis. Thus, in the development of suitable NPs for use as drug or biomolecules delivery platforms, induction of potential NPs in adverse cellular reactions must be considered (Kagan et al., 2005). Exposure to NPs typically occurs in the epithelia of the lung, skin, or gastrointestinal tract. The present study uses A549 cells, in vitro model of human lung epithelia, to analyze cytotoxicity, genotoxicity, and oxidative stress-induced by CuO NPs exposed. We determined the cytotoxicity by (3-(4,5-dimethyl thiazol-2-yl)-2,5-diphenyl tetrazolium bromide) MTT and lactate dehydrogenase (LDH) leakage in response to different concentrations of CuO NPs in A549 cells. Genotoxicity induced by CuO NPs in A549 cells was determined by Comet assay, a technique that detects single- and double-strand DNA breaks in individual nuclei by measuring the migration of denatured DNA fragments through an agarose gel (McKelvey-Martin et al., 1993; Plewa et al., 2002). In addition to comet assay, the micronuclei (MN) induction was also determined by flow cytometry that has the capability of analyzing thousands of events rapidly in three dimensions, leading to a reduction in false negative errors, a powerful technique over conventional measurement, which is usually conducted microscopically and is limited to 2- D (Dertinger et al., 2002; Smolewksi et al., 2001). Parameters of oxidative stress, in addition to cytogenotoxicity, have also been determined in the present study. Oxidative stress biomarkers including reduced glutathione (GSH) as a cellular antioxidant, reactive oxygen species (ROS) generation as a collective marker of hydrogen peroxide (H2O2), superoxide anion (O2-), hydroxyl radical (HO• ), and thiobarbituric acid reactive substances (TBARS) as an indicator of membrane lipid peroxidation (LPO) were determined. Fetal bovine serum (FBS), Dulbecco's modified eagle's medium (DMEM), Hank's balanced salt solution (HBSS), penicillin–streptomycin, and trypsin were bought from Invitrogen Co. (California, USA). GSH, MTT, 5,5-dithio-bis-(2-nitrobenzoic acid), 2,7-dichlorofluorescin diacetate (DCFH-DA), pyruvic acid, propidium iodide (PI), O-phthalaldehyde (OPT), 1,1,3,3-tetraethoxypropane, thiobarbituric acid (TBA), sodium dodecyl sulfate (SDS), and copper chloride (CuCl2) were purchased from Sigma-Aldrich (Missouri, USA). All other chemicals used were of the highest purity available from commercial sources. CuO NPs were synthesized by solvo-thermal method as given subsequently. Briefly, 3 mM of CuCl23H2O and 6 mM of 1, 10-phenanthroline were dissolved into the mixed distilled water and ethanol. Then 2 M sodium hydroxide (NaOH) aqueous solution was added under magnetic stirring. This alkaline solution was transferred into a Teflon-lined autoclave with about 80% capacity. The autoclave was then sealed and maintained at 160C for 24 h. After cooling to room temperature, the black precipitates were filtered, washed with distilled water and absolute ethanol several times. The resulting product was then dried at 60C for 6 h to get the dry nanopowder of CuO. Morphology of CuO NPs were evaluated by field emission transmission electron microscopy (FETEM, JEM-2100F, JEOL Inc., Japan) at an accelerating voltage of 200 kV. In brief, dry powder of CuO NPs was suspended in deionized water at a concentration of 1 mg/ml and then sonicated using a sonicator bath at room temperature for 15 min at 40 W to form a homogeneous suspension. For size measurement, sonicated 1 mg/ml CuO NP stock solution was then diluted to a 50–100 mg/ml working solutions. Then a drop of CuO NP suspension was placed onto a carbon-coated copper grid, air-dried, and observed with FETEM. Dynamic light scattering (DLS) used for the characterization of hydrodynamic size and zeta potential of CuO NPs in cell culture medium were performed on Malvern Instruments (Zetasizer Nano-ZS, UK) as described by Murdock et al. (2008). Briefly, dry powder of CuO NPs was suspended in cell culture medium at a concentration of 15 mg/ml for 24 h. Then suspension of CuO NPs was sonicated using a sonicator bath at room temperature for 15 min at 40 W and performed the measurements. Cells were used between passages 10–20. Cells were cultured in DMEM/F12 medium supplemented with 10% FBS and 100 U/ml penicillin–streptomycin at 5% CO2 and 37C. At 85% confluence, cells were harvested using 0.25% trypsin and were subcultured into 25 cm2 flasks, 6-well plates, or 96-well plates according to the selection of experiments. Cells were allowed to attach to the surface for 24 h prior to treatment. CuO NPs were suspended in cell culture medium and diluted to appropriate concentrations (5, 10, and 15 mg/ml). The appropriate dilutions of CuO NPs were then sonicated using a sonicator bath at room temperature for 10 min at 40 W to avoid NP agglomeration prior to administration to the cells. Selection of dosage range (5–15 mg/ml) and exposure time of CuO NPs was based on a preliminary doseresponse study (data not shown). Viability of cells after exposure to CuO NPs was assessed by MTT assay as described by Mossman (1983) with some specific modifications (Akhtar et al., 2010). The MTT assay assesses the mitochondrial function by measuring ability of viable cells to reduce MTT into blue-formazon product. Briefly, 1000 cells/well were seeded in 96-well plates and exposed to CuO NPs at the concentrations of 5, 10, and 15 mg/ml for 24 h. At the end of exposure, medium was removed from each well to avoid interference of NPs and replaced with new medium containing MTT solution in an amount equal to 10% of culture volume and incubated for 3 h at 37C until a purple colored formazan product developed. The resulting formazan product was dissolved in acidified isopropanol. Further, the 96-well plate was centrifuged at 2300g for 5 min so that the remaining NPs present in the solution settle. Then, a 100 ml supernatant was transferred to other fresh wells of 96-well plate and absorbance was measured at 570 nm by using a microplate reader (Synergy-HT, BioTek, Virginia, USA). Lactate dehydrogenase (LDH) is an enzyme widely present in cytosol, which converts lactate to pyruvate. When plasma membrane integrity is disrupted, LDH leaks into culture media and its extracellular level is elevated. LDH assay was carried out with the method described earlier (Welder et al., 1991; Wroblewski and LaDue, 1955). In brief, 10,000 cells per well were seeded in 96-well plate and exposed to CuO NPs at the concentrations of 5, 10, and 15 mg/ml for 24 h. At the end of exposure, 96-well plate was centrifuged at 2300g for 10 min to get the cell culture media. Then, a 100 ml of culture media transferred to new fresh tube containing 100 ml of sodium pyruvate (2.5 mg/ml phosphate buffer) and 100 ml of reduced nicotinamide adenine dinucleotide (NADH) (2.5 mg/ml phosphate buffer) in a total volume of 3.0 ml (0.1 M potassium phosphate buffer, pH 7.4). The rate of NADH oxidation was determined by following the decrease in absorbance at 340 nm for 3 min at 30 s interval using a spectrophotometer (Thermo-Spectronic, New York, USA). The amount of LDH released is represented as LDH activity (IU/l) in culture media. Comet assay was performed as described by Singh et al. (1988) with some specific modifications (Ali et al., 2010). In brief, 80,000 cells/well were seeded in a 12-well plate. After 24 h of seeding, cells were treated with different concentrations of CuO NPs (5–15 mg/ml) for 24 h. At the end of exposure, cells were trypsinized and resuspended in DMEM supplemented with 10% FBS and the cell suspension was centrifuged at 2300g for 5 min at 4C. The cell pellet was finally suspended in ice-chilled phosphate buffer saline for comet assay. Then, 15 ml of cell suspension (approximately 20,000 cells) were mixed with 85 ml of low melting point agarose (0.5%) and layered on one end of a frosted plain glass slide, precoated with a layer of 200 ml normal agarose (1%). Thereafter, it was covered with a third layer of 100 ml low melting point agarose (0.5%). After solidification of the gel, the slides were immersed in a freshly prepared lysing solution (2.5 M sodium chloride (NaCl), 100 mM disodium ethylenediaminetatraacetate (Na2EDTA), and 10 mM Tris pH 10 with 10% dimethyl sulfoxide (DMSO) and 1% Triton X-100) overnight at 4C. The slides were then placed in a horizontal gel electrophoresis unit. Fresh cold alkaline electrophoresis buffer (300 mM NaOH, 1 mM Na2EDTA, and 0.2% DMSO, pH 13.5) was poured into the chamber and left for 20 min at 4C for DNA unwinding and conversion of alkali-labile sites to single-strand breaks. Electrophoresis was carried out using the same solution at 4C for 20 min at 15 V. The slides were neutralized gently with 0.4 M tris buffer at pH 7.5 and stained with 75 ml ethidium bromide (20 mg/ml). The slides were stored at 4C in a humidified slide box until scoring. Slides were scored at a final magnification of 400 using an image analysis Komet 5.5 (Kinetic Imaging Limited, Liverpool, UK) attached to a fluorescent microscope (Leica-Optiphase, Germany) equipped with appropriate filters. The undamaged cell resembles an intact nucleus without a tail and a damaged cell has the appearance of a comet. The comet parameter was used to measure the mean tail moment of the cells. Images from 200 random cells (50 cells from each of 4 replicate slides) were analyzed for each experiment. MN formation due to exposure to CuO NPs was examined by flow cytometer as described by Nusse et al. (1994). Briefly, cells were exposed to CuO NPs at the concentrations of 5, 10, and 15 mg/ml for 24 h. After exposure, the cells were washed with cold phosphate buffer saline. Further cell suspension was centrifuged for 5 min at 500g and supernatant was removed, the cell pellet was suspended in solution I (10 mM NaCl, 3.4 mM sodium citrate, 25 mg/ml PI, 0.01 mg RNase from bovine pancreas, and 0.3 ml/ml triton-X). After 1 h at room temperature, an equal volume of solution II (78.1 mM citric acid, 40 mg/ml PI, and 0.25 M sucrose) was added. After 15 min, the suspension was filtered through a 53-mm nylon mesh and stored on ice until analyzed on Flow cytometer (Becton-Dickinson LSR II, California, USA) using 'Cell Quest' 3.3 analysis software. The production of intracellular ROS was measured using DCFH-DA (Wang and Joseph, 1999) with some specific modifications (Akhtar et al., 2010). The DCFH-DA passively enters the cell where it reacts with ROS to form the highly fluorescent compound dichlorofluorescein (DCF). Briefly, 10 mM DCFH-DA stock solution (in methanol) were diluted in culture medium without serum or other additive to yield a 100 mM working solution. After exposure to CuO NPs, the cells were washed twice with HBSS. Then the cells were incubated in 1 ml working solution of DCFH-DA at 37C for 30 min. Cells were lysed in alkaline solution and centrifuged at 2300g for 10 min to avoid interference of NPs and cell debris. A 200 ml supernatant were transferred to 96-well plate and fluorescence was measured at 485 nm excitation and 520 nm emission using a microplate reader (Synergy-HT, BioTek). The intensity of untreated control well was assumed to be 100% and data are represented in percentage of control. LPO was assessed by the TBARS assay, which detects mainly malondialdehyde (MDA), an end product of the peroxidation of polyunsaturated fatty acids and related esters. TBARS was measured by slight modification of the method of Ohkawa et al. (1979). Sub-confluent cells were scraped in 75 cm2 flasks, washed two times in isotonic trace element-free Tris-HCl buffer (400 mM, pH 7.3). A 200 ml aliquot of cell suspension were subsequently mixed with 800 ml of LPO assay cocktail containing 0.4% (w/v) TBA, 0.5% (w/v) SDS, 5% (v/v) acetic acid, pH 3.5, and incubated for 60 min at 95C. The sample was cooled using tap water and centrifuged at 2300g for 5 min. The absorbance of the supernatants was read at 532 nm against a standard curve prepared using the MDA standard (10 mM 1,1,3,3 tetramethoxypropane in 20 mM Tris-HCl, pH 7.4). Results were calculated as nmol of TBARS/mg protein. Intracellular GSH was quantified using the method of Hissin and Hilf (1976). Briefly, cells were cultured in 75 cm2 culture flask and exposed to CuO NPs at the concentrations of 5, 10, and 15 mg/ml for 24 h. After the exposure, the cells were lysed in 20 mM Tris (pH 7.0) and the centrifuged at 2300g for 15 min at 4C. Further, protein of the supernatant was precipitated using 1% perchloric acid and again centrifuged at 2300g for 10 min at 4C to get the supernatant. Then 20 ml of supernatant were mixed with 160 ml of 0.1 M potassium phosphate-5 mM EDTA buffer (pH 8.3), and 20 ml OPT (1 mg/ml in methanol) in a black 96-well plate. After 2 h of incubation at room temperature in the dark, fluorescence was measured at the emission wavelength of 460 nm and excitation wavelength of 350 nm. The amount of GSH was expressed as nmol GSH/mg protein. Protein concentration was measured by the Bradford method (Bradford, 1976) using Bradford reagent (Sigma-Aldrich, Missouri, USA) and bovine serum albumin as the standard. Figure 1(a) shows the typical TEM image of the CuO NPs. This picture shows that the majority of the particles were in spherical shape with smooth surfaces. TEM average diameter was calculated by measuring over 100 particles in random fields of TEM view. The average TEM diameter of CuO NPs was approximately 23 nm. The average hydrodynamic diameter and zeta potential of the CuO NPs suspension in culture medium determined by DLS were found to be 87 nm and -25 mV, respectively (Figure 1(b)). The higher size of CuO NPs in aqueous suspension than the TEM size might be due to the tendency of particles to aggregate in aqueous state. This finding is supported by other investigators (Bai et al., 2009) and has been briefly discussed in our previous publication (Ahamed et al., 2010). Cytotoxicity of CuO NPs against A549 cells was measured by MTT and LDH assays. MTT results show that cell viability was decreased by CuO NPs and the degree of reduction was dose dependent. Cell viability was decreased to 81, 56, and 31% for the concentrations of 5, 10, and 15 mg/ml, respectively (p < 0.05) (Figure 2(a)). Similarly, LDH leakage, an indicator of membrane damage was also induced by CuO NPs in A549 cells in a dose-dependent manner. LDH level in cytosol was found to be 1.5-, 2-, and 2.8-fold higher for the concentrations of 5, 10, and 15 mg/ml, respectively (p < 0.05) (Figure 2(b)). Cell morphology was assessed following exposure to different concentrations of CuO NPs using a Leica phase contrast microscope. Figure 3 shows the comparative morphologies of control and CuO NPs exposed A549 cells. A significant lowering of cell density and rounding of cells were observed in dosedependent manner supporting the cell viability data. One of the signs of genotoxicity is the induction of DNA damage that can be determined by comet assay, a widely used method for the detection as well as measurement of DNA strand breaks (Anderson and Plewa, 1998; Singh et al., 1988). Comet assay was carried out in A549 cells against CuO NPs at the concentrations of 5, 10, and 15 mg/ml for 24 h. A dose-dependent increase in DNA damage was observed in CuO NP-treated cells evident by an increase in olive tail moment (arbitrary unit). However, no significant DNA damage was observed in the untreated control cells (Figure 4(a) and (b)). MN induction by flow cytometry further confirmed genotoxic potential of CuO NPs in A549 cells. A significant (p < 0.05) increase in the frequency of MN was observed in A549 cells with increasing concentration of CuO NPs. CuO NP exposure (5, 10, and 15 mg/ml for 24 h) led to induction of MN frequency in A549 cells up to 9, 14, and 25, respectively, whereas in untreated control the frequency was 4 (Figure 5(a) and (b)). It has been suggested that ROS generation and oxidative stress mediate the genotoxicity of NPs (Ahamed et al., 2011; Akhtar et al., 2010, 2012; Nel et al., 2006). Therefore, the ability of CuO NPs to induce oxidative stress in A549 cells was assessed by measuring ROS, LPO, and GSH levels. As expected, we observed that CuO NPs have the potential to induce oxidant generation (ROS and LOP) and reduce antioxidant level (GSH) in A549 cells in a dosedependent manner (Figure 6(a), (b), and (c)). This study reported the genotoxic potential of CuO NPs determined by Comet assay and flow cytometry. Several studies have shown a positive correlation among oxidative stress, genotoxicity, and apoptosis (Calviello et al., 2006; Chen et al., 2005). Generation of oxidative stress may also cause DNA damage, which may lead to genotoxicity and cancer (Bump and Malakar, 1998). Results of the present study demonstrate that CuO NP exposure to A549 cells can induce cytotoxicity, genotoxicity, and oxidative stress in a dose-dependent manner. Our study suggests the involvement of oxidative stress in the mechanism of CuO NPs-induced genotoxicity. We found that CuO NPs significantly induced genotoxic effect in human lung cells. Currently, NP genotoxicity testing is based on in vitro methods established for hazard characterization of chemicals (Fubini et al., 2010; Landsiedel et al., 2010). Comet assay is one of the important and well-applied in vitro methods in genotoxicology and DNA damage studies. It is an in situ method in which embedded cell on agarose base is lysed and electrophoresed on neutral or alkaline conditions (Collins, 2004). The results obtained by the comet indicated that CuO NPs were able to cause a significant increase in tail moment in A549 cells when exposed for 24 h at the concentrations of 15 mg/ml. In addition to comet assay, the MN induction measurement is an established cytogenetic assay that can detect acentric fragments and lagging chromosomes induced by clastogens and aneugens (Mavournin et al., 1990). Manual measurement of MN induction is usually conducted microscopically and is limited to two dimensions and, hence, some MN are not counted due to a lack of visibility. The MN induction measurement by flow cytometry, however, is a powerful technique that has the capability of analyzing thousands of events rapidly in three dimensions leading to a reduction in false negatives errors. Progress has been made to automate the scoring of MN by flow cytometry (Dertinger et al., 2002; Roman et al., 1998; Smolewksi et al., 2001). The results obtained by the MN assay indicated that CuO NPs were able to cause a significant increase in MN induction in A549 cells when exposed for 24 h at the concentrations of 15mg/ml. Oxidative stress has been suggested to play an important role in the mechanism of toxicity of a number of compounds either by the production of ROS or by depleting cellular antioxidant capacity. Cellular integrity is affected by oxidative stress when the production of ROS overwhelms antioxidant defense mechanism (Halliwell and Gutteridge, 2007). ROS are oxygen-containing molecules, such as H2O2, O2-, and HO• , that have a greater chemical activity than molecular oxygen. There are many evidences showing that NPs increase ROS production and can cause cell death in different types of cultured cells (Akhtar et al., 2010; Park et al., 2008; Peters et al., 2007; Wang et al., 2009). Recently, we have reported oxidative stress as a major mechanism of toxicity induced by CuO NPs in mouse fibroblast cells (Balb 3T3) (Akhtar et al., 2012), whereas Sun et al. (2012) have reported autophagy as a major mechanism of cell death in A549 cells. Adding to this conflict, however, Song et al (2012) have reported oxidative stress-induced MN induction and DNA damage as the major mechanisms of cell death by several metal NPs including NPs of CuO. This discrepancy prompted us to investigate CuO NPs-induced MN induction and DNA damage, within our limit, in A549 cells along with other parameters of oxidative stress. In the present study, CuO NPs induced significantly higher MN induction and DNA damage, which correlated well with ROS generation in a concentration-dependent manner in A549 cells. Certain other nanomaterials, such as C60, silica, and talc NPs mediate cytotoxicity primarily through LPO, another marker of oxidative stress (Ahmad et al., 2012; Isakovic et al., 2006; Lin et al., 2006; Sayes et al., 2005). Recently, Scarfı` et al. (2009) have reported that plasma membrane contact with quartz, a kind of crystalline silica, is sufficient to trigger membrane LPO, tumor necrosis factor (TNF)-a release, and cell death in mouse macrophages (RAW-264.7). Our present data also suggest that contact of CuO NPs with cell membranes of A549 cells initiate in the generation of ROS and, in turn, LPO in a concentration-dependent manner. Antioxidant GSH is the most abundant nonproteinous tripeptide containing a sulfhydryl group in virtually all cells and plays a significant role in many biological processes. It also constitutes the first line of the cellular defense mechanism against oxidative injury and is the major intracellular redox buffer in ubiquitous cell types (Meister, 1989). GSH acts as a co-substrate in the GSH peroxidase-catalyzed reduction of H2O2 or LPO leading to its depletion. Previous studies demonstrated that ROS generation following GSH depletion caused mitochondrial damage (Martensson et al., 1989; Meister, 1995), which has been implicated in apoptosis (Green and Reed, 1998). Therefore, increases or decreases in these responses can be interpreted as evidence for oxidative stress as the cell compensates either for increased stress by upregulating the production of antioxidants or the exhaustion of cellular stores of antioxidants by oxidation from reactive nitrogen species (RNS) or ROS. There was a significant depletion of GSH induced due to CuO NP exposure in a concentration-dependent manner when compared with untreated A549. We found that CuO NPs significantly induced genotoxic effect in human lung cells. It has been demonstrated that Cu ions were released from the surface of CuO NPs when suspended in aqueous state (Midander et al., 2009). However, we have not examined the degree of ionization of CuO NPs in aqueous suspension and their biological effects. However, Fahmy and Cormier (2009) studied the cytotoxic response of CuO NPs in airway epithelial cells and found that the release of Cu ions in the cell culture media did not significantly contribute to the cytotoxic response and the oxidative damage of CuO NPs. Further, Griffitt et al. (2007) have also demonstrated that dissolved portion of Cu ions from NPs is insufficient to produce mortality in Zebrafish exposed to CuO NPs. Our study along with other reports, thus, suggests that cytotoxicity and oxidative stress are associated with NPs of CuO per se rather than the released Cu ions. Taken together, we have reported the potential of CuO NPs to cause genotoxicity in human lung epithelial cells (A549), which may be mediated through the induction of oxidative stress as well as compromising cellular antioxidant potential. This work suggests that the commercial and industrial application of CuO NPs should be carefully evaluated as to their potential hazardous effects to human health. The authors declared no conflicts of interest. The authors extend their appreciation to the Deanship of Scientific Research at King Saud University for funding this work through research group no. RGP-VPP-308. protein 70, oxidative stress and apoptosis in Drosophila melanogaster. Toxicology and Applied Pharmacology 242: 263–269. study of toxicologic agent. In Vitro Cellular and Developmental Biology 27: 921–926. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 87.0, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 5.0, \"viab\": 81.514}, {\"h\": 24.0, \"mgL\": 10.0, \"viab\": 56.477}, {\"h\": 24.0, \"mgL\": 15.0, \"viab\": 31.441}]}", "role": "assistant" } ]	61.json
[ { "content": "Extract sample context and measurement list. <doc> (wileyonlinelibrary.com) DOI ABSTRACT: Despite the widespread use of silica nanoparticles (SiO2 NPs) in biological and medical fields, their adverse effects have not been clearly elucidated. In this study, spherical SiO2 NPs with a 50 nm diameter were used to study their interaction with HaCaT cells. SiO2 NPs were found to be readily internalized into HaCaT cells and localized in the cytoplasm, lysosomes and autophagosomes. Decreased cell viability and damaged cell membrane integrity showed the cytotoxicity of SiO2 NPs. Significant glutathione depletion and reactive oxygen species generation, which reduced the cellular antioxidant level, could be the major factor of cytotoxicity induced by SiO2 NPs. 2013 John Wiley & Sons, Ltd. Keywords: Silicon dioxide nanoparticles; Transmission electron microscope (TEM); HaCaT cells; Cytotoxicity; Oxidative stress The unique physicochemical properties of engineered nanoparticles (NPs) are in part attributed to their small size, larger surface area, surface reactivity and surface structure (Auffan et al., 2009; Nel et al., 2006; Oberdorster et al., 2005). As a kind of non-metal oxide, silica NPs (SiO2 NPs) have been extensively used in biomedicine, pharmaceuticals and other industrial manufacturing fields, such as DNA delivery (Bharali et al., 2005; Roy et al., 2005), cancer therapy (Czarnobaj & Lukasiak, 2007; Kim et al., 2007; Ohulchanskyy et al., 2007) and enzyme immobilization (Qhobosheane et al., 2001). Recently, a number of studies have focused on the interaction between SiO2 NPs and biological systems to explore their biocompatibility at the systemic and cellular level (Vasir & Labhasetwar, 2008). In vitro experiments have revealed cytotoxic effects of NPs using various human cell types such as the endothelial cell line EAHY96 (Napierska et al., 2009), HaCaT keratinocytes (Yang et al., 2010) and A549 cells (Lison et al., 2008). SiO2 NPs were toxic to marine macrophages and keratinocytes (Park & Park, 2009; Park et al., 2009) and induced gene expression related to surface area (Waters et al., 2009). Moreover, SiO2 NPs caused a proinflammatory stimulation of endothelial cells (Peters et al., 2004). On the other hand, an in vivo study demonstrated that silica NPs were not toxic and, therefore, could be used in vivo or for other biomedical applications (Jin et al., 2007). As previously was well documented, NPs could be transported into cells through endocytosis (Conner & Schmid, 2003; Huang et al., 2010). In addition, the interaction between NPs and cell membranes could proceed by nonspecific cellular uptake, which should not be ignored (Rieux et al., 2006; Wilhelm et al., 2003). This study was undertaken to explore the interaction between SiO2 NPs and HaCaT cells, as characterized by transmission electronic microscopy (TEM), laser scanning confocal microscope (LSCM) and lactate dehydrogenase (LDH) assay. The oxidative stress induced by SiO2 NPs was determined by reactive oxygen species (ROS) and glutathione (GSH). SiO2 NPs were prepared according to the Stober sol–gel method at the Iron and Steel Research Institute (Beijing, China). The size distribution and crystalline phase of the NPs were characterized by TEM. SiO2 NPs without any further treatment were dispersed in serum-free medium, and the suspension was ultrasonicated for 30 min every 10 min for vortex to reach well dispersion. The dispersibility of SiO2 NPs was detected by a Hitachi TEM (H-7650; Hitachi, Ltd., Tokyo, Japan) with a GATAN digital camera (Pleasanton, USA). The HaCaT cell line, an immortalized keratinocyte cell line established from adult human skin cells, was purchased from the China center for type culture collection of Wuhan University. HaCaT cells were grown in 25 cm2 cell culture flask in minimal medium (Gibco BRL, Invitrogen AG, Basel, Switzerland) supplemented with 10% fetal bovine serum (Sijiqing Biological Engineering Materials Co., Ltd., Hangzhou, China) and 1% penicillin/streptomycin. Cell cultures were kept at 37 °C under 5% CO2 humidified atmosphere. For splitting and seeding, the cells were detached from the flask with 0.25% trypsin-EDTA (0.5 g *Correspondence to: Chan Jin, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, 201800, China. Email: a Physics College, Henan normal university, Xinxiang, 453007, China c Institute of Biophysics, Second Military Medical University, Shanghai, 200433, China trypsin, 0.2 g EDTA.4Na in Hanks' balanced salt solution; BoGuang Co., Ltd., Shanghai, China). The SiO2 NPs suspension was sonicated for 30 min every 10 min for vortex before diluted with serum-free medium to get different concentrations (25, 75, 100, 200, 400 and 500 μg ml–1 ). The SiO2 NPs suspension was co-incubated with HaCaT cells for 4 h at 37 °C under a 5% CO2 humidified atmosphere. HaCaT cells treated without SiO2 NPs were monitored as the control. After the treatment of SiO2 NPs for 4 h, HaCaT cells were washed with Hanks' balanced salt solution and fixed with 4% glutaraldehyde and 4% paraformaldehyde overnight at 4 °C. Then, the cells were centrifuged at 850 g for 5 min and washed with 0.1 M phosphate-buffered saline (PBS) three times. All the cells were fixed in 3% glutaraldehyde containing a small amount of blood plasma overnight at 4 °C for coagulating. Afterward, the cells were post-fixed with 1% osmium tetraoxide for 2 h at room temperature in 1 mm3 masses (Gupta & Gupta, 2005; Wilhelm et al., 2003). The cells were then dehydrated in a graded series of ethanol and acetone (Gupta et al., 2003; Kim et al., 2006; Wang et al., 2007) and embedded in Epon812. Ultrathin section (70 nm) were cut by ultramicrotome (UC6; Leica Ltd. Co., Solms, Germany), transferred on to 200-mesh copper grids, strained with uranyl acetate and lead nitrate and observed under TEM. HaCaT cells incubated with SiO2 NPs were washed three times with 0.1 M PBS and then fixed with 4% formaldehyde for 15 min. Once fixed, the sample was washed with PBS and a permeabilizing buffer (10.3 g of sucrose, 0.292 g of NaCl, 0.06 g of MgCl2, 0.476 g of Hepes buffer, 0.5 ml of Triton-X-100, in 100 ml of water, pH 7.2) was added for 5 min. Then the sample was incubated at 37 °C for 5 min in 1% bovine serum albumin/ PBS. Afterward, the sample was stained with 25 μmol l–1 Hoechst 33342 and 5 μmol ml–1 propidium iodide (PI) (Molecular Probes; Beyotime Institute of Biotechnology, Jiangsu, China) for 10 min. After washing with PBS, the HaCaT cells were observed by LSCM (TCS SP2; Leica Ltd., Co., Solms, Germany). The false-colored images were obtained with an excitation wavelength of 350 nm or 543 nm, and emission fluorescence was collected at wavelengths of 380–500 nm or 555–600 nm. LDH leakage, which was a measure of cytotoxicity based on membrane damage, was determined using a commercial LDH kit (Beyotime Institute of Biotechnology, Jiangsu, China). Released LDH catalyzed the oxidation of lactate to pyruvic acid with simultaneous reduction of NAD+ to NADH. The rate of NAD+ reduction was directly proportional to LDH activity in the cell medium. Different SiO2 NPs concentrations were used for LDH analysis. Absorption was measured using a Bio-Tek Elx800 (Vermont, USA) at 490 nm. Each experiment was repeated at least three times. In vitro cell viability was accessed by a methyl thiazolyl tetrazolium (MTT) toxic assay (Mosmann, 1983), which was widely used to evaluate the cytotoxicity. HaCaT cells in 96-well plates were incubated with SiO2 NPs for 4 h and then the cell viability was determined using a MTT proliferation assay kit. Briefly, 10 μl MTT reagent was added to each well of the plate. After 1 h, the medium was replaced by 100 μl dimethyl sulfoxide and mixed thoroughly to dissolve the formazan crystals. To eliminate potential problems of residual NPs, which could interfere with the assay, NPs were allowed to sediment for 1 h and were then transferred to a new plate. The absorbance was then measured at 490 nm by a multiskan ascent (MK3; Thermo Fisher Scientific Inc., Waltham USA). The intracellular GSH and ROS measurement was used to indicate the oxidative stress caused by SiO2 NPs. After treatment for 4 h, HaCaT cells were collected and centrifuged at 10 000 g for 10 min at 4 °C. The cells were resuspended in 20 μl cell medium. Ten microliters of cells was mixed with 30 μl 5% metaphosphoric acid, then frozen and thawed twice using liquid nitrogen and 37 °C water. GSH and ROS were respectively measured using the reagent kits purchased from Beyotime Institute of Biotechnology according to the manufacturer's instructions. Data were expressed as means ± SD from three independent experiments. Data were analyzed by a one-way ANOVA followed by Dunnett's t-test for comparisons between groups. P < 0.05 was considered significant compared to control. SiO2 NPs were prepared according to the Stober sol–gel method and were characterized by TEM (Fig. 1). The image indicated that SiO2 particles were spherical with a uniform size of about 50 nm in diameter. SiO2 particles were well dispersed in serum-free medium. The TEM technique was used to confirm that SiO2 NPs could enter HaCaT cells and determine the intracellular distribution of SiO2 NPs, as shown in Fig. 2. TEM images showed that most of the mitochondria in cells were injured. SiO2 NPs were found in the cytoplasm, lysosomes and autophagosomes. Noteworthy, no NPs were found in the nucleus. The LSCM images of HaCaT cells were shown in Fig. 3. Hoechst 33342 (blue in Fig. 3) stained DNA when cell membrane integrity was maintained. Once the cell membrane was damaged, nonpermeable PI (red in Fig. 3) could enter the cell. In the control group, only blue Hoechst stain was visible in Fig. 3(A–C), which indicated that cell membrane integrity was maintained. After exposure to 200 μg ml–1 SiO2 NPs, besides the blue stain, the red PI stain was visible in Fig. 3(D–F). The colocalization of red and blue in Fig. 3(F) illustrated that the cell membrane of the HaCaT cell was damaged. HaCaT cells were exposed to different concentrations of SiO2 NPs (25, 75, 100, 200, 400 and 500 μg ml–1 ) for 4 h. The cell viability decreased as a function of the dosage level (Fig. 5). The threshold concentration that induced significant cytotoxicity was 100 μg ml–1 . When the concentration of SiO2 NPs was greater than 100 μg ml–1 , the cell viability decreased to lower than 50%. The cell viability of the 500 μg ml–1 group decreased to about 20% compared to the control. A diverse array of applications take advantage of engineered SiO2 NPs, but their potential risks have not yet been fully evaluated. Recently, a number of studies have focused on the interaction between the SiO2 NPs and biological systems to understand their biocompatibility at the systemic and cellular level. The detailed interaction between SiO2 NPs and HaCaT cells was shown in our study. SiO2 NPs could enter HaCaT cells and be internalized in the cytoplasm, lysosomes and autophagosomes. These results are consistent with previous studies (Conner & Schmid, 2003; Huang et al., 2010). The intracellular red PI stain and increased LDH leakage indicated that the cell membrane integrity was damaged. This result was in accordance with the decreased cell livability detected by MTT assay. Previous studies have shown that SiO2 NPs could react with oxygen molecules and produce superoxide and other ROS through a disproportionate reaction (Al-Rawi et al., 2011; Lin et al., 2006). In this study, decreased GSH and increased ROS induced by SiO2 NPs could be the major factor of cytotoxicity. In the current study, spherical SiO2 NPs with a 50 nm diameter were used to study their interaction with HaCaT cells. SiO2 NPs were readily internalized into HaCaT cells and localized in the cytoplasm, lysosomes and autophagosomes. Decreased cell viability and damaged cell membrane integrity indicated the cytotoxicity of SiO2 NPs. Significant GSH depletion and ROS generation could be the major factor of cytotoxicity induced by SiO2 NPs. The Authors did not report any conflict of interest. This work was supported by National Science Foundation of China (grant no. 11105221), the Youth Foundation of Second Military Medical University (grant no. 2010QN02), State Key Development Program for Basic Research of China (grant no. 2011CB932803) and the National Public Benefit Research Sector (grant no. 201210284–02). </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 4.0, \"mgL\": 25.0, \"viab\": 99.588}, {\"h\": 4.0, \"mgL\": 75.0, \"viab\": 94.643}, {\"h\": 4.0, \"mgL\": 100.0, \"viab\": 52.473}, {\"h\": 4.0, \"mgL\": 200.0, \"viab\": 36.126}, {\"h\": 4.0, \"mgL\": 400.0, \"viab\": 22.253}, {\"h\": 4.0, \"mgL\": 500.0, \"viab\": 19.643}]}", "role": "assistant" } ]	54.json
[ { "content": "Extract sample context and measurement list. <doc> Contents lists available at SciVerse ScienceDirect journal homepage: www.elsevier.com/locate/talanta a Department of Chemistry and Institute of Biomedical Sciences, Fudan University, Shanghai 200433, China b Department of Chemistry and Biochemistry, Texas State University—San Marcos, San Marcos, TX 78666, USA Article history: Received 13 November 2012 Received in revised form 22 January 2013 Accepted 23 January 2013 Available online 29 January 2013 Keywords: Silica nanoparticles Cell uptake Particle size Cytotoxicity In this study, we investigated and reported the cellular uptake efficiency, mechanism, and cytotoxicity of silica nanoparticles (SNPs) with different sizes. Using confocal laser scanning microscope (CLSM), flow cytometry (FCM), and graphite furnace atomic absorption spectrometry (GFAAS), the qualitative and quantitative experimental results showed that the cellular uptake of SNPs toward HeLa cells is sizedependent. To further examine the uptake process, three different inhibitors including sucrose, Filipin III, and Cytochalasin D (Cyt D) were introduced to pretreat the HeLa cells. It appeared that the largest SNPs (SNPs-307.6) take an energy-dependent uptake pathway (clathrin dependent and caveolin independent) while that for the medium size SNPs-167.8 involves clathrin and caveolin dependent endocytosis. In contrast, the smallest SNPs (SNPs-55.6) follow not only energy required clathrin and caveolin dependent endocytosis but also an energy independent pathway to efficiently enter the cells. Moreover, the cellular uptake efficiency of SNPs, which also show excellent biocompatibility, is sizedependent in the order of 55.64167.84307.6 nm. This knowledge is fundamentally important and will facilitate more development of size-defined SNPs as the transporters for various purposes. & 2013 Elsevier B.V. All rights reserved. Although many applications have shown promising potential in disease diagnosis and therapy, the fundamental interactions of SNPs with cells still remain unknown, for a large part. It was reported that the internalization of fluorescein isothiocyanate (FITC) modified SNPs, which is mediated by clathrin and actin dependent endocytosis in both human mesenchymal stem cells 1 Jie Zhu and Lei Liao contributed equally to this work. (hMSCs) and 3T3-L1 cells, is concentration-, time-, and cell typedependent [20]. The possible mechanisms were studied for the cellular uptake of this material in the presence of specific endocytic trafficking inhibitors. It was demonstrated that the uptake of FITC-modified silica nanoparticles is energy-dependent through a clathrin mediated pathway toward HeLa cells and human pancreatic cancer cell line (PANC-1) [21,22]. Further studies revealed that the surface charge on SNPs can not only influence ED50 (the value for the effective dose at 50% of internalization) but also the mechanisms of internalization [20,21]. After surface modification, the silica nanoparticles may be internalized through a caveolin mediated process [21]. The effect of surface charge on the cellular uptake of SNPs toward hMSCs and 3T3-L1 cells was reported by Huang et al. [20]. Their results showed that the uptake of SNPs by hMSCs can be regulated by a threshold of positive surface charge and the modulation of surface charge on the uptake is specific to cell type. Besides the surface charge, the size of SNPs is another important factor which can affect the cellular uptake. It was reported that SNPs with a diameter of 50 nm could be the most suitable candidate to serve as a carrier into the cells [23]. Our previous work demonstrated that the ultra small hollow SNPs possess enhanced endocytosis properties when compared to conventional ones with relatively larger particle sizes [24]. It has also been shown that SNPs with a diameter of 50 nm or less can enter the nucleus and may be used in the application for improving the efficiency of anticancer drug delivery [25]. However, as far as it is concerned, the size effect of SNPs on the cellular uptake mechanism has not been studied in detail. To facilitate theoretical and experimental investigations, studies for cellular uptake of SNPs with consistency in Zeta potential and surface functionality are desired to elucidate the effect of size on the efficiency, mechanism, and cytotoxicity of the uptake process. Herein, we focused on examining the cellular uptake pathway as well as the corresponding efficiency and cytotoxicity of SNPs with similar surface charges but different sizes, in order to get insight into the intracellular properties of SNPs. Interestingly, we discovered that SNPs with different diameters do follow different uptake pathways and the subcellular localization of internalized materials is also size-dependent. We propose that silica nanoparticles with a diameter of less than 160 nm (which can enter the cell but not likely the nucleus) might be candidates for drug delivery and those SNPs having a diameter of 50 nm or less (which can enter the nucleus) could be carriers of siRNA and DNA for gene therapy. Nevertheless, there is no ''rule of thumb'' to predict which endocytic pathway that either SNPs or functionalized SNPs will follow when internalized by the cells. To further understand the pathway for cellular internalization of SNPs will be of utmost importance for many of its applications in biotechnology and biomedicine. All chemicals were used as received without further purification. Filipin III and Cytochalasin D (Cyt D) were purchased from Sigma-Aldrich. The HeLa cell line was provided by the Institute of Biochemistry and Cell Biology, SIBS, CAS (China). Cetyltrimethylammonium chloride (CTAC), tetraethoxysilane (TEOS), 3-aminopropyltrimethoxysilane (APTMS), diethanolamine (DEA), ammonium hydroxide (NH4OH), ethanol, and sucrose were purchased from Shanghai Chemical Corp. The mesoporous silica nanoparticles used in this work were synthesized according to literature procedures [26]. For the synthesis of SNPs-55.6, 7.2 mL of water, 0.9 g of ethanol, 0.26 g of CTAC, and 0.02 g of DEA were mixed and stirred in a water bath at 40 1C for 30 min. Subsequently, 0.73 mL of TEOS was added dropwise within 2 min under stirring. For the synthesis of SNPs-167.8, the same procedure was followed except that the amount of ethanol used was 1.65 g. The solution gradually turned to white and was stirred for another 2 h. The milky mixture was allowed to undergo dialysis for 48 h. The white powders were collected after evaporation of the solvent. The final products were obtained after calcination at 550 1C for 5 h. For the synthesis of SNPs-307.6 [8], 5.1 g of NH4OH solution and 4.5 g of water were added to 38 g of ethanol. Afterwards, 2.08 g of TEOS was added dropwise within 2 min under stirring. With further stirring for 12 h, the milky mixture was centrifuged and the product was obtained by evaporation of the solvent. 5 mg of FITC was mixed with 10 mL of APTMS and stirred for 2 h. The mixture was added to the aqueous solution of SNPs and allowed to react for another 2 h. The unreacted FITC and APTMS were removed by dialysis for 48 h. Transmission electron microscopy (TEM) images were obtained with the aid of a JEOL 2011 microscope operated at 200 kV. Scanning electron microscopy (SEM) images were collected by a Philips XL30 microscope operated at 20 kV. The size distribution and Zeta potential of SNPs were measured by dynamic light scattering (DLS) at 25 1C using Nanosizer ZS-90 (Malvern). Nitrogen sorption isotherms were measured at 77 K by a Micromeritics Tristar 3000 analyzer. Small-angle X-ray scattering (SAXS) patterns were recorded by a Bruker D4 X-ray diffractometer with Ni-filtered Cu Ka radiation (l1⁄41.54056 A) at a voltage of 40 kV and a current ̊ of 40 mA. HeLa cells were incubated with 40 mg/mL SNPs in a serum free medium for 3 h at 4 1C or 37 1C under 5% CO2. After incubation, the cells were fixed with 4% paraformaldehyde and the nuclei were stained with 5 mg/mL 40 ,6-diamidino-2-phenylindole (DAPI) in 10% glycerol. Confocal fluorescence images were obtained by a confocal laser scanning microscope (CLSM) imaging system (Leica TCS SP5) and a 63 oil-immersion objective lens. Excitation at 488 nm was provided by a Multi Ar laser. For flow cytometry (FCM), the cells were harvested by trypsinization and the analysis was carried out by an EPICS ALTRA automatic flow cytometer (Beckman Coulter). HeLa cells were also placed in Modified Eagle0 s Medium (MEM) on 14-mm glass coverslips (5- 108 cells/L for CLSM) or in 60-mm tissue culture dishes (5- 105 cells/dish for FCM) supplemented with 10% Fetal Bovine Serum (FBS) at 37 1C under 5% CO2 and allowed to grow for 24 h. After incubation, the cells were washed with phosphate buffer solutions (PBS, pH1⁄47.4) for three times. Serum free media containing 0.45 mol/L sucrose, 5 mg/L filipin III, or 1 mmol/L Cyt D were then added, respectively. The HeLa cells were incubated afterwards with 40 mg/mL SNPs for 3 h at 37 1C under 5% CO2 before CLSM and FCM analyses. For graphite furnace atomic absorption spectrometry (GFAAS) analyses, HeLa cells were seeded in a 10-cm Petri dish (2- 106 cells/dish) and cultivated with 40 mg/ mL SNPs for 3 h. Subsequently, the cells were washed with PBS for three times and trypsinized. After centrifugation and being washed once, the cells were resuspended in water and centrifuged again for collection. They were dried overnight before 100 mL of 48–51 wt% HF and 9.9 mL of 2 wt% HNO3 were added sequentially to allow dissolution of the SNPs and cells with the aid of sonication. The mass of FITC–SNPs in the HeLa cells was determined by measuring the silicon concentration using GFAAS (Unicam Solar 939). Cell viability was determined by MTT assay. Cells were placed in a 96-well flat bottom plate with 6 - 103 cells per well and allowed to grow overnight prior to exposure to SNPs at different concentrations. After 48 h of incubation, the MTT reagent was added to be converted to a purple formazan product by active mitochondria for 4 h at 37 1C. The product was then dissolved in DMSO and quantified by absorption spectrophotometry at 490 nm with the aid of an enzyme-labelled instrument (SUNOS-TIK SPR-960). SNPs with different sizes were synthesized according to the literatures [8,26]. In order to compare the cellular uptake performance of various SNPs and study the corresponding intracellular uptake mechanisms, FITC was covalently linked to the surface of SNPs for fluorescence detection. TEM and SEM images of FITC functionalized SNPs (FITC–SNPs) are shown in Fig. 1. By measuring the diameters of 100 individual SNPs directly from the TEM images, the narrow size distributions of approximately 55.6 nm (SNPs-55.6), 167.8 nm (SNPs-167.8), and 307.6 nm (SNPs-307.6) are presented in Fig. 2a–c. Comparatively, DLS analyses showed that the diameters of the three SNPs in phosphate buffer solutions (PBS, pH1⁄47.4) are centered at 61.8 nm, 164.2 nm, and 342.0 nm (Fig. 2d), which are slightly larger than the results obtained from TEM images except that of SNPs-167.8. This is reasonable because hydrodynamic diameters are generally larger than the core nanoparticle sizes observed by TEM [23]. Since previous data revealed that the surface charge can not only affect the ability of SNPs to internalize cells and escape endosomal entrapment [21] but also the corresponding cellular uptake mechanisms [20,21], the surface charges on the three SNPs were measured to check the uniformity of this parameter. Table 1 shows the Zeta potentials for various SNPs suspended in PBS before and after the surface modification with FITC. When compared with those of SNPs-55.6 and SNPs-167.8, the Zeta potential of SNPs-307.6 is a little more negative. This small difference can be attributed to the condensation of silanol groups of hydrated SNPs-55.6 and SNPs-167.8 during the calcination process [24]. After modification with FITC, the surface charges on FITC–SNPs increase slightly due to the presence of amino groups (from APTMS used to treat SNPs surface) which are not linked to FITC. In general, the surface charges are similar for FITC–SNPs of different sizes at the physiological pH. Fig. 3 shows the nitrogen adsorption–desorption isotherms of the SNPs before and after the Eukaryotic cells internalize extracellular materials inside cytoplasm through energy independent or energy-dependent uptake pathway [28–30]. Generally, low molecular weight solutes directly transport through the plasma membrane without consuming energy via the energy independent pathway. The energydependent pathway, also known as endocytosis for uptake of relatively large particles, can be hindered when incubations are performed at 4 1C. The endocytosis is divided into at least three subcategories: clathrin dependent, caveolin dependent, and clathrin/caveolin independent [31,32]. Extracellular materials or solutes following the clathrin dependent uptake pathway are first enclosed within clathrin coated vesicles derived from folds or invaginations of the plasma membrane, and then brought into the cells. Sucrose can disturb the formation of clathrin coated vesicles on the cell membrane and thus hinder the clathrin dependent endocytosis [33–35]. On the other hand, clathrin independent endocytosis can occur through the caveolae or lipid rafts. Filipin interacts with 3-b-hydroxysterols in the plasma membrane to form filipin–sterol complexes which subsequently cause the filamentous caveolin-1-coat to rapidly disassemble, leading to inhibition of caveolin mediated endocytosis [35,36]. Cyt D, a potent inhibitor of actin polymerization, was used to inhibit actin related phagocytosis and clathrin/caveolin independent endocytosis [32,37]. In the case of SNPs, most internalization is via endocytic pathways which could be different with regard to the nature of the surface functionalization and structural properties of SNPs (shown in Scheme 1) [38,39]. Once the silica nanoparticles have overcome the cell membrane barrier, they have to reach the cytoplasm to release the cargoes. To understand the corresponding intracellular pathways would help the design of more efficient drug delivery systems. Fig. 6 presents the CLSM and FCM data for HeLa cells after 3 h of incubation in solutions containing SNPs with various sizes under different conditions. Cells incubated at 37 1C without adding any SNPs were used as a control group. By incubating cells with SNPs at different temperatures, we found that the uptake of silica nanoparticles is higher at 37 1C than at 4 1C. The CLSM for HeLa cells cultivated with SNPs-307.6 at 4 1C does not give any green fluorescence and the relative fluorescence intensity obtained from FCM is 0.6, which is very close to the control group (0.4) and indicates that the uptake of SNPs-307.6 is through Three different inhibitors including sucrose, Filipin III, and Cyt D were used for cell pretreatments to study the endocytosis mechanisms for various SNPs. The resulting effects on fluorescence intensity were examined and compared with those cultivated at 37 1C without any pretreatment. As shown in Fig. 6, the relative fluorescence intensities in FCM for cells with the pretreatment of sucrose decrease from 51.6 to 29.2 for SNPs-55.6, 24.3 to 14.0 for SNPs-167.8, and 1.9 to 1.6 for SNPs-307.6. These results indicate that the cellular uptake of all three SNPs is clathrin dependent [40]. However, with the pretreatment of Filipin III, the relative fluorescence intensity decreases only for SNPs-55.6 and SNPs-167.8, indicating that these two SNPs also follow a caveolin dependent endocytosis pathway. Furthermore, Cyt D does not exert any obvious effect on the cellular uptake of SNPs toward HeLa cells. In summary, the cellular uptake of SNPs toward HeLa cells is size-dependent. The uptake of the largest SNPs (SNPs-307.6) is through an energy-dependent pathway (clathrin dependent and caveolin independent). While for SNPs-167.8, the process is mainly through energy required clathrin and caveolin dependent endocytosis. On the other hand, the uptake of the smallest SNPs (SNPs-55.6) follows not only clathrin and caveolin dependent endocytosis, but also an energy independent pathway. To evaluate the in vitro cytotoxicity, cell viability was examined by standard MTT assay for various SNPs at two different concentrations (10 mg/mL and 100 mg/mL). As presented in Fig. 7, all SNPs show very low cytotoxicity toward HeLa cells at these concentration levels. The results indicate that the biocompatibility of SNPs is very good. The cellular uptake efficiency, mechanism, and cytotoxicity of SNPs with various sizes toward HeLa cells have been investigated. The endocytosis pathway for large SNPs (307.6 nm) is mainly through clathrin coated pits, while smaller SNPs (167.8 nm) can be internalized through clathrin coated vesicles as well as the caveolin dependent pathway. Furthermore, ultra small SNPs (55.6 nm) can partly enter the cells through an energy independent pathway. The cellular uptake efficiency is also sizedependent in the order of 55.64167.84307.6 nm. On the basis of these results, we propose that silica nanoparticles with a diameter of less than 160 nm (which can enter the cell but not likely the nucleus) might be candidates for drug delivery and those SNPs having a diameter of 50 nm or less (which can enter the nucleus) could be carriers of siRNA and DNA for gene therapy. Further studies may facilitate more development of size-defined SNPs as transporters for different purposes. This research was supported by the NSFC (20925517, 21175028), SKLEAC201101, and Texas State University Faculty Development Leave Program. Supplementary data associated with this article can be found in the online version at http://dx.doi.org/. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 61.8, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 48.0, \"mgL\": 10.0, \"viab\": 98.63}, {\"h\": 48.0, \"mgL\": 100.0, \"viab\": 94.894}]}", "role": "assistant" } ]	51.json
[ { "content": "Extract sample context and measurement list. <doc> Experimental and Toxicologic Pathology xxx (2015) xxx–xxx Contents lists available at ScienceDirect Experimental and Toxicologic Pathology journal homepage: www.elsevier.de/etp Sarieh Ghasempoura , Mohammad Ali Shokrgozarb , Roghayeh Ghasempourc , Mohsen Alipoura, * aDepartment of Physiology & Pharmacology, School of Medicine, Zanjan University of Medical Sciences, Zanjan, Iran bNational Cell Bank of Iran, Pasteur Institute of Iran, Tehran, Iran c Renewable Energy of Environment Engineering Department, Faculty of New Science of Technology, University of Tehran, Tehran, Iran Article history: Received 21 April 2015 Received in revised form 7 July 2015 Accepted 13 July 2015 Keywords: Iron oxide nanorods Viability Liver Kidney In recent years, iron oxide nanorods find a lot of applications including drug delivery, cell separation, hyperthermia and magnetic resonance imaging. In this study the cytotoxicity of iron oxide nanorods was evaluated based on mouse fibroblast cell behavior and wistar rat's liver and kidney function. At first for modification, nanorods were added to Dulbecco's modified Eagle's medium (DMEM) which contained a lot of sources of vitamins, amino acids, proteins in Fetal Bovine Serum (FBS). The MTT assay was employed for evaluating the toxic effects of 200 and 400mg/mL modified and non-modified iron oxide nanorods on L929 mouse fibroblast cells in a 24 h period. Changes in cell granularity and size as well as cell cycle were investigated using flow cytometry. Moreover liver and kidney function test and serum iron level measurement were performed 24 h after the injection of modified iron oxide nanorods via the tail peripheral vein of wistar rats. Results indicated that greater concentration of modified iron oxide nanorods had no significant effect on cell viability while greater concentration of non-modified iron oxide nanorods significantly decreased cell viability. Modified iron oxide nanorods did not have significant effects on cell cycle. The results of liver and kidney function tests did not differ significantly while a significant increase in serum iron level was observed. After H&E staining of slices, there were no changes on morphology of rat's kidney and liver cells. This study suggests that short-time use of 200 and 400mg/mL iron oxide nanorods are probably safe. Further studies are needed for investigation of toxic effects of different concentrations, coatings, and exposure time periods of iron oxide nanorods. ã 2015 Elsevier GmbH. All rights reserved. Nanotechnology has many medical and biological applications such as drug delivery, cell separation, hyperthermia, magnetic resonance imaging, cancer diagnosis (Cheng et al., 2007; Gao et al., 2004), synthesis of nanostructure medical devices and drug carriers (Helmus, 2006; Mahmoudi et al., 2009b; Mao et al., 2013). For instance, nanostructures are used as drug carriers for minimizing the side effects and maximizing the effectiveness of chemotherapy agents. Such technology can reduce the therapeutic dose of drugs and it can also protect cells against their side effects (Kommareddy et al., 2005). Human beings mightily strive for protecting their health and improving their life quality. Health and well-being are directly dependent on the health of the most basic parts of human body, E-mail address: (M. Alipour). http://dx.doi.org/ 0940-2993/ã 2015 Elsevier GmbH. All rights reserved. i.e., the cells. The health of body cells is in turn related to the soundness of their condition, proliferation, and evolution. In order to proliferate and survive, each cell goes through different phases called cell cycle. The cell cycle is responsible for cell mitosis and proliferation. It is evident that the strict regulation of mitosis is vital to the health and well-being of all organisms. Accordingly, in order to maintain each complex organism's health, each cell needs to closely regulate its duplication process and have a healthy progression. Uncontrolled duplication due to the adverse effects of drugs, toxins, and harmful rays is considered as the underlying cause of cancer. Cancer is a common life-threatening disease which has a high mortality rate. Accordingly, human knowledge is striving to find effective ways for managing this disease (Gupta and Wells, 2004). Cancers are usually treated by using chemotherapy agents. Besides their therapeutic effects on cancerous cells, these agents have major adverse effects on normal cells of human body. One * Corresponding author. Fax: +98 2433449553. way for minimizing the adverse effects of chemotherapy agents on normal cells is targeted drug delivery to the cancerous tumors. Nanotechnology scientists have used different drug carriers such as iron oxide for targeted drug delivery. Iron nanoparticles, which are found in mineral and organic compounds, are vital to healthy cell growth. In human body, iron nanoparticles are coated with ferritin and stored in form of 5–7 nm particles (Chasteen and Harrison, 1999; Gupta and Wells, 2004). Super paramagnetic iron oxide nanoparticles (SPIONs) are widely used for biological purposes particularly for drug delivery. Bare and coated SPIONs have been investigated in different studies (Mahmoudi et al., 2011b; Mahmoudi et al., 2008). For instance, Mahmoudi et al. (2008, 2011) investigated the cytotoxicity of ferritin-coated SPIONs. However, due to their nano-scale sizes, nanoparticles can also have harmful effects on cells (Mahmoudi et al., 2011a,c). Accordingly, nanoparticles are usually coated with different coatings (An and Su, 2001; Bautista et al., 2005; Kang et al., 2002) such as polyethylene glycol (Kang et al., 2002), chitosan (An and Su, 2001; Khor and Lim, 2003), polyvinyl alcohol (Albornoz and Jacobo, 2006; Osada and Gong, 1998), gold and urea for decreasing their toxicity and increasing their biocompatibility and applicability (Latham et al., 1995). Moreover, super paramagnetic nanoparticles with Lactoferrin and Ceruloplasmin coated have been reported to show cytotoxicity effects on human dermal fibroblasts such as disruption of the cell membrane and disorganized cell cytoskeleton (Gupta and Curtis, 2004). To our knowledge, most of the previous studies have been done on spherical nanoparticles and there are few studies on the cytotoxic effects of rod-shaped ones. The aim of this study were to investigate the cytotoxicity effects of modified and nonmodified iron oxide nanorods on mouse fibroblast cells; effects of modified iron oxide nanorods on cell cycle life of fibroblasts and wistar rats liver and kidney function. In this experimental study, the effects of iron oxide nanorods at 200 and 400mg/mL concentrations were examined on mouse fibroblast cells (L929). According to laboratory facilities, the lowest iron oxide nanorods dose with high precision was 200mg/mL iron oxide nanorods. Moreover for greater certainty double dose (400mg/mL) were also examined. Each dose was compared with its control. The inclusion criterion was being living cells adhering to the plate while the exclusion criterion was having no contamination or morphologic changes. In this study, deionized (DI) water was used for preparation of solutions.Inorder to preventing fromoxidation, solutions deaerated using argon bubbling for 30min. Iron salts with the mole fraction of Fe3 to Fe2 equal to 2:1 were dissolved in 10 mL DI water containing 1.0M HCl. The second solution was obtained by dissolution of 23 g urea in40mL DIwater at roomtemp. Two mentioned solutions were added together and mixed with vigorous stirring. Thereafter, the reaction flask was placed into the hot oil bath and stirring was continued for 8 h at 90 C and 1300 rpm. The particle was washed several times with DI water containing 0.01 M HNO3; the particles were, then, disseminated in DI water containing 0.01 M HNO3 and maintained at 6 C for future use. After dispersion with vortex, nanorods were transferred into falcon tubes. Then Dulbecco's Modified Eagle Medium (DMEM, Gibco, USA) containing 10% Fetal Bovine Serum (FBS, Gibco, USA) was added to the tubes up to nine times more than the volume of nanorods. Tubes were incubated for 24 h. After 24 h, DMEM was refreshed by fresh DMEM. L929 mouse fibroblast cells were supplied by the Pasteur Institute, Tehran, Iran. These cells are stored in DMEM containing 10% FBS and then were incubated at 37.0 C, 90% humidity, and a CO2 concentration of 5%. Totally, 104 numbers of L929 cells were seeded onto each well of a 96-well plate and 100ml of DMEM + 10% FBS culture medium was added and incubated for 24 h for adhering cells to the plate. Then, the wells were divided into three six-well groups. Thereafter, 40ml of non-modified nanorods at a concentration of 200 and 400mg/ mL were added, respectively, to groups one and two. The third group, i.e., the control one, was treated with 40ml of DMEM + 10% FBS. All groups were incubated for 24 h and then (3-(4,5 dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide) assay (MTT assay) was performed on them. MTT assay was repeated for three times. Cell exposure to modified nanorods was performed in the same way as cell exposure to non-modified nanorods. After 24 h of incubation, the plate was removed from incubator. Supernatants were removed under the hood and MTT assay protocol performed as the same way had been written in the leaflet. The concentration of substance solved in isopropanol was calculated by using an ELISA-reader (STAT FAX, USA) at 545 nm. We went through all these phases for both modified and non-modified nanorods. 2.6. Cell preparation for investigating the effect of nanorods on cell cycle Totally, 5 105 cells were seeded onto three 25 mL flasks-5 105 cells onto each flask and 200mg/mL modified nanorods was added to each flask. The control flasks were treated with only the culture medium containing no nanorods. All flasks were incubated for 24 h at 37.0 C, 90% humidity, and a CO2 concentration of 5%. Then, supernatants were removed and cells were washed with PBS. Cells were detached from the plate by using 2–3 mL of trypsin 0.25% (Gibco, USA) solution. The content of each flask was transferred to one falcon. Then, were washed with 5 mL of DMEM + 10% FBS culture medium two times, next supernatants were removed after that were washed by PBS twice. Thereafter, nine mL of ethanol alcohol was added for fixation finally cells were stored at a temperature of –20.0 C for 48 h. These cells can be stored for six months. Cells fixed in alcohol were prepared by washing with PBS two times. Then, supernatants were removed and the collected cells were washed with PBS and then suspended in 1 mL PI/Triton X-100 staining solution with RNase A. Then they were kept either at 37 C for 15 min or at room temperature for 30 min 50ml of PI (Sigma–Aldrich), 20ml of RNaseA (SinnaGen), and one ml of Triton X-100 (Sigma–Aldrich) were added. The volume of suspension was increased to one mL by adding PBS. Falcon tubes containing cells and the aforementioned substances were covered with foil to protect them from light. Then, cell flow cytometry was performed by using Partec (Cyflow, Germany) and cell cycle graphs were analyzed by using the Flowjo software. Totally, 14 wistar rats weighed 240–260 g were recruited to this study. Rats were randomly allocated to either experimental or control groups (seven rats in each group). Rats were numbered and kept in threesome or foursome cages. Each rat was anesthetized by intraperitoneal injection of ketamine (75 mg/kg) and xylazine (100 mg/kg). Then, 200mg/mL modified nanorods was injected via the tail vein by using an insulin syringe having a needle size of 28. Rats in the control group received 0.9% normal saline. One hour after nanorods injection, a 1-mL blood sample was obtained from the heart of each rat through the trans-cardial route and transferred to a test tube. Rats were then returned to their cages. Twenty four hours after the injection of nanorods, rats were re-anesthetized. 1 mL of blood was sampled from the heart and transferred to a test tube. The sample was placed in an ice bag and sent to laboratory for performing liver and kidney function tests (serum alanine aminotransferase (ALT), aspartate transaminase (AST), alkaline phosphatase (ALP), blood urea nitrogen (BUN), and creatinine (Cr)) and measuring serum iron level. These parameters were measured by using COBASINTEGRA kits (Roche, USA). 24 h after injection of modified nanorods to rats, they were perfused transcardially with heparinized saline for 20 min, and then liver and spleen were removed from the body and were put in paraformaldehyde 4%. Then Tissue sections of 5mm thickness were stained with hematoxylin and eosin (H & E). The morphology of the tissues was studies by light microscope. The statistical Package for Social Sciences (SPSS) was used for data analysis. Groups were compared by using the independentsamples t, the one-way analysis of variance (ANOVA), and the Tukey's post-hoc tests. The significance level for all comparisons was set at P < 0.05. Fig. 1 shows the Scanning and Transmission Electron Microscopy (SEM and TEM) images of nanorods (Fig. 1a and b). Urea coated nanorods were rod-shaped with a length and diameter of 15 and 150 nm, respectively. Fig. 2 revealed that the viability of cells which were exposed to non-modified nanorods was significantly lower than the cells which were exposed to modified nanorods (P < 0.001). The findings of this test also showed that compared with the control group, the viability of the cells which were exposed to 200 and 400mg/mL modified and non-modified nanorods was significantly lower (P < 0.05). Moreover, the 200 and the 400mg/mL modified nanorods group did not differ significantly in terms of cell viability. Furthermore, the cells which were exposed to 400mg/mL nonmodified nanorods had a significantly higher viability compared with the cells exposed to 200mg/mL non-modified nanorods (P < 0.05). In flow cytometry images, forward scattering (FSC) and side scattering (SSC), respectively, stand for cell size and cell granularity. Fig. 3 shows the shape and the amount of cell distribution. As Fig. 3 shows, 96.1% of the cells were investigated. In the test group, cells were exposed to 200mg/mL modified nanorods for 24 h. Data showed that 52.5% of these cells remained in the same area as the control group and the remaining 43.6% of cells shifted to either left or upward (Fig. 3). This finding denotes increased cell granularity and decreased cell size. Fig. 4 represents the percentage of cells in different phases of cell cycle. In Flowjo analysis, G2/M, S, G0/G1, and subG0/G1 phases are depicted. The subG0/G1 phase reflects cell apoptosis while the other parameters stand for the different phases of cell cycle. As Fig. 4 illustrates, 3.4% of cells (subG0/G1) among 43.6% of the cells which showed Fig. 2. Viability comparison of L929 which exposed with modified and nonmodified Iron oxide nanorods at 24 h. \|, P < 0.05 for each dose with its own control. *, P < 0.05 for similar dose in different groups. #, P < 0.05 for different doses in same group. Fig. 6 indicates the laboratory findings of the experimental (exposed to 200mg/mL modified nanorods) and the control groups. The results revealed that there was no statistically significant difference between the experimental and the control groups in terms of serum levels of ALT, AST, ALP, BUN, and Cr in one and in 24 h after injection. 24 h after injection, the serum iron level in the experimental group was significantly higher than the first hour. In addition as Fig. 7 shows that in histopathology findings there were no changes on morphology of glomerulus and tubular structure and there were not any inflammation in kidney and also liver cells. The findings in Fig. 2 revealed that compared with modified nanorods, the viability of the cells which were exposed to nonmodified nanorods was significantly lower. It seems after the exposure of nanoparticles to protein-containing media such as biologic fluids or FBS-supplemented culture mediums, proteins are absorbed to the surface of nanoparticles (Mahmoudi et al., 2009a, 2010) and hard corona is formed. The formation of this proteincontaining layer probably changes the pH of the culture medium which in turn results in cell death. However, substitution of the old culture medium, which has experienced pH changes, with a fresh culture medium normalizes the pH. It seems despite the great availability of biologic substances in the fresh culture medium, there is no free location on the surface of nanoparticles for protein binding which accordingly results in the stabilization of pH in the new culture medium (Mahmoudi et al., 2009a, 2010). In this study, we also placed nanorods in a FBS-supplemented culture medium for 24 h and then substituted the old medium with a fresh one. This technique provided a more precise measure of cell death once exposed to nanorods. Mahmoudi et al. (2010) investigated the cytotoxic effects of modified and non-modified spherical nanoparticles. They found that modified nanoparticle was safer than non-modified nanoparticles (Mahmoudi et al., 2010) Moreover, MTT assay findings showed that compared with the corresponding control groups, exposure to all concentrations of modified and non-modified nanorods caused a significant decrease in cell viability which denotes the death of some cells. Cells suffer injury or death immediately after exposure to nanoparticles. This can be attributed to physical or chemical characteristics of nanoparticles which may affect either the cell membrane or the intracellular organelles and cause injury or death. For example, nanoparticlesinduced mitochondrial respiratory chain injury decreases ATP, causes the formation of reactive oxygen species (ROS), and creates oxidative stress which finally increases the probability of cell injury or death (Xia et al., 2006). We also found that an increase in the concentration of non-modified nanorods from 200 to 400mg/ mL caused a significant increase in cell viability. Maybe this significant increment in cell viability once exposed to greater concentrations of nanoparticles can be attributed to iron oxide. Iron is vitalto the health and development of bodily organs. Studies showed that iron deficiency can be detrimental to cell cycle as well as cell cycle proteins such as CDKs and cyclins (Cheng et al., 2007, Helmus, 2006; Mahmoudi et al., 2009b, Mao et al., 2013). Accordingly, the availability of iron would improve the condition for cell growth. In this study, nanorods contained iron. Therefore, increment of cell viability after being exposed to greater concentrations of non-modified nanorods was probably due to the abundance of iron in the medium. Perhaps the iron which has been released from the beneath of urea coating to the culture medium probably improves the condition for cell growth and decreased cell death rate (Huang et al., 2009, Mahmoudi et al., 2009a). However, this finding was not observed for those cells which were exposed to modified nanorods. This is possibly due to the formation of hard corona (Mahmoudi et al., 2009a, 2010). Maybe hard corona acts as an added coating over urea-coated nanorods and prevents iron release from them within a 24 h period, resulting in a higher cell death rate. increased granularity after being exposed to Super Paramagnetic Iron Oxide Nanoparticles (SIPONs). In another study, polyvinyl alcohol-coated SPIONs also caused increased granularity which was confirmed by Comet assay (Mahmoudi et al., 2009b). Cells suffer injury or death immediately after exposure to nanoparticles. This result can be related to physical or chemical characteristics as mentioned previously. As mentioned in Fig. 5, by utilizing flow cytometry we could find cell population in cell cycle phases. As Fig. 4 shows, cell distribution in different phases of cell cycle in the experimental group is similar to the control group. The 0.9% increase and the 0.5% decrease in, respectively, the G0/G1 and the G2/M phases are not significant. However, the 3.1% decrease in phase S deserves further investigation. Cai et al. (2006) also reported cell distribution in G2/M phase following the administration of paclitaxel prodrug (Cai et al., 2006). Iron metabolism degrades the surface of nanoparticles and causes iron release. The released iron ions cause oxidative stress (Chasteen and Harrison, 1999; Gupta and Wells, 2004). In our study, nanorods were iron oxide. We modified iron oxide nanorods by placing them in DMEM and FBS 10% culture medium which caused the formation of hard corona. Accordingly, in vivo studies showed maybe iron could not pass the double-layer coating (i.e., urea and hard corona layers) of nanorods and enter blood in one hour. However, after 24 h, we found a significant difference between the experimental and the control groups in terms of serum iron level (Fig. 6). This finding denotes that after 24 h, the double-layer coating could not prevent iron release from nanorods. According to Mahmoudi et al. (2011a,b,c), the biologic effects of nanoparticles mainly depend on the type of the coating proteins (Mahmoudi et al., 2008). For example, studies have shown that 55% of oleic acid and ploronic-coated nanoparticles aggregate in liver while 20–25% of dextran-coated nanoparticles are excreted in the urine and feces (Weissleder et al., 1989). These differences can be attributed to the type of surface proteins and coatings of nanoparticles (Mahmoudi et al., 2011c). Moreover, the type of surface coating can increase the bioavailability and the safety of nanoparticles. In this regard, Mahmoudi et al. (2009b) found that polyvinyl alcohol and polyethylene glycol increased the safety of iron nanoparticles (Mahmoudi et al., 2009b). In the current study, nanorods were coated with and modified by proteins. Therefore, the safety of nanorods after 24 h may be due to their coatings. Accordingly, modification of nanorods can be considered as a way for increasing their biologic safety. In our study we evaluated kidney and liver function by measuring of ALT, ASTand ALP for liver and BUN and Cr for kidney. The results in Fig. 6 showed there are not any changes in these tests so these confirmed the safety of this dose in this time application. Also in histopathology founding we found health kidney and liver tissues (Fig. 7). Lozano et al. (2012) found that in acute toxicity study of silicon carbide there are no changes on liver and kidney functions by measuring level of ALT, AST, ALP, BUN and Cr. Furthermore they did not see any damages on kidney, liver, stomach and intestines which was referred to Sic (Lozano et al., 2012). Moreover, Park et al. (2009) compared iron oxide nanosphere and nanorods. They found that parameters such as targeting ligand density, kind of target receptor, and nanoparticles physical shape can play an dedicated role in determining the extent of accumulation (Park et al., 2009). Therefore result of safety in our study maybe due to the size, coverage, nanoparticle shape and organs which were examined. So to complete this study it needs more studies. In this study, exposure to modified and non-modified nanorods significantly decreased cell viability. The toxic effects of 200 and 400mg/mL modified nanorods did not differ significantly. However, compared with 200mg/mL concentrations, 400mg/mL nonmodified nanorods significantly increased cell viability. The reason was the improvement of condition for cell growth because of iron release. As mentioned, nanorods were made of iron and additionally we found improvement of viability stat so it is probably because of iron release from under the urea layer. Moreover, 200mg/mL modified nanorods had toxic effect neither on the phases of cell cycle nor on the function of liver and kidney. The only effect of 200mg/mL modified nanorods was a significant increase of serum iron level after 24 h. There were not any changes on liver and kidney functions and they were healthy and safe. These findings maybe suggest that the formation of a proteincontaining layer around nanorods minimizes their toxic effects and enhances their safety in a 24 h period. None. This work was supported by Vice Chancellor for Research and Technology of Zanjan University of Medical Sciences, Zanjan, Iran. (Grant No. A-12-141-3). </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 200.0, \"viab\": 55.577}, {\"h\": 24.0, \"mgL\": 400.0, \"viab\": 72.401}]}", "role": "assistant" } ]	49.json
[ { "content": "Extract sample context and measurement list. <doc> Contents lists available at ScienceDirect journal homepage: www.elsevier.com/locate/toxinvit Ruchi Roy a,d , Vyom Parashar b , L.K.S. Chauhan c , Rishi Shanker a,d,e , Mukul Das a,d , Anurag Tripathi a,⇑ , Premendra Dhar Dwivedi a,d,⇑ a Food, Drug and Chemical Toxicology Group, Indian Institute of Toxicology Research, M.G. Marg, Post Box No. 80, Lucknow 226001, India bNanotechnology Application Centre, Faculty of Science, University of Allahabad, Allahabad 211002, India c Electron Microscopy Facility, CSIR-Indian Institute of Toxicology Research, M.G. Marg, Post Box No. 80, Lucknow 226001, India d Academy of Scientific and Innovative Research (AcSIR), New Delhi, India e Environmental Toxicology Division, CSIR-Indian Institute of Toxicology Research, M.G. Marg, Post Box No. 80, Lucknow 226001, India Article history: Received 5 July 2013 Accepted 11 December 2013 Available online 22 December 2013 Keywords: Zinc oxide nanoparticles Proinflammatory cytokines Macrophages Phagocytosis Endocytosis The inflammatory responses after exposure to zinc oxide nanoparticles (ZNPs) are known, however, the molecular mechanisms and direct consequences of particle uptake are still unclear. Dose and timedependent increase in the uptake of ZNPs by macrophages has been observed by flow cytometry. Macrophages treated with ZNPs showed a significantly enhanced phagocytic activity. Inhibition of different internalization receptors caused a reduction in uptake of ZNPs in macrophages. The strongest inhibition in internalization was observed by blocking clathrin, caveolae and scavenger receptor mediated endocytic pathways. However, FcR and complement receptor-mediated phagocytic pathways also contributed significantly to control. Further, exposure of primary macrophages to ZNPs (2.5 lg/ml) caused (i) significant enhancement of Ras, PI3K, (ii) enhanced phosphorylation and subsequent activation of its downstream signaling pathways via ERK1/2, p38 and JNK MAPKs (iii) overexpression of c-Jun, c-Fos and NF-jB. Our results demonstrate that ZNPs induce the generation of reactive nitrogen species and overexpression of Cox-2, iNOS, pro-inflammatory cytokines (IL-6, IFN-c, TNF-a, IL-17 and regulatory cytokine IL-10) and MAPKs which were found to be inhibited after blocking internalization of ZNPs through caveolae receptor pathway. These results indicate that ZNPs are internalized through caveolae pathway and the inflammatory responses involve PI3K mediated MAPKs signaling cascade. -2013 Elsevier Ltd. All rights reserved. Living beings have been exposed to nanoparticles (NPs) ever since the origin of life on earth. NPs are produced from various natural as well as anthropogenic sources such as, volcanic eruptions, physical and chemical weathering of rocks, diesel exhaust, electroplating, and welding (Rietmeijer and Mackinnon, 1997; Murr et al., 2004; Wigginton et al., 2007; Siegmann et al., 2008). These particles may aggregate to form large sized particles or may persist in air, water, soil and biological systems as such (Beduneau et al., 2009). Therefore, we may assume that living organisms have co-evolved with nanomaterials in the environment and may have developed efficient disposal or toxicity mitigating mechanisms for NPs. Nonetheless, it is also known that as particles approach the nano-scale, they often become more reactive, owing to increase in the surface area and predominance of surface related phenomenon (Monteiller et al., 2007). Importantly, materials that are inert in bulk form may become reactive and toxic in nano-sized form (Nel et al., 2006). For example, even an inert compound like gold tends to elicit a biological response when introduced in nano-form (Goodman et al., 2004). This clearly emphasizes the need for a careful evaluation of the toxicological manifestations rendered by NPs. Zinc oxide nanoparticles (ZNPs) are among the most commonly used nanomaterials, with a wide range of applications in industrial and consumer products. Recently, ZNPs have been shown to specifically target and kill cancer cells and can possibly be developed as an alternative anticancer therapeutic agent (Hanley et al., 2008; Moos et al., 2010). ZNPs have also been reported to have the potential to activate dendritic cells and stimulate the release of proinflammatory cytokines (Heng et al., 2011). In particular the workers in the manufacturing of ZNPs can be subjected to low level chronic exposure to ZNPs via different routes viz. inhalation, ingestion or dermal route. Ryman-Rasmussen et al. (2006), have shown that sub micrometer particles can penetrate even through the outer layers of skin. In the present study, macrophages were chosen as an in vitro model to study the exposure to ZNPs because they are the primary scavenger cells of the body and form the first line of defense in the immune response to foreign materials (Germain and Margulies, 1993). When NPs enter the systemic circulation, they encounter a complex web of immune cells and plasma proteins. The recognition of NPs as non-self by the immune cells may result in a multilevel immune response against the NPs leading to generation of reactive oxygen species (ROS), reactive nitrogen species (RNS) and altered cytokine levels. The oxidative stress response and cytokine milieu in the body is delicately balanced and orchestrated according to the nature of physiological stress, infections and diseases. On exposure to xenobiotics, inflammatory cells produce hyper levels of ROS which may cause airway hyper-responsiveness, sepsis/multiple organ dysfunction, vascular dysfunction and reduced antioxidant levels (Busija et al., 2006; Dong et al., 2005; Hou et al., 2010). Similarly, abnormal levels of cytokines like TNF-a, IL-6, IL-17 and IFN-c have been reported in rheumatoid arthritis, active psoriasis and allergic asthma (Wong et al., 2001; Arican et al., 2005; Gratacos et al., 1994). Macrophages are professional phagocytes, which are capable of efficient uptake of foreign invaders by phagocytosis. Phagocytosis is a particle size-dependent phenomenon which is induced by the interaction of different specialized cell surface receptors with their specific ligands (Haberzettl et al., 2007; Kwiatkowska and Sobota, 1999). Scavenger receptors (SR), mannose receptors (MR), and complement receptors (CR) are some of the specialized receptors with respective specificities (Aderem and Underhill, 1999; Janeway, 1992; Underhill and Ozinsky, 2002). Phagocytosis could further be mediated by the family of Fc receptors (FcR), which recognize immunoglobulin coated particles and complexes (Aderem and Underhill, 1999). Though macrophages do not phagocytose NPs as efficiently as they do for larger particles, but the phagocytic process may get facilitated by the adsorption of opsonins (plasma proteins) onto the particle surface (Owens and Peppas, 2006). Previously, we have reported that ZNPs have the immunomodulatory potential by altering the cytokines like TNF-a, IL-6, IL-1 and IL-17 in macrophages (Roy et al., 2011). Therefore, the aim of the present study was to identify the classic phagocytosis or endocytosis pathways involved in the uptake of ZNPs and then subsequent inflammatory consequences. Internalization of ZNPs was analyzed in ZNPs treated macrophages either in the presence or absence of inhibitors of different potentially involved receptors. Furthermore, we examined the morphological characteristics by TEM, mitogen activated protein kinases (MAPKs) signaling cascade, and the inflammatory responses, such as phagocytic activity, RNS generation and related inflammatory marker such as Cyclooxygenase-2 (Cox-2) and inducible form of Nitric oxide synthases (iNOS) expression in macrophages. Antibodies (CD16/32 and CD11b) were purchased from BD Biosciences. Anti-PI3K, anti-Ras, anti-p-JNK, anti-c-Fos and anti-a-Tubulin were purchased from BD Biosciences, USA. Anti-p-ERK1/2, antip-p38, anti-p-NF-jB, anti-c-Jun and anti-b-actin were purchased from Santa cruz, USA. Anti-Cox-2 and anti-iNOS were obtained from Calbiochem, USA. Phosphate buffered saline (PBS), fetal bovine serum (FBS) and antibiotic-antimycotic solution (10,000 U/ml penicillin, 10 mg/ml streptomycin sulfate) were purchased from Gibco, Invitrogen Cor. (Grand Island, N. Y., USA). Cell culture plastic-wares were obtained from Nunc (Roskilde, Denmark). 100 lg/ml concentration of ZNPs suspension was prepared in RPMI 1640 containing 10% FBS by sonication (Sonics Vibra cell, Sonics and Material Inc., New Town, CT, USA) for 1.5 min at 32 W amplitude followed by the break for 1 min, and subsequently the same process was repeated twice. The nanoparticle suspensions were characterized by the following methods. 2.2.2.1. Transmission electron microscopy (TEM). The microstructure of ZNPs was studied using a TEM and selected-area electron diffraction was carried out on a Technai 30 G2 S-Twin electron microscope operated at 200 kV accelerating voltage. For TEM analysis, samples of ZNPs were prepared by solution-casting onto carbon coated copper grid. 2.2.2.2. Dynamic light scattering (DLS). The average hydrodynamic size, size distribution and zeta potential of ZNPs in suspension were determined by DLS and phase analysis light scattering, using a Zetasizer Nano-ZS equipped with 4.0 mW, 633 nm laser (Model ZEN3600, Malvern Instruments Ltd., Malvern, UK). Inbred strains of female Balb/c mice (8-10 weeks old) were sacrificed according to the guidelines for the care and use of laboratory animals of CSIR-Indian Institute of Toxicology Research, Lucknow, India. Peritoneal exudate cells were collected from the peritoneal cavity of mice by injecting chilled RPMI 1640 medium and added to 96-well cell culture flat bottom plate. After 3 h of incubation in a CO2 incubator (5% CO2) at 37 C, the non-adherent cells were removed by vigorous washing (three times) with warm RPMI 1640 medium. Furthermore, adhered cells were incubated overnight in RPMI 1640 medium supplemented with heat-inactivated FBS (10%), penicillin (100 U/ml) and streptomycin (100 U/ ml) at 37 C in humid air containing 5% CO2 to form macrophage monolayers. More than 95% of the adherent cell populations were macrophages as determined by morphology and non-specific esterase staining. Stock suspension of 100 lg/ml particle concentration in RPMI1640 (supplemented with 10% FBS) was serially diluted to 1, 2.5, 5 and 10 lg/ml and vortexed before distribution in the culture wells. Macrophage monolayers (4 - 105 cells/well) were prepared as described above, and treated with the indicated concentrations of ZNPs for 24 h. The medium was removed and cells were washed with PBS. One tenth of total volume of MTT dye solution was added to each well. The 96-well plates were incubated in a humidified, 5% CO2 incubator at 37 C for 2 h. Next, 100 ll of the solubilization solution was added to the wells and then the plates were kept in the dark for 1 h at room temperature. After brief gentle mixing of contents in each well, the absorbance (OD) of each well was recorded at 570 nm using a 96-well plate reader (BioTek, USA). Macrophages were plated in 12-well culture plate at a concentration of 2 - 106 cells/ml. Cultured cells were treated with NPs at concentrations of 10, 25, 50 or 100 lg/ml for 1 or 2 h. Cells were washed twice with 500 ll of warm 1X PBS and gently scraped from the culture dishes on ice. The uptake of particles was determined by flow cytometer. In all 10,000 events were acquired. For calculation, cell debris was excluded by gating live macrophages. Macrophages (2 - 106 cells/well) were plated in 12-well cell culture plate. After overnight incubation, the cells were exposed to ZNPs (10 lg/ml) for 1 h. After exposure, the media were aspirated and the cells were washed twice with 1x PBS. Cells were then centrifuged and the pellet was fixed with 2.5% glutaraldehyde and 2% paraformaldehyde. After two washes with 0.1 M sodium cacodylate buffer, the pellet was post-fixed in 1% osmium tetraoxide for 3 h. Fixed pellet was then washed and dehydrated through different grades (30–100%) of acetone. The sample was infiltrated with Araldite resin overnight at room temperature and finally embedded in pure resin. The blocks were incubated at 60 C for 72 h. After incubation, ultrathin sections were prepared using Reichert-Jung ultra microtome (Vienna, Austria). The sections were stained with uranyl acetate and Reynold's lead citrate. To avoid confusions of any staining artifacts, the grids were examined before and after staining under TEM (JEM-2100, JEOL Ltd., Tokyo, Japan) at 60 or 80 kV. In this study, we investigated the participation of different classic phagocytosis receptors on the uptake of particles. Macrophages (2 - 106 cells/ml RPMI 1640, 10% FCS) were pre-incubated with different pathway inhibitors: FcR-Ab and CR-Ab at 1:200 dilution (Haberzettl et al., 2008); SR agonist: 400 lg/ml Poly-G for 30 min (Kanno et al., 2007); macropinocytosis, 10 lg/ml Amiloride for 30 min; F-actin polymerisation, 10 lg/ml Cytochalasin D for 30 min (Zhang and Monteiro-Riviere, 2009); clathrin mediated endocytosis, 10 lg/ml Chlorpromazine for 30 min (Zhang and Monteiro-Riviere, 2009); caveolae mediated endocytosis, 5 lg/ml Filipin for 30 min (Schnitzer et al., 1994) and 5 lg/ml Genistein for 1 h followed by treatment with 50 lg/ml particle concentrations for 1 h. The influence of different receptor inhibitors on uptake of ZNPs by macrophages was examined using a flow cytometer. Measurement of IgG binding with FBS opsonised nanoparticles (ZNPs disperse in 10% FBS supplemented RPMI media, incubated at 37 C for 0.5 h) was performed by enzyme linked immunosorbent assay (ELISA). In brief, 100 ll of (5, 50, 100 and 200 lg/ml) NPs were coated onto 96-well micro plates and kept overnight at 4 C and then blocked with 200 ll of 3% BSA. 1:1000 dilution of IgG level was added to each well and incubated for 2 h at 37 C and then washed 3 times with washing buffer (PBS-T). To each well, 100 ll of HRP conjugated goat antimouse IgG antibody (1:1000; Southern Biotech, Birmingham, USA) was added and incubated for 1 h at 37 C. To obviate the interference of NPs with substrate or other reagents, a set having ZNPs only was taken in which antibody was not added. The plates were washed with washing buffer. A 50 ll substrate solution (5 mg ortho-phenylenediamine in 10 ml substrate buffer and 10 ll H2O2) was added to each well and the plate was incubated for 30 min at 37 C in the dark. The reaction was stopped by the addition of 50 ll of stopping solution (5 N H2SO4) and the absorbance was taken at 492 nm in an ELISA plate reader (Biotek, Power Wave XS2). For western blot analysis of phosphatidylinositol 3-kinase (PI3K), Ras, p-ERK1/2, p-p38, p-JNK, p-NF-jB, c-Fos and c-Jun (after 0.5, 3, 6, 12 and 24 h; 2.5 lg/ml of ZNPs alone) the macrophages were harvested, washed with cold phosphate-buffered saline and lysed with ice-cold radio immunoprecipitation assay buffer supplemented with protease and phosphatase inhibitors. p-ERK1/2, p-p38, p-JNK, Cox-2 and iNOS were analyzed in the cell lysates of untreated and treated groups (after 0.5, 3, 6, 12 and 24 h of 2.5 lg/ml of ZNPs alone; 400 lg/ml Poly G or 5 lg/ml Filipin treated cells followed by exposure to ZNPs). Proteins were resolved on 10% sodium dodecyl sulfatepolyacrylamide gel and electrotransferred on polyvinylidene fluoride membranes. The blotted membranes were blocked with 5% BSA in phosphate-buffered saline containing 0.1% Tween 20 (blocking solution) and incubated with the desired antibodies at dilutions mentioned by the manufacturer, followed by incubation with horseradish peroxidase-conjugated secondary antibody (Sigma, St. Louis, USA). Bound secondary antibody was detected by enhanced chemiluminescence using Amersham ECL western blotting detection reagents as described in manufacturers' protocol (Amersham, Fairfield, CT). All the blots were stripped and reprobed with b-actin or b-tubulin to ensure equal loading of protein. For profiling of cytokine production, macrophage monolayer was prepared in 96-well cell culture plate. The cells were treated with 2.5 lg/ml concentration of ZNPs for 24 h. In other groups, cells were pretreated with Filipin (5 lg/ml for 30 min) followed by 2.5 lg/ml ZNPs. Untreated macrophages were taken as control. After the completion of incubation, the plate was centrifuged and 50 ll of the supernatant was collected from each well for the assessment of cytokine IL-6, IL-10, IL-17, TNF-a and IFN-c using the cytometric bead array NHP Th1/Th2 cytokine kit (BD Bioscience, USA) as per manufacturers' instructions. Analysis was done by flow cytometer (FACS CantoTM II, BD BioSciences, San Jose, CA, USA). Data are expressed as mean ± SE. The results were analyzed using analysis of variance (one-way ANOVA). All the statistical analyses were performed using Prism version 5, Graph Pad Software Inc. A value of p < 0.05 was considered as statistically significant. The particle dimensions were established using TEM and DLS. As shown in Fig. 1A, the size of ZNPs measured by TEM was typically around 50 nm and the morphology was roughly spherical. Since serum supplemented medium was used for macrophage culture, the DLS size of ZNPs was also determined. DLS yielded the hydrodynamic size of the particles in solution allowing measurement of particles bearing adsorbed corona of molecules such as adsorbed proteins. Fig. 1B, shows the DLS size distribution with an average diameter of 278.8 nm in culture media supplemented with 10% FBS. The zeta potential of ZNPs was 11.5 mV in complete medium (Fig. 1C). The macrophages were treated with increasing doses (1, 2.5, 5, 10 lg/ml) of ZNPs for 24 h. It was observed that at 1 lg/ml dose, cell viability (>80%) was similar to control (Fig. 2A). However, 2.5, 5 and 10 lg/ml doses showed approx. 40%, 60% and 80% cell death, respectively. Therefore, we have selected 2.5 lg/ml dose (at this dose minimum cell death with cytotoxicity) for the evaluation of cytotoxicity related inflammatory responses. The uptake of ZNPs in cells was assessed by flow cytometry on the basis of side scatter (SSC) intensity which represents the granularity of a cell and forward scatter (FSC) represents the size of a cell. Upto 100 lg/ml of ZNPs incubated for 2 h was found completely non-toxic to macrophages (data not shown). These measurements revealed significantly increased uptake of 50 and 100 lg/ml doses of ZNPs at 1 and 2 h of exposure (Fig. 3A). Therefore, lower dose of 50 lg/ml was used for flow cytometery related experiments. Electron micrographic images demonstrated significant uptake of ZNPs in macrophages (Fig. 3B). Multiple membrane foldings and plasma membrane (pseudopodial) extensions were fused together to engulf the ZNPs. It was also clearly observed that some NPs were adsorbed on the cellular membrane surface. The cytoplasmic region of macrophages showed accumulation of ZNPs within micron-sized vesicles resembling late endosomes and lysosomes. To further address the mechanism of the particle uptake, macrophages were treated with different inhibitors of phagocytic pathways prior to particle incubation. As shown in Fig. 4A and B – reduction in uptake of NPs was found as follows: FcR-Ab (49% ;), CR-Ab (30% ;) and Poly-G agonists (75% ;). To examine the role of proteins opsonisation of ZNPs with IgG in phagocytosis, FBS opsonised particles were incubated with IgG and its binding was measured by ELISA. Binding of NPs to serum IgG was significantly increased at 200 lg/ml (Fig. 4C). For identification of uptake of ZNPs through endocytic mechanisms, macrophages were treated with different inhibitors of endocytic pathway prior to particle incubation (Fig. 5A and B). Reduction in uptake of ZNPs was found as follows: Cytochalasin D (96% ;), Filipin (88% ;), Chlorpromazine (71% ;), Amiloride (66% ;), Genistein (66% ;). To verify the importance of endosome formation for uptake of ZNPs and its implication on actin recruitment to initiate the process, we focused on the specific signaling cascade described by the activation of PI3K and Ras GTPases. Treatment of macrophages with ZNPs (2.5 lg/ml) resulted in the persisted elevation of PI3K protein level from 0.5 h to 3 h, which thereafter started depleting, while Ras increased from 0.5 h to 12 h and down-regulated on 24 h (Fig. 6A). MAPKs are serine/threonine specific kinases that are activated in response to extracellular as well as intracellular stimuli. Hence, it was argued that ZNPs may activate MAPKs signaling in macrophages. As shown in Fig. 6B, ZNPs cause activation of ERK1/2, JNK and p38 MAPKs in macrophages at 0.5 h and which remained enhanced upto 24 h. 3.9. Effect of ZNPs on translocation of transcription factors (NF-jB, c-Jun and c-Fos) in macrophages Several studies have reported that MAPKs play a critical role in the activation of NF-jB by phosphorylating it. It is also reported that MAPKs cause transcriptional activation of c-Jun and c-Fos, the subunits of AP-1 transcription factors. Therefore, the effect of ZNPs on the phosphorylation of NF-jB, and expression of c-Fos and c-Jun proteins in macrophages was studied. The results showed enhancement of phosphorylation of NF-jB at 0.5 h after ZNPs exposure and which remained enhanced up to 24 h (Fig. 6C). The AP-1 subunits, c-Jun and c-Fos, were all found to be overexpressed at 6 h, 12 h and 24 h of exposure to ZNPs (Fig. 6C). Pretreatment of Poly G gradually inhibited the expression of p-ERK1/2 and p-p38, while its effect on p-JNK1/2 was found to be inhibited after 12 h only (Fig. 6B), Filipin had pronounced inhibitory effect on the activation of p-ERK1/2, p-JNK1/2 and p-p38 as compared to Poly G. We observed that macrophages treated with ZNPs had elevated levels of Cox-2 and iNOS protein expressions 0.5 h after exposure and remained elevated upto 24 h in case of Cox-2 and upto 12 h in case of iNOS. Pretreatment of Poly G completely inhibited Cox-2 expression at 24 h, whereas no observable change was noted in the iNOS protein expression from 0.5 h to 24 h (Fig. 7A). Cox-2 expressions inhibited by Filipin was more in comparison to Poly G, whereas iNOS inhibition started from 6 h and was complete by 24 h. A significant enhancement of RNS at 24 h of ZNPs incubation was observed which was inhibited to the control level by Filipin pretreatment but not by Poly G (Fig. 7B). Therefore, further analysis of pro-inflammatory cytokines profiling with Filipin only was performed. Macrophages exposed to ZNPs showed significant enhancement in the levels of pro-inflammatory cytokines IL-6, IFN-c, TNF-a, IL-17 and anti-inflammatory cytokine IL-10 as compared with the untreated control group. Interestingly, on inhibiting nanoparticles internalization with Filipin, there was reduction in IL-6, IFN-c, IL-17 and anti-inflammatory cytokine IL-10 (Fig. 7C). the major functions of macrophages, the aim of the present study was to identify the classic phagocytosis or endocytosis pathways involved in the uptake of ZNPs and to investigate the potential of ZNPs to induce an inflammatory response in macrophages. For this investigation, primary peritoneal macrophages were chosen because they are the first line of defense against foreign assaults, but the extremely small size of nanoparticles may help them to retard or evade the phagocytic process. However, nanoparticles internalization may occur either by direct penetration of the macrophage cell membrane or by specific mechanisms like macropinocytosis (Ruan et al., 2007). The size of ZNPs observed by TEM was 50 nm, while the mean hydrodynamic size obtained by DLS was found to be more in culture medium (278.8 nm) as compared to the size obtained by TEM, this may be due to the coating of ZNPs with proteins in serum. Further elaborating our findings, we studied the uptake pattern of ZNPs by macrophages that may have some bearing upon the functional responses of macrophages. It was observed that the uptake of ZNPs was much higher in macrophages after the first and second hour of exposure to ZNPs at 50 and 100 lg/ml doses. The results obtained from flow cytometry were further validated by TEM where ZNPs were found to get internalized into the macrophages. ZNPs were found to form clusteres in the membrane bound vesicles and multiple membrane foldings were visible. It appears that ZNPs were internalized through the endocytic pathway and were concentrated within micron-sized vesicles resembling late endosomes and lysosomes. We speculated that the reason underlying the internalisation of ZNPs could be attributed to the phagocytic property of macrophages. As expected, the phagocytic activity of macrophages was found to be significantly enhanced on exposure to ZNPs. We further explored the different phagocytic pathways responsible for particle uptake by macrophages using the specific pathway inhibitors. Macrophages express a number of receptors, such as scavenger receptor, FcR and complement receptors that facilitate the recognition of foreign particles (Geier and Celli, 2011). Inhibition of the cellular association of particles with scavenger receptor type A agonist (Poly G) significantly reduced the uptake of ZNPs (75% ;) indicating that scavenger receptor mediated pathway plays an important role in the recognition and internalization of ZNPs by macrophages. Our data further revealed that ZNPs have the tendency to bind with IgG rendering them recognizable by FcR and CR. It was clearly observed that nearly 50% of the ZNPs were found to be internalized through FcR mediated phagocytosis. The current findings also demonstrate the involvement of CR mediated phagocytic pathway in the nanoparticle internalization, though less efficiently (30% ;) as compared to scavenger receptor or FcR mediated uptake pathways. As far as the non-phagocytic process (pinocytosis or endocytosis) is concerned for the internalization of ZNPs, different endocytosis mediated pathways were identified by using specific pathway inhibitors. Endocytosis occurs in normal cells by macropinocytosis, clathrin-mediated endocytosis and caveolae-mediated endocytosis, however, macropinocytosis generally occurs in complementation to clathrin or caveolae mediated endocytosis (Harush-Frenkel et al., 2008). Our results have shown that on pre-treatment of macrophages with Cytochalasin D and Amiloride which interfere with macropinocytosis by preventing membrane ruffling, (Ivanov et al., 2004) and inhibiting Na+ /H+ exchanger isoform 1 activity, the uptake of ZNPs was prominently inhibited (96% ;, 66% ;). As macropinocytosis is an actin cytoskeleton dependent process which aids in the non-specific, non receptor dependent way (Swanson and Watts, 1995). To find out the involvement of clathrin-mediated endocytosis in ZNPs uptake, macrophages were pre-treated with Chlorpromazine which perturbs clathrin-processing (Wang et al., 1993). We have found that clathrin mediated endocytosis has a major contribution in the cellular uptake of ZNPs (71% ;). Our results are in accordance with the findings of Rejman et al. (2004) that clearly showed that internalization of NPs having a diameter below 200 nm occur by clathrin-mediated endocytosis (Rejman et al., 2004). Another important endocytic pathway operational in macrophages is mediated through caveolae, where Filipin and Genistein as inhibitors were used that distort the structure and functions of cholesterol-rich membrane domain required for the maintenance and sealing off, of caveolae plasma membrane (Ros-Baro et al., 2001). Genistien diminished the uptake of ZNPs by 66%, whereas inhibitory effect of Filipin was even more pronounced as it reduced the ZNPs internalization by 88%. Size of the caveolae known to internalize particles or nanoparticles aggregate ranged from 200 nm to 500 nm. As we found that the size of ZNPs aggregates in the FBS Pro-inflammatory effects of ZNPs may have been mediated by the activation of transcription factors (NF-kB, c-Fos and c-Jun) and subsequent pro-inflammatory gene expression. Compared to the untreated cells, ZNPs showed a significant increase in the activation of transcription factors where they further induce Cox-2 and iNOS expression and proinflammatory cytokines such as IL-6, TNFa, IL-17 regulatory cytokine IL-10. Our study suggests that the inflammatory manifestations due to ZNPs may be attributed to their internalization mainly by scavenger receptor and caveolae pathways. Therefore, Poly G and Filipin were investigated for their ability to mitigate the MAPKs and inflammatory responses. Effect of Filipin was more than the Poly G on the key mediators (ERK1/2/, JNK1/2 and p38) and inflammatory markers. Henceforth, these results suggest that caveolae pathway has direct impact on MAPKs and consecutive inflammatory responses. Our results revealed that non-phagocytic caveolae mediated endocytosis mechanism was primarily responsible for uptake of ZNPs by macrophages. In addition to this, ZNPs enhanced the expression of MAPKs, Cox-2, iNOS and production of inflammatory cytokines. All these responses got supressed by inhibiting caveolae pathway. These findings are ought to be of relevant consideration before their potential use in biomedical applications. The authors declare that they have no conflict of interest. The authors wish to thank Dr. K.C. Gupta, Director, CSIR-IITR for his support. Financial support from CSIR Research Grant No. OLP- 009 and Nano-SHE (BSC-0112) are acknowledged. Ruchi Roy is thankful to the University Grants Commission (UGC), New Delhi for the award of Senior Research Fellowship and conveys her gratitude to Academy of Scientific and Innovative Research (AcSIR), New Delhi. We are thankful to Mr. B.D. Bhattacharji, Head, RPBD, CSIR-IITR, for his editorial assistance. This is manuscript No. 3001, allotted by CSIR-IITR. Transparency Documents associated with this article can be found, in the online version, at http://dx.doi.org/. 12.004. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 278.8, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 1.0, \"viab\": 91.342}, {\"h\": 24.0, \"mgL\": 2.5, \"viab\": 61.905}, {\"h\": 24.0, \"mgL\": 5.0, \"viab\": 31.818}, {\"h\": 24.0, \"mgL\": 10.0, \"viab\": 15.152}]}", "role": "assistant" } ]	34.json
[ { "content": "Extract sample context and measurement list. <doc> Title: In vitro cytotoxicity of gold nanorods in A549 cells Author: Ying Tang Yafeng Shen Libin Huang Gaojian Lv Changhai Lei Xiaoyan Fan Fangxing Lin Yuxia Zhang Lihui Wu Yongji Yang Please cite this article as: Tang, Y., Shen, Y., Huang, L., Lv, G., Lei, C., Fan, X., Lin, F., Zhang, Y., Wu, L., Yang, Y.,In vitro cytotoxicity of gold nanorods in A549 cells, Environmental Toxicology and Pharmacology (2015), http://dx.doi.org/ This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. Page 1 of 16 a Department of Biophysics, College of Basic Medical Sciences, Second Military Medical University, Shanghai, 200433, China b Department of Science & Research, Second Military Medical University, Shanghai, 200433, China c College of Material Science and Engineering, East China University of Science and Technology, Shanghai, 200237, China d Department of Cardiothoracic Surgery, Changzheng Hospital, Second Military Medical University, Shanghai, 200433, China Corresponding author. Tel.: +86 21 81870923. E-mail address: (Y. Yang), (L. Wu). 1 All the authors contributed equally to this work. Accepted ManuscriptGold nanoparticles, which have unique physicochemical characteristics, are being used for an increasingly wide range of applications in biomedical research. In this study, gold nanorods (width of 25 nm, length of 52 nm) were found to be internalized by A549 cells and were primarily localized in the lysosomes and membranous vesicles. The integrity of the membranes of A549 cells exposed to gold nanorods for 4 h was damaged, as indicated by laser scanning confocal microscopy (LSCM). Increased lactate dehydrogenase (LDH) leakage and decreased cell viability further indicated the concentration-dependent cytotoxicity of the gold nanorods to the A549 cells. Reactive oxygen species (ROS) production was induced in the A549 cells by the gold nanorods, and this effect was positively correlated with the concentration of the gold nanorods. The results of this study indicated that exposure to gold nanorods caused dose-dependent cytotoxicity in A549 cells and that oxidative stress may be the main factor causing cytotoxicity. Keywords: Gold nanorods, A549 cells, Laser scanning confocal microscope (LSCM), Transmission electron microscope (TEM), Cytotoxicity Nanoparticles are generally defined as particles that have at least one dimension less than 100 nm. Because of their small sizes and unusual structures, these materials have many unique physical and chemical properties, including small-size effects, high specific surface area, high reactivity, and quantum effects. In recent years, gold nanoparticles have gradually become a hot topic of research due to their shape, the simplicity of the control of their synthesis, large surface area, high chemical stability, good biocompatibility and their strong absorption and scattering of light. Gold nanoparticles are widely used in various fields including biosensors, cell and tissue biology, diagnosis, drug delivery, imaging and tumor hyperthermia ((Aibo liu and ye, 2013; Catherine. Murphy et al., 2008; Daniel and Astruc, 2004; Huang et al., 2008; Murphy et al., 2008). At present, a number of in vitro studies on the cytotoxicity of gold nanoparticles have been performed, but the results are inconclusive. Some have reported that the cytotoxicity of gold nanoparticles was related to their size, with smaller gold nanoparticles demonstrating greater cytotoxicity (Pan et al., 2007; Yen et al., 2009). Other studies have proposed that rod-shaped gold nanoparticles have much greater cytotoxicity than spherical ones of the same size, although there are insufficient data regarding the toxicity of nanomaterials of different shapes (Roy et al., 2014). In HeLa cells, exposure to gold nanorods (width and length of 40 × 14 nm and 74 × 14 nm) for 6 h resulted in greater internalization than was observed for spherical nanoparticles (14, 30, 50, 74, and 100 nm sizes)(B. Devika Chithrani et al., 2006; Chithrani and Chan, 2007). One study also indicated that gold nanorods were cytotoxic to most mammalian cells, but the reaction of cells to gold nanorods depended primarily on the cell type rather than on the gold nanorod size and cellular uptake(Chuang et al., 2013). Accepted ManuscriptAlthough the potential toxic impact of gold nanoparticles was controversial, gold nanoparticles were widely used in nanomedicine, such as diagnosis, drug delivery and tumor therapy. In the present work, A549 cells were used to assess cytotoxicity of gold nanorods. The A549 tumor-cell line, a human lung adenocarcinoma derived by explants-culture of lung cancer tissue from a Caucasian male, was established in 1972 and had characteristic features of Type II cells of the pulmonary epithelium, including lamellar bodies (Lieber et al., 1976). This cell line has been widely used in in vitro cytotoxicity and drug metabolism studies(Beer et al., 2012; Chuang et al., 2013; Foldbjerg et al., 2011; Kimberly A. Foster et al., 1998; Uboldi et al., 2009). We aimed to explore the interaction between the gold nanorods and A549 cells using transmission electron microscopy (TEM), laser scanning confocal microscopy (LSCM) and molecular biology techniques. The results showed a dose-dependent cytotoxic effect of gold nanorods on A549 cells and suggested that the oxidative stress induced by the nanoparticles may be the main cause of toxicity. Gold nanorods were obtained from Sigma Aldrich (St. Louis, MO, USA). The gold nanorods were 22.5-27.5 nm in width and 42-52 nm in length. The chemicals and reagents were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China) unless otherwise specified. Dulbecco Modified Eagle Medium (DMEM) was obtained from Thermo Fisher Scientific (Shanghai, China). Fetal bovine serum (FBS) was purchased from Hangzhou Sijiqing Biological Engineering Materials Co., Ltd. Trypsin was purchased from Shanghai Boguang Biology Technology Co. Ltd (Shanghai, China). Laboratory equipment was purchased from Thermo Fisher Scientific (Shanghai, China). Cell counting kit-8 (CCK-8) were purchased from Dojindo Chemical Co. Ltd. (Japan); LDH assay kit, ROS assay kits and GSH assay kit were purchased from Beyotime Biotechnology Research Institute (Jiangsu, China); and the fluorescent dyes, propidium iodide (PI) and Hoechst 33342 were purchased from Sigma Aldrich (St. Louis, MO, USA). The morphology and physicochemical properties of the gold nanorods were examined by transmission electron microscopy (H-7650, Hitachi, Japan) and energy dispersive spectrometry (EDS). The formation of gold nanorods was further confirmed by energy dispersive X-ray spectroscopy (EDX). Accepted ManuscriptA 15 μg/ml interim suspension of the gold nanorods was prepared using serum-free DMEM medium by agitation with a vortex mixer for 2 minutes. The suspension was then dispersed using ultrasonication (power of 300 W, temperature between 25 and 30 °C) for 10 min, repeated 3 times. Finally, the gold nanorod suspension was diluted to 2.5 μg/ml, 5μg/ml, 7.5 μg/ml, 10 μg/ml, 12.5 μg/ml, and 15 μg/ml using serum-free DMEM culture medium. The suspension of gold nanorods was freshly prepared for each experiment. A549 cells, a human lung adenocarcinoma cell line, were obtained from the China Center for Type Culture Collection (CCTCC) (Wuhan University, China). After recovery, the A549 cells were cultured in a 5% CO2 atmosphere using DMEM culture medium containing 10% fetal bovine serum, 100 IU/ml penicillin and 100 μg/ml streptomycin at 37 °C. When A549 cells reached a density of approximately 70%, they were subcultured with 0.25% trypsin. The cell viability was detected by cell counting kit-8 according to the manufacturer's instruction. A549 cells (5×104 A549 cells/ml; 100 μl/well in DMEM) were seeded in 96-well plates. After culture overnight, the cells were incubated with gold nanorods for 4 h, and then 10 μl of 2-(2-methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium monosodium salt (WST-8) was added for 45 min. The absorbance was measured at 450 nm using a microplate reader (Synergy2, Bio-tek, USA). The mean absorbance value for each gold nanorod concentrations was divided by the mean value of control cells and thus presented as a percentage of control (control treated as 100%). A549 cells (5×104 A549 cells/ml; 180 μl/well in DMEM) were seeded in 96-well plates with 5 wells for each concentration. After 24 h, the negative and blank control groups of the cells and those to which the gold nanorods were added were incubated for 4 h. The supernatant was collected by centrifugation at 400 × g for 5 minutes in a 96-well-plate centrifuge, and 120 μl of the supernatant from each well was combined with 60 μl of the LDH working fluid. After incubating the plate for 30 minutes at room temperature, the absorbance was measured at 490 nm using the microplate reader. A549 cells were seeded onto coverslips in 12-well cell-culture plates and incubated with the gold nanorods. The samples were stained with 5 μg/ml Hoeschst 33342 and 3 μg/ml PI fluorescent dyes at room temperature in the dark for 10 min. After washing with 0.1 M PBS, the A549 cells were examined using LSCM (Leica TCS SP2, Leica Inc, Germany). 2.7. Transmission electron microscopy A549 cells were incubated with various concentrations of gold nanorods for 4 h, collected by scraping, and then fixed overnight at 4 °C with a fixative mixture consisting of 2% paraformaldehyde and 2% glutaraldehyde. The cells were then centrifuged at 850 × g for 5 min and washed three times with 0.1 M PBS. The cells were then coagulated with glutaraldehyde and a small amount of blood plasma. The cell masses were cut into small pieces (1 mm3 ), washed with 0.1 M PBS, and then post-fixed with 1% osmium acid for 2 h. The samples were then dehydrated in a graded series of ethanol and acetone and embedded in Epon812 epoxy resin. Ultrathin sections (thickness, 70 nm) were obtained using an ultramicrotome (UC6, Leica Inc, Germany) and stained with uranyl acetate and lead nitrate. The ultrastructure images of A549 cells were obtained using a TEM (H-7650, Hitachi, Japan). Accepted ManuscriptThe A549 cells were seeded into 12-well culture plates at a cell density of approximately 105 cells and incubated with different concentrations of the gold nanorods. Cells incubated with serum-free DMEM were used as a control group. After a 4-h exposure, the cells were washed three times with PBS and then incubated with 10 μM 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA) for 20 min in the dark at 37 °C. After washing with 0.1 M PBS, the samples were mounted in 50% glycerol and observed using LSCM. The detection parameters were an excitation wavelength of 488 nm and an emission wavelength were 500-550 nm. All experimental data are presented as the mean ± standard deviation (SD) of three separate experiments and analyzed by one-way ANOVA followed by Dunnett's t-test for comparisons between groups. P < 0.05 was considered a significant difference. The gold nanorods were characterized by TEM and EDAX (Fig. 1). The TEM results (Fig. 1A) showed that the gold nanorods were rod-like, having widths ranging between 23 and 26 nm and lengths ranging from 35 to 58 nm. The gold nanorods were well dispersed in the serum-free medium. The spectroscopic analysis of the gold nanorods by energy dispersive X-ray spectroscopy (Fig. 1B) showed the characteristics peaks for gold (Au). Internalization of the gold nanorods in A549 cells was investigated using TEM imaging. The results (Fig. 2) demonstrated the uptake and intracellular distribution of the gold nanorods in the A549 cells. Gold nanorods mainly enter A549 cells by phagocytosis (Fig. 2A). In the cells, the gold nanorods were present in the cytosol and in membranous vesicles and lysosomes in the form of aggregates (Fig. 2B-2D), in addition to small single particles in the lysosomes and cytoplasm (Fig. 2B). Gold nanorods were not observed in the nucleus. Most of the mitochondria in the gold nanorod-exposed A549 cells were swollen to various degrees, and the number of lysosomes increased significantly (Fig. 2B-2D). In the group of A549 cells treated with 10 μg/ml gold nanorods, the cells were observed to be disintegrated (Fig. 2D). The number of necrotic cells increased significantly with increasing concentrations of the gold nanorods. A549 cells were treated with a range of concentrations of gold nanorods (2.5 μg/ml, 5 μg/ml, 7.5 μg/ml, 10 μg/ml, 12.5 μg/ml, or 15 μg/ml) for 4 h. The gold nanorods caused a dose-dependent decrease in the viability of the A549 cells (Fig. 3A). The threshold concentration that induced significant toxicity was 5 μg/ml. With further increases in the concentration of the gold nanorods, the cell viability decreased significantly. In the 7.5 μg/ml group, the cell viability was only approximately 20% of that of the control group. Fig. 3 (A) Viability of A549 cells incubated with different concentrations of gold nanorods for 4 h, (B) LDH leakage from A549 cells exposed to various concentrations of gold nanorods for 4 h. The data are shown as the means ± SD values obtained from three separate experiments. All treated groups showed statistically significant differences from the control group (p<0.05, p<0.01). Lactate dehydrogenase (LDH) is abundant in the cytoplasm and cannot pass through the normal cell membrane. When the cells are damaged or dead, LDH is released into the extracellular medium. An elevated level of LDH leakage thus reflects cell membrane damage. The LDH leakage results for the A549 cells exposed to the various concentrations of gold nanorods are shown in Fig. 3B. The results showed that gold nanorods were able to induce LDH leakage of A549 cell in all groups in a dose-dependent manner. The amount of LDH leakage from the 5 μg/ml group was twice that of the 2.5 μg/ml group. Accepted ManuscriptThe cell membrane integrity was also examined using the PI-Hoechst 33342 double-fluorescent staining method. Fluorescence images of the A549 cells were obtained using LSCM, as shown in Fig. 4. Hoechst 33342 (blue in Fig. 4) stains the DNA of living cells independent of the presence of cell membrane injury. PI (red in Fig. 4), a non-membrane-permeable dye, can only enter the cell when the cell membrane is damaged. In the control group (Fig. 4A), only blue fluorescence was visible, which indicated that cell membrane integrity was maintained. In the 5 μg/ml group, in addition to the blue stain, a small amount of red fluorescence (Fig. 4B) was visible, indicating that the cell membranes suffered a certain degree of damage. In 15 μg/ml group, the brightness of the red fluorescence increased significantly (Fig. 4D), indicating that the membranes of most of the cells were more seriously damaged. Fig. 4 Laser scanning confocal microscopic images of A549 cells exposed to various concentrations of gold nanorods for 4 h. The cells were stained with PI (red) and Hoechst 33342 (blue). (A) control group, (B) 5 μg/ml group, (C) 7.5 μg/ml group, (D) 15 μg/ml group. 3.5. Cellular oxidative stress level Oxidative stress induced by gold nanorods in the A549 cells was examined using LSCM, as shown in Fig. 5. ROS generation is indicated by DCF fluorescence (green). The results showed that the brightness of green fluorescence in the A549 cells was clearly enhanced in cells treated with 10 μg/ml or higher concentrations, indicating that ROS generation increased. The ROS generation in the A549 cells increased with the further increases in the gold nanorod concentrations (Fig. 6). Fig. 5. Laser scanning confocal microscopic images of ROS generation in A549 cells exposed to various concentrations of gold nanorods for 4 h. (A) control group, (B) 5 μg/ml group, (C) 10 μg/ml group, (D) 15 μg/ml group. Fig. 6. The fluorescence intensity of ROS generation of A549 cells exposed to various concentrations of gold nanorods for 4 h. The data are expressed as the means ± SD. p<0.01: the response of the experimental group was significantly different from that of the control group. As the potential for exposure to nanomaterials in daily life increases, there has been an increased level of concern regarding their potential to cause harm and a demand for a full evaluation of their safety (Brumfiel.G, 2003; Serice.R.F, 2003). Gold nanoparticles are widely used in biomedical fields, including drug delivery, diagnostics and therapy. The potential toxicity of gold nanoparticles to organisms has not yet been fully evaluated. In recent years, although some studies have reported in vivo and in vitro toxicity and biological effects of gold nanoparticles in animals and various cell lines, the results have been inconsistent. Accepted ManuscriptIn this study, we demonstrated that gold nanorods were readily internalized into A549 cells and damaged the ultrastructure of these cells, and the gold nanorods were taken up by these cells primarily by phagocytosis (Fig. 2A). Earlier studies suggested that gold nanoparticles could be easily transported into A549 cells through endocytosis, and the manner of transport was not associated with nanoparticle size (Uboldi et al., 2009). There are a lot of studies about existence forms and locations of the gold nanoparticles in the cells, but have no unified opinion. Our study showed that the gold nanorods in A549 cells existed in two forms, single particles and aggregates, although most were present as aggregates. The gold nanorods were mainly located in membranous vesicles and occasionally in the cytoplasm and lysosomes. Similarly, some other studies reported that gold nanoparticles were freely dispersed in the cytoplasm (Rothen-Rutishauser et al., 2007). However, our results are inconsistent with some previous studies that proposed that the gold nanoparticles were internalized into cells and stored in phagocytic vesicles (Connor et al., 2005; Pernodet et al., 2006; Takenaka et al., 2006; Uboldi et al., 2009; Verma et al., 2008). In addition, swollen mitochondria and increasing numbers of lysosomes showed that the gold nanoparticles were toxic to the A549 cells. When the concentration of the gold nanorods was increased to 10 μg/ml and above, the number of necrotic cells increased significantly. The data from the LDH release experiments showed that at an even lower concentration (2.5 μg/ml), the gold nanorods caused apparent LDH leakage. At concentrations of the nanogold particles greater than 7.5 μg/ml, PI (red fluorescence) staining was observed in the A549 cells. PI, a non-membrane-permeable fluorescence dye, is observed within the cells only when the cell membranes are damaged and their permeability is increased. The results of the experiment with this dye indicated that the gold nanorods have dose-dependent toxic effects on the A549 cells. The CCK-8 assay is utilized to study cell toxicity and primarily detects mitochondrial damage. The results from this assay showed that the cell viability after exposure to 7.5 μg/ml gold nanorods decreased significantly to approximately 20% of that of the control cells. This result was consistent with the observations of changes in the cellular ultrastructure, which further confirmed the threshold concentration of gold nanorods that caused overt toxicity was 7.5 μg/ml. However, Accepted ManuscriptPrevious studies proposed that the underlying mechanism of gold nanoparticle-induced cytotoxicity is oxidative stress and that ROS generation is the major factor inducing cell toxicity. Paino et al. suggested that gold nanoparticles could cause obvious increases in ROS generation in HepG2 cells and PBMC and that this effect was closely correlated with enhanced cytoxicity (Paino et al., 2012). The results of this study are consistent with our studies in which the DCF measurements showed that A549 cells exposed to 5 μg/ml gold nanorods generated only a small amount of ROS. When the concentration of gold nanorods was increased to 10 μg/ml, the amount of ROS induced in the A549 cells exceeded the normal ROS generation required for cell survival, thus imposing toxic effects to the cells. However, a different conclusion was reached in another recent study(Aueviriyavit et al., 2014). Aueviriyavit et al. suggested that gold nanoparticles had no significant effect on ROS generation or GSH levels in colonic adenocarcinoma cells (Caco-2), and the gold nanoparticles did not induce oxidative stress in the cells. In this paper, the interaction between gold nanoparticles having a length-width ratio of 2 (42-52 nm: 22.5-27.5 nm) and A549 cells was studied. Gold nanorods could be internalized by the A549 cells and were primarily localized in the lysosomes and membranous vesicles but were also found scattered in the cytoplasm. In A549 cells exposed to gold nanorods, various levels of mitochondrial swelling were observed, and the numbers of lysosomes increased significantly. The decrease in cell viability and the damage to the cell membrane integrity indicated a dose-dependent toxicity of the gold nanorods to the A549 cells. High levels of ROS production suggested that oxidative stress may be the main factor causing cytotoxicity. The Authors report no conflicts of interest. 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Verma, A., Uzun, O., Hu, Y., Hu, Y., Han, H.S., Watson, N., Chen, S., Irvine, D.J., Stellacci, F., 2008. Surface-structure-regulated cell-membrane penetration by monolayer-protected nanoparticles. Nature materials 7, 588-595. Yen, H.J., Hsu, S.H., Tsai, C.L., 2009. Cytotoxicity and immunological response of gold and silver nanoparticles of different sizes. Small 5, 1553-1561. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 4.0, \"mgL\": 2.5, \"viab\": 96.783}, {\"h\": 4.0, \"mgL\": 5.0, \"viab\": 69.437}, {\"h\": 4.0, \"mgL\": 7.5, \"viab\": 21.716}, {\"h\": 4.0, \"mgL\": 10.0, \"viab\": 15.013}, {\"h\": 4.0, \"mgL\": 12.5, \"viab\": 7.7748}, {\"h\": 4.0, \"mgL\": 15.0, \"viab\": 6.9705}]}", "role": "assistant" } ]	29.json
[ { "content": "Extract sample context and measurement list. <doc> 1 Department of Chemical and Materials Engineering, University of Dayton, Dayton, OH 45469, USA 2 Molecular Bioeffects Branch, Bioeffects Division, Airmen Systems Directorate, Wright-Patterson Air Force Base, OH 45433, USA 3 Integrative Science and Engineering Center, University of Dayton, Dayton, OH 45469, USA Correspondence should be addressed to Kristen K. Comfort; Received 18 July 2018; Accepted 17 September 2018; Published 1 October 2018 Guest Editor: Nikhat J. Siddiqi 2018 Claudia J. Labrador-Rached et al. Tis is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. Due to their distinctive physicochemical properties, platinum nanoparticles (PtNPs) have emerged as a material of interest for a number of biomedical therapeutics. However, in some instances NP exposure has been correlated to health and safety concerns, including cytotoxicity, activation of cellular stress, and modifcation to normal cell functionality. As PtNPs have induced diferential cellular responses in vitro, the goal of this study was to further characterize the behavior and toxicological potential of PtNPs within a HepG2 liver model. Tis study identifed that a high PtNP dosage induced HepG2 cytotoxicity. However, lower, subtoxic PtNP concentrations were able to elicit multiple stress responses, secretion of proinfammatory cytokines, and modulation of insulin-like growth factor-1 dependent signal transduction. Taken together, this work suggests that PtNPs would not be overtly toxic for acute exposures, but sustained cellular interactions might produce long term health consequences. In recent years, nanoparticles (NPs) have emerged as a major research thrust, with extensive resources and eforts focused on generating and characterizing a library of unique nanosized materials. Due to their enhanced surface area to volume ratio, NPs display diferential behavior from their bulk counterparts, such as augmented catalytic potential, distinctive plasmonic signatures, and enhanced transport capabilities [1]. Te unique physicochemical properties and behaviors associated with NPs have led to their incorporation into hundreds of consumer, medical, and industrial applications [2]. Tis surge in NP usage is accompanied by a corresponding rise in human exposure to these novel materials. Te feld of nanotoxicology was founded to explore the safety of NPs following introduction within biological systems [3]. Years of investigations have revealed the mounting challenge facing this feld, as observed bioresponses are dependent upon unique, NP-specifc physicochemical properties, including size, surface moiety, core composition, and morphology [4]. Documented nanotoxicological responses vary widely and can include cellular death, activation of numerous stress responses, genotoxicity, and developmental abnormalities in vivo [5, 6]. Beyond traditional toxicological events, NPs have been shown to modify basal cell functionality, even in the absence of cytotoxicity, including the activation of infammatory and immune responses, modifcation to gene transcription patterns, and modulation of signal transduction pathways following external stimulation [7, 8]. Many noble metals NPs, including platinum (PtNPs), have gained considerable attention due to their unique plasmonic and catalytic potentials, making them attractive for emerging nano-based developments [9]. PtNPs, which were originally employed as a fuel catalyst additive, are now being researched as a core material for numerous biomedical applications. Potential therapeutics span the vast biomedical feld and include diagnostic mediators, contrast agents for imaging, medical implants, drug delivery vehicles, and photothermal therapy compounds [10–12]. While other metal NPs, such as gold and silver, have undergone extensive nanotoxicological investigations, limited studies have thoroughly explored the safety of PtNPs. More importantly, there appears to be some discrepancy between published PtNP safety analyses. Previous works demonstrated either a biocompatibility or a great degree of cytotoxicity following PtNP exposure, depending on the cell model [13, 14]. Interestingly, PtNPs are well documented as reactive oxygen species (ROS) scavengers and have been shown to reduce intracellular reactive oxygen in the presence of other stressors [15, 16]. However, in some cases, PtNPs elicited negative bioresponses including the activation of cellular stress, as well as DNA damage and genotoxicity efects in vitro [17–19]. In a zebrafsh model, PtNP addition resulted in developmental alterations and a concentration dependent drop in heart rate, demonstrating that PtNPdependent efects translated into in vivo models [20]. Te existence of conficting reports regarding PtNP-induced reactions suggests that additional evaluations are required to better elucidate biological responses and ensure the safety of PtNP-derived applications. Te goal of this study was to enhance the current state of knowledge regarding cellular response following PtNP introduction in vitro. Tis work employed the human liver, HepG2, cell model as PtNPs have been shown to accumulate in the liver, making it a relevant nanotoxicological target [17]. Select endpoints included both traditional toxicological evaluations and biological responses frequently overlooked in safety evaluations, such as the activation of an infammatory response and modulation of signal transduction pathways. Tis work demonstrated that PtNPs induced cytotoxicity and activated several stress pathways in a dose-dependent fashion. Moreover, PtNP exposure elicited a mild proinfammatory response and augmented phosphorylation of the critical signaling protein Akt, suggesting that chronic PtNP exposure could lead to long term health concerns. 2.1. Liver Cell Culture. HepG2, the human liver model utilized in this study, was purchased from American Type Culture Collection. Te cells were maintained in tissue culture treated petri dishes and grown in RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin.Te HepG2s were housed in a humidifed incubator maintained at 37∘ C and 5% carbon dioxide. 2.2. Platinum Nanoparticle Characterization. Te citrate coated, 70 nm PtNPs were purchased from nanoComposix in concentrated solution form.Te PtNP stock was stored at 4∘ C in the dark to minimize modifcations to physicochemical properties over time. Primary particle size and morphology were verifed using transmission electron microscopy (TEM) on a Hitachi H-7600 microscope. Te spectral signature of the PtNPs was visualized through ultraviolet visible (UV-Vis) spectroscopy on a Synergy 4 BioTek microplate reader. For the remainder of PtNP characterization experiments, the particles were diluted to 25 g/mL in either water or media immediately before assessment. Extent of PtNP agglomeration was quantifed via dynamic light scattering (DLS) on an Anton Paar Litesizer 500. Te surface charge of the particles was determined via a zeta potential analysis, also carried out on the Anton Paar Litesizer. For ionic dissolution experimentation, the PtNP samples were incubated at 37∘ C for 24 hours, followed by NP removal via centrifugation at 10,000 rpm for 15 minutes. Te ion containing supernatant was collected and analyzed for platinum content via inductively coupled plasma optical emission spectrometry (ICP-OES) on a Termo Fisher iCAP 7200. 2.3. HepG2 Viability Assessment. HepG2 cells were seeded into 96-well plates at a density of 3x104 cells per well and returned to the incubator. Te following day the cells were washed and dosed with the denoted PtNP concentration or fresh media as a negative control. Afer a 24-hour exposure the cells were washed and the viability was determined using the CellTiter 96 AQueous One Solution Cell Proliferation Assay (MTS) from Promega, in accordance with the manufacturer's protocol. Te untreated cells served as a control to determine percent viability of experimental conditions. 2.4. Intracellular Stress Evaluations. Stress levels within the HepG2 cells were assessed via two endpoints: reactive oxygen species (ROS) and actin levels. For both metrics HepG2 cells were seeded into black 96-well plates at a density of 3x104 cells per well and returned to the incubator overnight. For ROS analysis, the cells were washed and incubated with DCFH-DA probe (Termo Fisher Scientifc), washed again, and exposed to the stated PtNP conditions for 24 hours. For actin evaluation, the HepG2s were exposed to the denoted conditions for 24 hours, washed, and fxed with 4% paraformaldehyde. Te cells were then probed for actin using Alexa Fluor 555-phalloidin (Termo Fisher Scientifc), in accordance with the manufacturer recommendations. Afer proper treatment and staining, both ROS and actin levels were quantifed via fuorescence analysis on a Synergy 4 BioTek microplate reader. PtNP dosed conditions were normalized against an untreated, negative control. 2.5. HepG2 Inflammatory Response to PtNPs. Evaluation of proinfammatory cytokine production was used to assess activation of the HepG2 infammatory response following PtNP exposure. Specifcally this study examined the secretion of interleukin- (IL-) 1, IL-6, IL-8, and tumor necrosis factor- (TNF-) . For these evaluations, HepG2 cells were plated into 6-well plates at a density of 1x106 cells per well and exposed to the stated conditions the following day. As FBS contains cytokines, PtNP-dosed and control experiments were carried out in serum free media. Afer an exposure duration of 24 hours the supernatants were collected and underwent cytokine analysis using protein specifc ELISA kits from Termo Fisher Scientifc, in accordance with the manufacturer's directions. 2.6. Akt and Erk Signaling Activation. Te liver cells were seeded at a density of 1x106 cells per well and grown overnight in a 6-well plate. Te cells were washed and dosed with the stated PtNP conditions in serum free media for a duration of 24 hours. Te HepG2 cells were then stimulated with 10 ng/mL of insulin-like growth factor- (IGF-) 1 for 1 hour at 37∘ C in order to activate the PI3K/Akt and Ras/Erk pathways. Te cells were then washed and lysed in a nondenaturing lysis bufer containing phosphatase and protease inhibitors (Cell Signaling Technology). Te phosphorylation levels of Akt and Erk were determined using ELISA kits from Cell Signaling Technology, which targeted Ser473 and Tr202/Tyr204, respectively. Phosphorylation levels were normalized by the total amount of the same protein, determined via ELISA analysis (Cell Signaling Technology). 2.7. Statistical Analysis. All experiments were performed in three independent trials for the purpose of carrying out statistical analysis. All data is expressed as mean ± the standard error of the mean. Graphpad Prism was used to run a one-way ANOVA for statistical analysis, with a p value threshold set to 0.05, and an asterisk (∗) indicating signifcance from untreated controls. 3.1. PtNP Characterization. Tis study focused on evaluating the safety of PtNPs, as they have emerged as particles of interest spanning both the medical and commercial sectors [10–12, 21]. To date, however, conficting reports exist regarding the safety of PtNPs within biological systems, thereby warranting additional evaluation. Prior to cellular exposure it was essential that the PtNPs underwent a standard battery of characterization assessments, as nanotoxicological potential has been correlated to the unique physicochemical properties of each experimental particle set [4]. TEM imaging, shown in Figure 1(a), verifed the spherical morphology of the 70 nm PtNPs utilized in this study. Using multiple images, a primary particle size of 68.3 ± 3.5 nm was calculated, demonstrating the uniformity of the PtNP stock. Next, the spectral profle of the PtNPs was produced using UV-Vis analysis (Figure 1(b)). Tis spectral signature displayed a single sharp peak, at approximately 260 nm, aligning with previous reports [22]. Moreover, the presence of a single peak confrmed the uniformity of the PtNPs, in agreement with TEM analysis. Next, the PtNPs underwent characterization for behavioral trends following dispersion in water or cell culture media, thereby capturing both stock and exposure fuid conditions. As all nanomaterials will agglomerate to some degree in solution, it was necessary to verify that extensive agglomeration, and a loss of particle stability, did not occur. As shown in Table 1, the PtNPs displayed minimal agglomeration in water, with a slight increase noted within media, verifying particle stability. Moreover, the small polydispersion index (PdI) values indicated that the particle set was monodisperse. Te increase in agglomerate size and PdI associated with cell culture media was due to the formation of a protein corona, which instantaneously forms around NP agglomerates within protein rich environments [23]. Zeta potential analysis was run to determine the surface charge of the particles (Table 1). In water the PtNPs displayed a negative charge, in accordance with the citrate coating. However, following dispersion in media, the PtNP surface charge was increased to approximately -10 mV, which aligns with the formation of the protein corona and the innate protein charge [23]. Finally, the percent of ionic dissolution was quantifed over a 24-hour time period, to determine the rate of platinum ion production that occurred from the PtNP surface. Quantifying ion production has emerged as a critical characterization assessment, as the secretion of metallic ions has been correlated to cytotoxicity, in particular with silver NPs [24]. As seen in Table 1, the rate of ionic dissolution within water was relatively minimal and was further reduced following dispersion in media due to the presence of a protein coating. 3.2. HepG2 Viability and Stress Activation. Following PtNP characterization, a dose-dependent cytotoxicity analysis was carried out. HepG2 cells were specifcally selected for this study owing to the considerations that PtNPs have been shown to accumulate in the liver and that this cell line has become a model for nanotoxicological investigations [13, 25]. As seen in Figure 2, PtNP exposure resulted in a dosedependent decrease in HepG2 viability. At dosages of 25 g/mL or less, no toxicity was identifed. However, at the high exposure concentration an approximate 25% cytotoxic response transpired. One of the areas of conficting reports following PtNP exposure is the activation of intracellular stress, with studies identifying both pro- and anti-oxidant efects [14–17]. Terefore, the next goal was to characterize the HepG2 stress response following exposure to the experimental, 70 nm PtNPs (Figure 3). First, intracellular ROS levels were monitored, as its production is a documented precursor for apoptosis and a known cellular response following NP exposure [26, 27]. As shown in Figure 3(a), the PtNPs induced ROS production in a dose-dependent fashion, with a substantial response associated with the 25 g/mL condition. In addition to ROS, actin expression was evaluated as a metric for cellular stress. Actin becomes disorganized and infamed during stress, making an increase in its expression directly proportional to cellular distress [28]. Following PtNP exposure, the actin expression displayed a dose-dependent increase, closely mirroring the ROS results (Figure 3(b)). Taken together, these fndings demonstrated that citrate coated, 70 nm PtNPs were able to activate a signifcant stress response in HepG2 liver cells, even in the absence of cytotoxicity. 3.3. Inflammatory Response to PtNP Exposure. Beyond activation of stress, NP exposure has been shown to trigger infammatory and immune responses in mammalian cells [5, 29]. Assessing infammatory activation is not a traditional nanotoxicological outcome; however, sustained infammation can introduce serious health implications, including heart disease, hypertension, and even cancer [30]. To assess PtNP-induced infammation in HepG2 cells, the secreted levels of IL-1, IL-6, IL-8, and TNF- were measured, which are early markers of an active infammatory response [31]. Te cytokine production levels following a 24-hour exposure to PtNPs are provided in Figure 4. Secretion of both IL-1 and IL-8 increased, in a dose-dependent manner, with the 25 g/mL dosage raising production approximately 50% over the untreated control. PtNP upregulated TNF- formation, approximately 25%, but appeared to be independent of exposure concentration. Tis study identifed no signifcant changes to IL-6 levels following PtNP exposure. 3.4. Modified IGF-1 Signaling. Lastly the ability of PtNPs to disrupt signal transduction was explored, as nanomaterials have previously been shown to modulate signaling pathways following growth factor stimulation [7, 17]. Signal transduction is a foundational aspect of cellular functionality as its activation and regulation control numerous outcomes including proliferation, migration, and survival. Moreover, unregulated signaling has been correlated to severe health concerns including cancer, respiratory conditions, and neurological diseases [32]. IGF-1 is a known growth factor for HepG2 cells, with ligand-receptor binding inducing the critical signaling pathways of PI3K/Akt and Ras/Erk [33]. Activation of these pathways was quantifed by evaluating phosphorylation levels of Akt and Erk, respectively (Figure 5), which are recognized central players in cellular functionality. As seen in Figure 5(a), both experimental PtNP dosages augmented Akt activation, in a dose-dependent fashion. Following 25 g/mL PtNP exposure, Akt phosphorylation was increased by approximately 50% over untreated controls. On the contrary, Erk phosphorylation levels were the same for all exposure conditions, suggesting that IGF-1 dependent activation of the Ras/Erk cascade was not a PtNP cellular target in HepG2 cells. 3.5. Implications of ese Findings. Te goal of this study was to further elucidate the biological response of liver cells following exposure to PtNPs. To date the literature has produced conficting reports regarding PtNP safety [13– 16]. Te most likely explanation is that nanotoxicological outcomes are dependent upon both cell type and the unique physicochemical properties of the experimental particles. Tis work utilized a human liver model, HepG2, as all NPs, including PtNPs, are known to accumulate in the liver [13]. Characterization assessments demonstrated that the citrate coated 70 nm PtNPs were of high quality, uniform in size, and stable within solution (Figure 1 and Table 1). Te work presented here focused on two distinct areas: (1) traditional toxicological endpoints and (2) evaluation of cellular functionality in the absence of toxicity. A pictorial summary of all observed HepG2 responses following PtNP exposure is shown in Figure 6. Initial eforts focused on traditional toxicological endpoints, with the frst evaluation metric exploring PtNPinduced cytotoxicity of HepG2 cells (Figure 2). A dosedependent toxicity was identifed, with a 25% loss of viability only occurring at the highest experimental PtNP concentration, 100 g/mL. While this observed HepG2 cytotoxicity is signifcant, PtNPs are not renowned for their toxicity potential versus some other core compositions, such as silver. In a recent study, 10 nm silver NPs were correlated with an approximate 60% loss of human lung cell viability following a 24-hour exposure to 25 g/mL concentration [7]. A previous study compared the direct toxicity of several NPs and identifed that silver, copper, and zinc were some of the most potent NP cores, inducing exceptionally high degrees of cytotoxicity [34], helping to highlight that PtNPs can cause cell death, but not to the same degree as other experimental nanomaterials. However, at lower, application-relevant dosages, full viability was maintained, suggesting that a direct toxicity response would not transpire if PtNPs were incorporated into products and therapeutics. However, even in the absence of cytotoxicity, PtNPs were found to induce intracellular stress pathways including ROS (Figure 3). ROS is a prooxidant cellular response to NPs and is a well-documented precursor of additional cellular distress, activation of infammatory and immune responses, and genetic modifcations [27, 35]. One known downstream response of ROS production is actin infammation and disorganization [28]. Tis response was verifed following PtNP exposure, with a 50% augmentation of actin expression identifed.Terefore, this study confrmed that at low, subtoxic concentrations, PtNPs were still capable of inducing multiple stress responses. Of greater interest was the investigation into whether PtNP exposure could disrupt normal HepG2 functionality. In addition to being a nontraditional toxicity evaluation, the examination of infammatory responses and signaling activation would provide insight into whether PtNPs could alter basal cell function and activity. Following PtNP exposure, secretion of IL-1, IL-8, and TNF- was all upregulated (Figure 4), indicating an active infammatory response. Cytokine production is a defence mechanism that is triggered following extensive cellular stress [30, 31] and therefore aligns with the ROS results. Moreover, this study identifed that PtNP incubation interfered with the HepG2 response to IGF-1 stimulation, increasing Akt phosphorylation levels. Tis result aligns with a previous report which identifed that 5.8 nm PtNPs increased activation of multiple signaling pathways, including PI3K/Akt, Ras/Erk, and Ras/JNK, in keratinocytes [17]. Tis work, which examined the efect of 70 nm PtNPs on a liver model, did not uncover any changes to the Ras/Erk cascade, further supporting the supposition that PtNP-dependent biological responses are reliant upon both cell type and NP physicochemical properties. Taken together, this work identifed that PtNP exposure augmented both infammation and signal transduction within HepG2 cells. Combined these responses could lead to serious health concerns if chronic exposure occurred, as alterations to these functional processes have been correlated to numerous diseases, including cancer, heart disease, and neurological disorders [30, 32, 35]. Moreover, increased Akt phosphorylation can trigger a deeper infammatory response [36], thereby establishing a synergistic intracellular reaction. Tis study explored the safety of 70 nm, citrate coated PtNPs within a liver mammalian model. Te PtNPs induced a potent cytotoxic response only at high dosages, well above incidental exposure levels. However, at lower concentrations, even in the absence of cellular death, PtNP exposure elicited a signifcant stress response, notably a marked increase in ROS production. Aligning with the stress activation, a strong infammatory reaction was observed, through both actin infammation and augmented secretion of IL-1, IL-8, and TNF-. Following exposure, it was elucidated that PtNPs disrupted basal cellular functionality as assessed via modulation to the PI3K/Akt signaling pathway following IGF-1 stimulation. Terefore, while low level exposure to PtNPs may not induce direct toxicity, the combined presence of intracellular stress, active infammatory responses, and upregulated signal transduction have the potential to introduce long term health hazards. However, as PtNPdependent biological responses are correlated to cell type and physicochemical parameters, further investigations are required prior to their regulated utilization in consumer applications or as biomedical therapeutics. Te data used to support the fndings of this study are included within the article. Te authors declare that there is no confict of interest regarding the publication of this paper. Claudia J. Labrador-Rached received funding from the University of Dayton Honors Program and the Verhof Family Research Award. Rebecca T. Browning and Kristen K. Comfort were funded in part from the Defense Associated Graduate Student Innovators (DAGSI). Rebecca T. Browning was also supported from the Henry Jackson Foundation. by SIRT1 through NF-B deacetylation in aseptic loosening,\" International Journal of Nanomedicine, vol. 12, pp. 3617–3636, 2017. www.hindawi.com Volume 2018 Medicinal Chemistry International Journal of www.hindawi.com Volume 2018 Hindawi Hindawi Pain Hindawi Hindawi BioMed http://www.hindawi.com Volume 2013 Hindawi Publishing Corporation Hindawi www.hindawi.com www.hindawi.com Volume 2018 Hindawi Emergency Medicine International Hindawi www.hindawi.com Volume 2018 Addiction Journal of Hindawi www.hindawi.com Volume 2018 Research and Treatment www.hindawi.com Volume 2018 Research and Treatment Hindawi www.hindawi.com Volume 2018 www.hindawi.com Volume 2018 Research International Research International Hindawi www.hindawi.com Volume 2018 www.hindawi.com Volume 2018 Hindawi Journal of Hindawi Hindawi Submit your manuscripts at www.hindawi.com Hindawi www.hindawi.com Volume 2018 Anesthesiology Research and Practice www.hindawi.com Volume 2018 www.hindawi.com Volume 2018 www.hindawi.com Volume 2018 Hindawi </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": 112.4, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 5.0, \"viab\": 100.0}, {\"h\": 24.0, \"mgL\": 25.0, \"viab\": 94.475}, {\"h\": 24.0, \"mgL\": 100.0, \"viab\": 68.646}]}", "role": "assistant" } ]	24.json
[ { "content": "Extract sample context and measurement list. <doc> Article Department of Stem Cell and Regenerative Biotechnology, Konkuk University, Seoul 05029, Korea; (M.J.); (M.-H.K.) * Correspondence: (S.G.); (J.-H.K.); Tel.: +82-450-0581 (S.G.); +82-450-3687 (J.-H.K.) Received: 18 April 2019; Accepted: 8 May 2019; Published: 10 May 2019 Abstract: Generally, platinum nanoparticles (PtNPs) are considered non-toxic; however, toxicity depends on the size, dose, and physico-chemical properties of materials. Owing to unique physico-chemical properties, PtNPs have emerged as a material of interest for several biomedical applications, particularly therapeutics. The adverse effect of PtNPs on the human monocytic cell line (THP-1) is not well-established and remains elusive. Exposure to PtNPs may trigger oxidative stress and eventually lead to inflammation. To further understand the toxicological properties of PtNPs, we studied the effect of biologically synthesized ultra-small PtNPs on cytotoxicity, genotoxicity, and proinflammatory responses in the human monocytic cell line (THP-1). Our observations clearly indicated that PtNPs induce cytotoxicity in a dose-dependent manner by reducing cell viability and proliferation. The cytotoxicity of THP-1 cells correlated with an increase in the leakage of lactate dehydrogenase, generation of reactive oxygen species, and production of malondialdehyde, nitric oxide, and carbonylated proteins. The involvement of mitochondria in cytotoxicity and genotoxicity was confirmed by loss of mitochondrial membrane potential, lower ATP level, and upregulation of proapoptotic and downregulation of antiapoptotic genes. Decreases in the levels of antioxidants such as reduced glutathione (GSH), oxidized glutathione (GSH: GSSG), glutathione peroxidase (GPx), superoxide dismutase (SOD), catalase (CAT), and thioredoxin (TRX) were indicative of oxidative stress. Apoptosis was confirmed with the significant upregulation of key apoptosis-regulating genes. Oxidative DNA damage was confirmed by the increase in the levels of 8-oxodG and 8-oxoG and upregulation of DNA damage and repair genes. Finally, the proinflammatory responses to PtNPs was determined by assessing the levels of multiple cytokines such as interleukin-1β (IL-1β), IL-6, IL-8, tumor necrosis factor-α (TNF-α), granulocyte-macrophage colony-stimulating factor (GM-CSF), and monocyte chemoattractant protein 1 (MCP-1). All the cytokines were significantly upregulated in a dose-dependent manner. Collectively, these observations suggest that THP-1 cells were vulnerable to biologically synthesized ultra-small PtNPs. Keywords: platinum nanoparticles; oxidative stress; genotoxicity; proinflammatory response; cytokines; apoptosis; DNA damage Owing to their catalytic and unique surface-related physicochemical properties, including high surface area, platinum nanoparticles (PtNPs) have garnered considerable interest in the automotive sector, chemical industry, and biomedical device manufacturing industry [1–3]. In addition, Pt catalysis is used in hydrogenation reactions, oxidation during organic acid production, isomerization, and dehydrogenation, as well as in the oxidation of carbon monoxide [4–6]. Application of PtNPs in industry and biomedicine depends on several factors such as their size, shape and elemental composition, electronic surface structure, presence of capping agent, dispersion state, solubility, and cell type [3,7,8]. Several methods have been developed for PtNPs synthesis; however, these methods have several disadvantages, such as unnecessary toxic effects of nanoparticles due to the presence of various potential harmful contaminants or hazardous chemicals, production of high amounts of waste, and uncontrolled size. Among the physical, chemical, and biological methods of synthesis, the biological method is most viable and ecofriendly. For instance, PtNPs are synthesized using bacteria [9], cyanobacteria [10,11], seaweeds [12], fungi [13], plants [14], and honey [15]. Although biological system-assisted synthesis of PtNPs appears to be simple, non-toxic, and cost-effective, it has certain limitations for biomedical applications; for example, the presence of endotoxins and fragments of biological materials with unwanted biological activity necessitate expensive and time-consuming purification procedures, including downstream processing. Furthermore, generation of biocompatible and stable products is laborious. Therefore, new and efficient methods utilizing suitable reducing agents are required to produce nanomaterials of controllable size, shape, stability, catalytic properties, and surface functionalization within a short period [1]. For example, PtNPs encapsulated within apoferritin exhibited high efficiency of cellular uptake and caused lower level of membrane damage. To avoid unnecessary impurities in nanoparticle preparation, Gurunathan and colleagues used various purified biomolecules such as saponin [16], quercetin [17], resveratrol [18], lycopene [19], R-phycoerythrin [20], and luteolin [21] during the synthesis of various nanoparticles, including silver, palladium, graphene, and gold. Generally, nanoparticles induce toxicity in human cells via oxidative stress, DNA damage, cell cycle arrest, and mitochondrial dysfunction. Biological activity of nanomaterials is dependent on their size, shape, polarity, porosity, and surface chemistry, type of capping agents, surfactants, and organic solvents used for synthesis. In addition, pristine PtNPs or PtNPs coated with cell-compatible materials exhibit less toxicity. PtNPs in the size range of 5–10 nm did not induce cytotoxicity, oxidative stress, and cell death in A549 and HaCaT cells [22]. Similarly, PtNPs synthesized using citrate as the reducing agent, with particle sizes ranging from 5 to 20 nm, showed significant cytocompatibility, and all the internalized particles accumulated in endosomal/lysosomal vesicles [23]. Functionalization of nanomaterials plays a significant role in cellular toxicity. Human peripheral blood monocytes (THP-1) treated with polystyrene nanoparticles functionalized with carbonyl group (PS-COOH) exhibited no cytotoxicity, whereas polystyrene nanoparticles functionalized with PS-NH2 not only inhibited THP-1 proliferation, but also induced apoptosis in THP-1 cells [24]. The THP-1 cell line was obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA). The cells were maintained as described in Section 2.3. All cells were cultured in 75 cm2 tissue culture flasks (Sigma-Aldrich, St. Louis, MO, USA) at 37 ◦C in the presence of 5% CO2 and 95% relative humidity. Dried and hydrated hexachloroplantinic acid (H2PtCl6·6H2O) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Penicillin-streptomycin, trypsin-EDTA, Roswell Park Memorial Institute (RPMI)-1640 cell culture medium, fetal calf serum (FCS), and antibiotic-anti-mycotic reagents were obtained from Life Technologies/Gibco (Grand Island, NY, USA). The in vitro toxicology assay kit was purchased from Sigma-Aldrich. The reagent kits for the measurement of malondialdehyde (MDA), protein carbonyl content, and antioxidant assay were purchased from Sigma-Aldrich. All other chemicals were purchased from Sigma-Aldrich unless otherwise stated. PtNPs were synthesized by reducing PtCl6 2- ions into PtNPs by mixing 10 mL of 1 mg/mL apigenin with 90 mL of 1 mM aqueous H2PtCl6.6H2O. The mixture was maintained at 100 ◦C (on a hotplate) in a sealed flask to avoid evaporation for 1 h, as temperature catalyzes the reduction process. For control experiments, identical amounts of platinum solution and apigenin were maintained separately under the same reaction conditions. The reduced platinum solution was sonicated for 10 min to separate platinum nanomaterials from the biomolecules. After sonication, the solution was filtrated with a 0.2-μm syringe filter. The reduced platinum metal was purified by repeated centrifugation at 5000× g for 30 min and the pellets were washed with distilled water to remove the impurities. Scheme 1 indicates the various steps involved in synthesis and purification of the PtNPs. Purified PtNPs were characterized using various analytical techniques such as UV-vis spectroscopy, X-ray diffraction (XRD), Fourier transform infrared spectroscopy (FTIR), dynamic light scattering (DLS), scanning electron microscopy (SEM), and transmission electron microcopy. THP-1 cells were cultured in RPMI-1640 cell culture medium supplemented with 10% FCS, 2 mM L-glutamine, 10 mM HEPES, 1 mM pyruvate, 100 U/mL penicillin, and 0.1 mg/mL streptomycin (Sigma-Aldrich). The cells were sub-cultured usually twice a week with 1 × 106 viable cells/mL and incubated at 37 ◦C in a 5% CO2 atmosphere. The medium was replaced the next day with 100 μL fresh 1 media and the cells were incubated for 24 h prior to PtNP exposure. Experiments were performed in 96-, 24-, and 12-well plates and 100-mm cell culture dishes, as required. Cells were treated with various concentrations of PtNPs or the required dose of PtNPs. Cell viability was measured using cell counting kit-8 (CCK-8; CK04-01l; Dojindo Laboratories, Kumamoto, Japan). Briefly, THP-1 cells were plated in 96-well flat-bottom culture plates containing various concentrations of PtNPs. After 24 h culture at 37 ◦C in a humidified 5% CO2 incubator, the CCK-8 solution (10 μL) was added to each well, and the plate was incubated for another 2 h at 37 ◦C. The absorbance was measured at 450 nm using a microplate reader (Multiskan FC; Thermo Fisher Scientific, Waltham, MA, USA). Cell proliferation was determined according to manufacturer's instructions (Sigma-Aldrich, St. Louis, MO, USA). Cells were incubated with various concentrations of PtNPs for 24 h; the BrdU labeling solution was added to the culture medium 2 h before the end of the incubation. The cells were fixed and the level of incorporated BrdU was determined using the BrdU enzyme-linked immunosorbent assay (ELISA) kit (Roche, Basel, Switzerland) following the manufacturer's instructions. Proliferation of the untreated cells at 0 h was considered 100%. The membrane integrity of THP-1 cells was evaluated using a lactate dehyrogenase (LDH) cytotoxicity detection kit. Briefly, the cells were exposed to various concentrations of PtNPs for 24 h. Subsequently, 100 μL of cell-free supernatant from each well was transferred in triplicate into the wells of a 96-well plate, and 100 μL of the LDH reaction mixture was added to each well. After 3 h of incubation under standard conditions, the optical density of the final solution was determined at a wavelength of 490 nm using a microplate reader. Cell mortality was evaluated using the trypan blue assay as described previously [34]. THP-1 cells were plated in the wells of 6-well plates (1 × 105 cells per well) and incubated for 24 h with various concentrations of PtNPs. Cells cultured in medium without PtNPs were used as controls. After 24 h, the cells were detached using 300 μL trypsin–EDTA solution, and both adherent and suspended cells were collected. The mixture of the supernatant and detached cells was centrifuged at 1200 rpm for 5 min. The pellet was mixed with 700 μL trypan blue solution and dispersed. After 5 min of staining, the cells were counted using a cytometer. The viable cells were unstained and the dead cells were stained blue. Three independent experiments were performed in triplicate. The mean and standard deviation were calculated. Cell proliferation is expressed as the percentage of viable cells relative to the appropriate control. ROS were estimated as described previously [35]. THP-1 cells were seeded into wells of 24-well plates at a density of 5 × 104 cells per well and cultured for 24 h. After washing twice with phosphate-buffered saline (PBS), fresh medium containing various concentrations of PtNPs was added and incubated for 24 h. The cells were supplemented with 20 μM DCFH2-DA, and incubation was continued for 30 min at 37 ◦C. The cells were rinsed with PBS, 2 mL PBS was added to each well, and the fluorescence intensity was determined using a Gemini EM spectrofluorometer (Molecular Devices, Sunnyvale, CA, USA) at an excitation wavelength of 485 nm and an emission wavelength of 530 nm. MDA levels were determined using a thiobarbituric acid-reactive substances assay as previously described with suitable modifications. NO production was quantified spectrophotometrically using Griess reagent (Sigma-Aldrich). The absorbance was measured at 540 nm and nitrite concentration was determined using a calibration curve prepared with sodium nitrite as the standard [36]. Carbonylated protein content was measured according to a previously described method [37]. 8-oxo-dG content was determined as described previously [35,39] and also per the manufacturer's instructions (Trevigen, Gaithersburg, MD, USA). Total RNA was extracted from the LNCaP cells treated with various concentrations of PtNPs for 24 h using the PicoPure RNA isolation kit (Arcturus Bioscience, Mountain View, CA, USA). Samples were prepared according to the manufacturer's instructions. RT-qPCR was conducted using a Vill7 device (Applied Biosystems, Foster City, CA, USA) and SYBR Green as the double-stranded DNA-specific fluorescent dye (Applied Biosystems). Target gene expression levels were normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) expression, which was unaffected by the treatment. The sequences of the PCR primers are shown in Supplemental Table S1. THP-1 cells were treated with PtNPs for 24 h and cytokine levels were determined using multianalyte inflammatory cytokine ELISA array (Qiagen, Valencia, CA, USA). Sample values were normalized to control values and are displayed as fold change over control. Independent experiments were repeated at least three times such that data are represented as mean ± standard deviation (SD) for all duplicates within an individual experiment. Data were analyzed using the t-test, multivariate analysis, or one-way analysis of variance (ANOVA), followed by Tukey's test for multiple comparisons to determine the differences between groups denoted by an asterisk. In this study, we investigated the cellular responses to PtNPs in THP-1 cells. To demonstrate the cytotoxic, genotoxic, and pro-inflammatory effects of PtNPs, we first confirmed bio-reduction of Pt ions using phenolic compounds such as apigenin as the reducing and stabilizing agent. Bio-reduction was visually verified by initial color change from pale yellow to dark brown, and finally by the appearance of a brownish-black precipitate. The intensity of the color depended on the period of incubation. The color change is due to the reaction between hexachloroplatinic acid and apigenin and indicates the formation of platinum nanoparticles [40]. The entire reaction was completed within 2 h due to the presence of pure phenolic compounds in the solution. The single sharp peak at 300 nm was attributed to the excitations of surface plasmon vibrations, indicating the synthesis of PtNPs (Figure 1A), which is in agreement with the results of previous reports [41,42]. Several studies have reported that metal nanoparticles can be synthesized using biomolecules as reducing and stabilizing agents. For example, silver and copper nanoparticles were prepared using ascorbic acid as the reducing and stabilizing agent [43,44]. In addition to color change, the formation of PtNPs was confirmed by a continuous absorption spectrum in the range of 300–600 nm in a UV-visible spectrophotometer (Figure 1A). The surface plasmon resonance of PtNPs synthesized using apigenin was observed at 302 nm. PtNPs synthesized using Ajwa and Barni dates exhibited resonance at 321 and 329 nm, respectively; however, the absorption spectra are dependent on the size and morphology of the particles. The purity and crystal structure of the synthesized PtNPs were assessed using XRD. As shown in Figure 1B, the broad reflections of the synthesized nanoparticles indicated the nanocrystalline nature of the powder. Broad diffraction peaks of the XRD pattern were observed at 2θ = 40.6, 48.4, and 68.1, corresponding to reflections (111), (200), and (220), respectively, which are consistent with the face centered cubic (fcc) structure of platinum and can be assigned to JCPDS Card 04-0802, demonstrating the presence of crystalline. Shah et al. [45] reported synthesis of PtNPs with an XRD pattern similar to that reported in this study, using a low-temperature economically feasible method. The sharpening of the peaks clearly indicated that the particles were in the ultra-small size range. The size of the platinum nanocrystals as estimated from the full width at half maximum of the (111) Pt peak using the Scherrer formula was 1–2 nm. The XRD pattern clearly illustrates that the PtNPs formed using this present synthesis method were crystalline in nature. The bio-reduction and capping of PtNPs was confirmed using FTIR. The typical characteristic bands at 1660 and 3350 cm−1 represented the C−N (amide) and O−H bonds, respectively (Figure 1C). The amino acids show the possible interaction of PtNPs and stretching modes for C=O and OH groups. The bio-reduction and capping of PtNPs using water-soluble apigenin was evident from the disappearance of a strong band at 1760 cm−1 . Furthermore, the presence of a strong band at 1200 cm−1 indicated the presence of C−O bond. The binding of biomolecules with platinum nanoparticles via amino acid groups was evident from the shift in NH frequency from 2900 cm−1 . Al-Radadi et al. [46] demonstrated synthesis of PtNPs using date fruit extracts, which contain polyphenols, organic acids, amino acids, and antioxidants such as apigenin and glutathione, which are responsible for the reduction of Pt salts. Accurate determination of the size of synthesized nanoparticles is crucial for developing nanoscale technologies and biological studies, as size affects many of the physical and chemical properties of these materials. DLS has been widely used for determining particle size [47]. As shown in Figure 1D, the average size of the particles was 2 nm, which is in accordance with the data obtained from TEM images. The results obtained from DLS exhibited unimodal distribution of nanoparticles with maximum percentage intensity of ~2 nm. The surface morphology of PtNPs was determined using SEM. As shown in Figure 1E, SEM showed individual PtNPs as well as aggregates. The PtNPs aggregated into large irregular structures with no well-defined morphology and all the particles possessed almost identical morphology. Finally, the structure and size of the PtNPs were analyzed using TEM. The TEM images showed that PtNPs had a uniform spherical shape and regular size, and were well dispersed with a narrow size distribution from 1.0 to 2.0 nm. DLS showed that the particles were 2 nm in diameter (Figure 1F). TEM exhibits significantly smaller size of the particles and good correlation with DLS, as TEM involves imaging of dry nanoparticles, whereas DLS is a measure of the hydrodynamic diameter. First, to evaluate the cytotoxic effects of PtNPs on THP-1 cells, we performed a mitochondria-based viability assay (CCK-8) using different doses of PtNPs. The higher doses of PtNPs (100–150 μg/mL) had a drastic effect on THP-11 viability, reducing cell viability to 60 ± 6.5% after 24 h of exposure (p < 0.05). This tendency continued, reaching 90% ± 7% after 24 h (Figure 2A). In contrast, compared to the control cells, low doses of PtNPs (25 μg/mL) did not significantly affect cell viability during the first 24 h of exposure (p > 0.05). Viability of THP-1 cells decreased by 90–95% after 24 h of treatment. Comparison of cell viability kinetics of THP-1 indicated that all PtNP treatments reduced the total number of viable cells after 24 h of treatment. Results of the CCK-8 assay showed that PtNPs reduced the number of viable cells in a dose-dependent manner in the concentration range of 25–150 μg/mL. Next, we evaluated the potential anti-proliferative effect of PtNPs on THP-1 cells. For this purpose, we cultured THP-1 cells in the presence of different concentrations of PtNPs. Treatment of THP-1 with PtNPs (25–150 μg/mL) for 24 h reduced cell proliferation to 25% at 50 μg/mL and to 85% at 150 μg/mL. These observations indicate that PtNPs exhibit dose-dependent cytotoxicity of THP-1 cells by inhibiting cell proliferation (Figure 2B). Yogesh et al. [48] reported in vitro and in vivo activities of biosynthesized PtNPs against human lung cancer, and PtNPs inhibited tumor growth by 66% in Severe combined immunodeficient mice SCID mice. Recently, Gurunathan et al. [38] reported that graphene oxide and reduced graphene oxide reduced the viability and proliferation of THP-1 cells. Changes in cell morphology by PtNPs were assessed using a digital microscope (Figure 3). Compared to the control, cells treated with various concentrations of PtNPs displayed significant changes in morphology, including loss of uniformity and remarkable shrinkage around the cell clusters. (Figure 3). The consistency in results obtained for cell viability, cell proliferation, and LDH leakage assay, trypan blue staining, and digital microscopy after PtNP treatments indicated that PtNPs affect cell survival and eventually cause cytotoxicity. Similarly, U87 glioblastoma cells treated with various concentrations of PtNPs showed characteristic cell death morphology with long branched protrusions. Compared to that in the control, PtNPs treatment reduced the cell number and the length of the cell protrusions [49]. At higher concentrations, the cells exposed to PtNPs clearly show membrane blebbing and loss of plasma membrane integrity. Collectively, the results suggest that PtNPs can alter cell morphology. Chronic exposure of human bronchial cells BEAS-2B cells with crystalline silica Min-U-Sil 5 induces epithelial-to-mesenchymal transition via argpyrimidine-modified Hsp70, miR-21 and SMAD signaling [50]. Figure 3. PtNPs altered the morphology of THP-1 cells. The effect of PtNPs on cell morphology was determined after 24 h exposure to different concentrations of PtNPs (25–150 μg/mL) using an optical microscope. The results are expressed as mean ± standard deviation of three independent experiments. At least three independent experiments were performed for each sample. The treated groups showed statistically significant differences from the control group per the Student's t-test (* p < 0.05). Scale bar 200 μm. THP-1 cells were treated with different concentrations (25–150 μg/mL) of PtNPs for 24 h, and LDH leakage, which acts as a sensitive and integrated tool for measuring the integrity of the cell membrane, was determined [35]. Biotoxicity of nanomaterials can be estimated from the leakage of LDH, which is released in culture medium from cells after plasma membrane damage. LDH is commonly known as an indicator of NP penetration into cells; it is ubiquitously present in nearly all cells and is expelled through the damaged plasma membrane into the extracellular space [51]. Plasma membrane damage is the major cause of cell death. Several studies have shown that nanoparticles increase LDH leakage in various cell types, including human breast cancer, ovarian cancer, adenocarcinoma, and neuroblastoma cells [34,47,52,53]. Our results also demonstrated that PtNPs induced LDH leakage in a dose-dependent manner (Figure 4A). Similarly, Wright et al. [54] had demonstrated that carbon nanodots increased LDH leakage in THP-1 cells. To further confirm the results obtained from the LDH assay, we counted the number of cells after staining with trypan blue. Incubation of cells with various concentrations of PtNPs for 24 h significantly reduced cell viability (Figure 4B). Results suggest that membrane integrity was compromised in PtNPs-treated cells. Kutwin et al. [49] demonstrated the effect of PtNPs on U87 glioma cells; their results suggested that the cells treated with PtNPs showed morphological deformations, DNA damage, decreased metabolic activity, and genotoxic effects. Exposure of human cervical cancer SiHa cells to platinum-copper alloy nanoparticles inhibited cell proliferation and enhanced nuclear morphological changes, including cell shrinkage, intranucleosomal DNA fragmentation, and chromatin condensation [55]. Collectively, PtNPs potentially induce membrane damage in THP-1 cells. Similarly, carbon nanoparticles such as graphene oxide and reduced graphene oxide increased LDH level and decreased THP-1 proliferation [38]. Oxidative stress results from an imbalance between ROS production and ROS scavenging activity in cells. The balance between pro- and anti-oxidants determines various cellular functions, signal transductions pathways, and cellular metabolism [56]. ROS play an important role in the progression of several diseases, including inflammation, atherosclerosis, aging, and age-related degenerative disorders. We determined the effect of PtNPs on ROS induction in THP-1 cells. PtNPs induced the generation of significant levels of ROS in THP-1 cells (Figure 5A). PtNPs modulate oxidative stress by impairing receptor activator of nuclear factor-κB ligand (RANKL) signaling [57]. Our results showed that ROS production is a common mechanism of toxicity both in vitro and in vivo. Furthermore, oxidative stress induced by PtNPs has been correlated with lipid peroxidation (Figure 5B), which increases MDA levels in cells, eventually leading to oxidative damage of mitochondrial DNA, alterations in the mitochondrial membrane potential, and changes in the expression of key antioxidant enzymes such as SOD2 [58–61]. The exposure of THP-1 cells to PtNPs increased MDA concentration in a dose-dependent manner after 24 h. As shown in Figure 5B, Protein carbonylation, a marker of oxidative stress, is involved in various metabolic diseases. Carbonylation is dependent upon the severity of oxidative stress in cells. For instance, in highly oxidative cells such as macrophages, ROS produced by the NADPH oxidase system may lead to the formation of significant amounts of lipid aldehydes that covalently modify endogenous enzymes and proteins [67]. Therefore, we determined the dose-dependent effect of PtNPs on carbonylation of proteins in THP-cells. Results showed that increasing the dose of PtNPs directly increased intracellular protein carbonyl content by 10–18-fold compared to control cells (Figure 5D). The results regarding these oxidative stress biomarkers indicate a close relationship between cytotoxicity and oxidative stress generated by PtNP treatment. Antognelli et al. (2009) [68] demonstrated the effect of crystalline silica Min-U-Sil 5 on oxidative stress, efficiency of antiglycation and antioxidant enzymatic defenses. Similarly, after treatment with silver nanoparticles, THP-1-derived macrophages showed an approximately 3-fold higher level of protein carbonyls than control cells [69]. Thus, we concluded that PtNP-induced oxidative stress is responsible for the observed cytotoxic effects. Recently, Gurunathan et al. (2019) [38] reported that graphene oxide and reduced graphene oxide increased the level of oxidative stress in THP-1 cells by increasing ROS levels, lipid peroxidation, nitric oxide levels, and protein carbonylation. The most common negative outcome of the use of nanoparticles as therapeutic molecules for cancer is the excessive generation of ROS, which is a key factor in NP-induced toxicity [70]. Imbalance in oxidative status occurs due to production of higher levels of ROS in the system. Thus, antioxidants are required to scavenge the excess free radicals. Reoccurrence of oxidative stress eventually leads to inflammation. Studies have reported that PtNPs are suitable agents for reducing ROS levels under certain conditions [71]. Hence, we attempted to determine the effect of higher concentration and smaller size (larger surface area) of PtNPs on THP-1 cells. To determine the effect of PtNP size on the cellular levels of various antioxidants, we analyzed the expression of markers such as GSH, CAT, SOD, GPx, GST, and GR. Results revealed that the levels of all the tested markers were significantly lower in the treated cells than in the control, and the reduction occurred in a dose-dependent manner (Figure 6). Figure 6. Effect of PtNPs on anti-oxidant markers. THP-1 cells were treated with different concentrations of PtNPs (25–150 μg/mL) for 24 h. After incubation, the cells were harvested and washed twice with ice-cold phosphate-buffered saline solution. The cells were collected and disrupted by ultrasonication for 5 min on ice. (A) Glutathione (GSH) concentration was expressed as percentage of control. (B) Catalase (CAT) was expressed as percentage of control. (C) Superoxide dismutase (SOD) was expressed as percentage of control. (D) Glutathione peroxidase (GPx) concentration was expressed as percentage of control. (E) Glutathione S-transferase (GST) concentration was expressed as percentage of control. (F) Glutathione reductase (GR) concentration was expressed as percentage of control. Results are expressed as mean ± standard deviation of three independent experiments. There was a significant difference between treated and untreated cells per Student's t-test (* p < 0.05). concentrations of PtNPs was lower than that in the control group (Figure 7A). Almeer et al. (2018) [75] suggested that a decrease in MMP level is cytotoxic to HEK293 cells. Figure 7. PtNPs decreased mitochondrial membrane potential (MMP) and ATP content. (A) THP-1 cells were treated with PtNPs (25–150 μg/mL) for 24 h, and the mitochondrial membrane potential (MMP) was determined using the cationic fluorescent indicator JC-1. (B) THP-1 cells were treated with PtNPs (25–150 μg/mL) for 24 h and the intracellular ATP content was determined according to the manufacturer's instructions (Catalogue Number MAK135; Sigma-Aldrich). The results are expressed as the mean ± standard deviation of three independent experiments. The treated groups showed statistically significant differences from the control group per the Student's t-test (* p < 0.05). The ATP level is crucial for regulating cellular metabolic activity; therefore, we investigated the dose-dependent effect of PtNP on ATP level. The results showed that ATP content decreased in cells treated with PtNPs, which is in agreement with the decrease in cellular metabolism in the cell viability test (Figure 7B). Similarly, the exposure of human colon cancer cells to silver nanoparticles altered MMP, lowered ATP concentration, and increased Bax expression. Recently, Gurunathan et al. (2019) [38] reported that graphene oxide and reduced graphene oxide treatment lowers ATP level in THP-1 cells. To investigate the mechanisms associated with DNA damage and apoptosis in PtNPs-treated THP-1 cells, we determined the expression of key genes involved in mitochondria-mediated apoptosis. Results showed significantly higher expression of proapoptotic genes such as those encoding p53, p21, Bax, Bak, caspase 9, and caspase 3 (Figure 8). A previous study showed that exposure of THP-1 cells to ZnONPs increased the expression of endoplasmic reticulum (ER) stress- and apoptosis-related gene expression, which is dependent on the interactions between ZnONPs and biological molecules [76]. Similarly, Almeer et al. [75] reported increase in the levels of Bax and caspase 3 and decrease in the level of Bcl2 in PtNP-treated HEK293 cells. The downregulation of Bcl-2 mRNA and concomitant upregulation of proapoptotic genes indicated that Bcl-2 and Bax play a key role in the execution of apoptosis and that the Bcl-2 family is involved in the regulation of apoptosis. Similarly, carbon nanoparticles such as graphene oxide and reduced graphene oxide increased the expression of proapoptotic genes and downregulated antiapoptotic genes in THP-1 cells. Figure 9. PtNPs increase oxidative damage to DNA and RNA. (A) THP-1 cells were treated with various concentrations of PtNPs (25–150 μg/mL) for 24 h. 8-Oxo-dG level was measured after 24 h of exposure. (B) THP-1 cells were treated with various concentrations of PtNPs (25–150 μg/mL) for 24 h. 8-Oxo-G level was measured after 24 h of exposure. Results are expressed as mean ± standard deviation from three independent experiments. There was a significant difference between treated cells and untreated cells per Student's t-test (* p < 0.05). Oxidative stress is one of the most common mechanisms of nanoparticle-induced DNA damage, and oxidatively-induced DNA lesions are predominantly repaired by the base excision repair (BER) pathway. Damage to DNA bases due to oxidative stress are deleterious, leading to stalled replication forks, mutations, and eventually cell death [84]. To understand the genotoxicity of PtNPs, we assessed the expression of various genes involved in DNA repair as an adaptive response to PtNPs. We determined the expression levels of relevant DNA glycosylases involved in BER, such as OGG1, APEX1, CREB1, POLB, UMG, and GADD45A. PtNP treatment upregulated all the tested genes in a dose-dependent manner (Figure 10). These results indicated that the PtNPs modulate the expression of DNA glycosylases. Titanium dioxide NPs (TiO2 NPs) induce the activation of the serine/threonine kinase ATM/Chk2, which is involved in the DDS signaling pathway [85]. TiO2 NPs-induced increase in the expression of ATM in hepatocellular carcinoma cells (HepG2) leads to induction of double strand breaks (DSBs), chromatin condensation, nuclear fragmentation, and apoptosis due to increased ROS production and subsequent DNA damage [86]. The exposure of cells to engineered nanomaterials affects the functioning of the entire cellular system, including fidelity of DNA replication and cell division, resulting in wide ranging DNA lesions that include genome rearrangements, single strand breaks (SSBs), DSBs, intra/inter strand breaks (SBs), and formation of modified bases. DNA lesions can lead to chromosomal aberrations, mutations, apoptosis, carcinogenesis, or cellular senescence [87–90]. Gurunathan and colleagues observed that carbon nanomaterials such as graphene oxide and reduced graphene oxide cause oxidative DNA damage by increasing the expression of DNA damage and repair genes in THP-1 cells [38]. ROS cause oxidative damage to various macromolecules, including proteins, lipids, and nucleic acids [91]. Repair of DNA lesions is crucial for maintaining genomic integrity, whereas other oxidatively damaged macromolecules undergo degradation [92]. Previous studies have reported that proinflammatory cytokines are induced after exposure of RAW264.7 macrophages [93] and THP-1 monocytes [94,95] to PM2.5. However, studies on the effect of PtNPs on THP-1 cells are lacking so far. Therefore, we investigated the molecular mechanism of macrophage inflammatory responses to PtNPs. To determine whether proinflammatory cytokines are upregulated in THP-1 cells after exposure to PtNPs, we determined the protein levels of interleukin-1β (IL-1β), IL-6, IL-8, tumor necrosis factor-α (TNF-α), granulocyte-macrophage colony-stimulating factor (GM-CSF), and monocyte chemoattractant protein 1 (MCP-1). Exposure of THP-1 cells to PtNPs significantly increased the expression levels of IL-1β, IL-6, IL-8, TNF-α, GM-CSF, and MCP-1 in a dose-dependent manner (Figure 11). Gold nanoparticles (GNPs) altered proinflammatory cytokine expression in rat liver and kidneys. Exposure of innate immune cells such as macrophages, dendritic cells, and monocytes to SiNPs increased production of pro-inflammatory cytokines such as IL-1b and IL-18 [96]. The expression levels of IL-1, IL-6, and TNF-α were significantly increased in silver nanoparticle-treated THP-1 cells and primary blood monocytes [97]. Recently, Khan et al. [98] reported that a single intraperitoneal injection of small-sized GNPs (5 nm) significantly increased IL-1 beta and IL-6 mRNA expressions in mouse brain, whereas the larger GNPs (20 and 50 nm) did not produce any inflammatory response. This indicates that the size of the nanoparticles plays a critical role in cytokine production. Gomez et al. [99] evaluated the potential in vitro immunomodulatory effect of 12-nm and 200-nm SiNPs on the expression of pro-inflammatory cytokines and NLRP3 inflammasome components in human primary neutrophils and peripheral blood mononuclear cell PBMCs. They observed that SiNPs induced the production of pro-inflammatory cytokines in a dose-dependent manner. Exposure of macrophages to atmospheric particulate matter with aerodynamic diameter less than 2.5 μm (PM2.5) significantly increased the intracellular levels of TNF-α, IL-1β, and IL-6, in both a dose- and time-dependent manner. Recently, Gurunathan et al. [38] suggests that graphene oxide and reduced graphene oxide aberrantly increase the expression of proinflammatory cytokines and chemokines. Our findings demonstrate that the smaller size of PtNPs is responsible for the production of proinflammatory cytokines in a dose-dependent manner. Owing to their unique physical and chemical properties, metallic nanoparticles (NPs) are being used in biomedical applications and as consumer products. Pt is also used as a vehicle exhaust catalyst, leading to possible exposure via inhalation. Despite their use, data on the cytotoxic, genotoxic, and proinflammatory responses to them and possible size-dependent effects are limited, particularly for PtNPs. Therefore, we analyzed the effect of ultra-small PtNPs (1–2 nm) on cytotoxicity, genotoxicity, and proinflammatory responses in THP-1 cells. First, we synthesized PtNPs using apigenin and characterized them using various analytical techniques. PtNPs caused cytotoxicity by decreasing cell viability and proliferation in a dose-dependent manner. The potential cytotoxicity of PtNPs on cellular redox systems was determined by determining LDH leakage, ROS generation, lipid peroxidation, and expression of antioxidants. Interestingly, LPO-generated 4-hydroxynonenal (HNE) and its protein adducts could play crucial role in cytoto, geno and immunotoxicity and these products could be a possible mechanism of PtNPs-induced toxicity (Figure 12). Studies are required to address these biological effects of PtNP which will be a focus point of future research. The results suggest that PtNPs potentially induce oxidative stress by increasing the levels of oxidative stress markers such as ROS and lipid peroxidation, and decrease the levels of antioxidant markers. The involvement of mitochondria in oxidative stress and genotoxicity was determined by assessing MMP, ATP level, and gene expression. The results of all these cellular assays clearly suggest that PtNPs treatment potentially leads to loss of mitochondrial functions. The possible mechanism of PtNPs-induced cell death was confirmed by gene expression analysis, which revealed upregulation of apoptotic markers such as p53, p21, Bax, Bak, caspase 9, and caspase 3, and simultaneous downregulation of antiapoptotic genes encoding Bcl-2 and Bcl-xl. The impact on DNA damage was confirmed by increased expression of DNA damage and repair genes associated with increases in the levels of 8-oxoG. PtNP treatment increased oxidative DNA damage and impaired DNA integrity. Altogether, PtNPs were able to stimulate multiple stress responses such as oxidative stress, cytotoxicity, genotoxicity, and secretion of proinflammatory cytokines. However, further in vivo and molecular mechanistic studies are necessary to understand the trafficking patterns of these nanoparticles inside cells and the mechanism underlying the proinflammatory response. Figure 12. The possible mechanism of PtNPs on induced cyto, geno and immunotoxicity through oxidative stress, LPO-generated 4-hydroxynonenal (HNE) and its protein adducts and DNA damage in THP-1 cells. Supplementary Materials: The following are available online at www.mdpi.com/xxx/s1, Table S1: The sequences of the PCR primers. Author Contributions: S.G. proposed the idea for this study and was involved in the experimental design, data analysis, synthesis of platinum nanoparticles, cellular assays and writing of the manuscript; M.J. performed the cell morphology assays; M.-H.K. ensured the quality of figures and reference arrangements; J.-H.K. provided the project and facility for the study, and supervised all of the work done. All authors read and approved the final manuscript. Funding: This work was supported by a grant from the Science Research Center (2015R1A5A1009701) of the National Research Foundation of Korea. Acknowledgments: Sangiliyandi Gurunathan is a recipient of the Konkuk University Research Professor Programme. Conflicts of Interest: The authors declare no conflicts of interest. 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/). </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 25.0, \"viab\": 100.0}, {\"h\": 24.0, \"mgL\": 50.0, \"viab\": 80.027}, {\"h\": 24.0, \"mgL\": 75.0, \"viab\": 59.783}, {\"h\": 24.0, \"mgL\": 100.0, \"viab\": 40.082}, {\"h\": 24.0, \"mgL\": 125.0, \"viab\": 19.973}, {\"h\": 24.0, \"mgL\": 150.0, \"viab\": 10.054}]}", "role": "assistant" } ]	23.json
[ { "content": "Extract sample context and measurement list. <doc> Contents lists available at ScienceDirect journal homepage: www.elsevier.com/locate/procbio Process Biochemistry a Department of Physics and Nanotechnology, SRMIST, Kattankulathur, Chennai, 603203, Tamil Nadu, India b Innovative Green Product Synthesis and Renewable Environment Development Research Group, Faculty of Environment and Labour Safety, Ton Duc Thang University, Ho Chi Minh City, Viet Nam Keywords: Titanium dioxide nanoparticles Cytotoxicity Cytokines Immune cells Inflammation Nanoparticles (NPs) can interact with the immune system in several ways, which can either enhance or suppress the functions, depending on their characteristics and composition. Though widely used in different biomedical applications, not much is known about the immunomodulatory effects of NPs on human cell lines. In this work, the immunomodulatory effects of titanium dioxide (TiO2) nanoparticles on human monocytic leukemia (THP-1) and human mast (HMC-1) cell lines were studied in a dose-dependent manner. TiO2 NPs of different molar concentrations were synthesized by a simple sol-gel method and analyzed by X-ray diffraction (XRD), Fourier Transform Infrared (FTIR) technique, UV–vis spectroscopy, Field Emission Scanning Electron Microscopy (FE-SEM), Transmission Electron Microscopy (TEM), Dynamic Light Scattering (DLS) and Zeta Potential. The cell viability of THP-1 cells treated with TiO2 NPs was significantly reduced at higher doses as observed by MTT assay. The expressions of Pro and anti-inflammatory cytokines on THP-1 and HMC-1 cells using TiO2 NPs were studied using ELISA method. The results suggested that the secretion of cytokines (IL-6 and IL-10) on human cell lines greatly depended on the concentration of TiO2 NPs. Lower concentrations of TiO2 NPs exhibited immunomodulatory effects by inducing inflammatory responses on the treated human cell lines. Nanoscience has become an emerging research field with wide applications in material sciences, electronics and biomedical sciences [1]. However, several reports were available to understand the characteristics of different transition metal nanoparticles in the biological system [2,3]. Nanoparticles (NPs) are able to stimulate or suppress the immune system via chemotaxis, macrophage activation and cytokine secretion [4,5]. When metal oxide NPs interact with cells, it generates reactive oxygen species (ROS) or some toxic metal ions are released at the surface of the NPs leads to immunotoxicity in the immune system [6]. Effects of immune activation and immune suppression on the immune system have very interesting applications in medicine. Inflammation is the initial response by the damaged or infected cells induced by cytokines and eicosanoids [7]. In order to mediate the normal cellular process and to regulate the immune response against some invaders, the cytokines are interacting with the cells of the immune system [8]. Cytokines are small signaling proteins originated from specific cells of the immune system had specific effect on the cells interaction and intercommunication [9]. Depending on their effects on immunocytes and biological responses, the cytokines have been classified into proinflammatory and anti-inflammatory cytokines. The major pro and antiinflammatory cytokines are lymphokines, interleukins (IL), monokines, interferons (INF), colony stimulating factor (CSF) and tumor necrosis factor (TNF) [10]. Among the cytokines mentioned above, interleukin (IL-6) and interleukin (IL-10) possess vital roles in the stimulation and suppression of the immune system. IL-6 performs like a pro as well as an anti-inflammatory cytokine and it defends the body against infections. It promotes inflammation via development and stimulation of T-cells, differentiation of B-cells and initiation of acute-phase reactants by hepatocytes [11]. Cytokine synthesis inhibitory factor (CSIF) also known as IL-10 has several roles in the immune system. Its key roles include the stimulation of cytokines by macrophages and some accessory functions of macrophages in T-cell activation [12]. T-cells are the major ⁎ Corresponding Authors. E-mail addresses: (A. Pugazhendhi), (K. Thirunavukarasu). 1 The authors contributed equally as corresponding author to this work. https://doi.org/ Received 24 February 2019; Received in revised form 8 July 2019; Accepted 7 August 2019 Available online 16 August 2019 1359-5113/ 2019 Elsevier Ltd. All rights reserved. sources of cytokines that regulate immune responses. Titanium dioxide (TiO2) NPs have been used in several fields viz. as photocatalysts [13], sensors, in biomedicine for cancer treatment [14], photovoltaic cells [15], electrochemical devices [16], cosmetics and pharmaceuticals [17] owing to excellent properties such as high biocompatibility, resistance to body fluids and durability against photo and chemical corrosions [18]. Several reports have shown the cytotoxicity and gene expression of TiO2 NPs and their metabolic activities on human cell lines and it was concluded that the TiO2 NPs have induced inflammatory responses [17,19,20]. TiO2 NPs induced innate immunity and long-lasting lymphocyte responses in the Dark Agouti rats upon exposure [21]. Owing to its high photocatalytic efficiency, TiO2 acts as a photosensitizer for the photodynamic therapy of cancer (PDT), which kills the cancer cells and some pathogenic organisms such as fungi and bacteria by ROS production [22]. Faddah et al. treated adult rats with TiO2 NPs, idebenone and quercetin and reported that the TiO2 NPs triggered oxidative stress and enhanced the expression of inflammatory cytokines [23]. Mano et al. demonstrated the effects of the size of TiO2 NP aggregates on two different human cell lines [19]. This work reports the cytotoxicity and the production of cytokines (IL-6 and IL-10) by the activation of cultured THP-1 and HMC-1 cells mediated by TiO2 NPs in a dose-dependent manner. synthesized particles [24]. 0.025 M of titanium (IV) isopropoxide, dissolved in 1 M of glacial acetic acid and deionized water, was added dropwise on stirring. It was then ultrasonicated for 30 min in a water bath and again kept for stirring. After the reaction was completed, the white coloured precipitate was collected by washing; it was repeated several times with distilled water and dried overnight at 100 °C. Finally, the sample was fine powdered and calcined at 500 °C for 2 h in a muffle furnace. Similarly, TiO2 NPs were prepared with different concentration of titanium isopropoxide (0.05 M and 0.1 M) by following the above mentioned procedure. The synthesized TiO2 samples under different molar concentrations were named as T1 (0.025 M), T2 (0.05 M) and T3 (0.1 M), respectively. The crystal structure of the prepared TiO2 NPs was analyzed by Xray diffraction (X' pert PAN analytical) with Cu – Kα radiation (λ =1.5405 Å) in the 2θ range of 20°–80°. The particle size and morphology of the TiO2 NPs were observed and the images were obtained by Field Emission Scanning Electron Microscope (FESEM) with Energy Dispersive X-ray spectroscopy (EDX) by FEI Quanta FEG 200 and Transmission Electron Microscope (TEM) by FEI-Philips TECNAI 10. Infrared spectra were obtained by Bruker alpha-T Fourier Transform Infrared (FTIR) spectrometer and the optical absorption spectra were recorded by (UV3000+) UV–vis spectrophotometer. The hydrodynamic size of the NPs and their surface charges in solution were measured by Dynamic light scattering (DLS) using Horiba Nanopartica SZ-100. The average hydrodynamic size of the prepared TiO2 NPs in aqueous medium was evaluated by dynamic light scattering experiment. The scattering angle was fixed at 173°. It would also provide information about the polydispersity index (PDI) used to quantify the colloidal dispersion level of the TiO2 NPs. Zeta potential measurement would provide the surface charge and the colloidal stability of the NPs in the aqueous medium. The temperature of the sample holder was maintained at 25 °C. Each result of the NPs suspension was expressed as the average value obtained from three consecutive measurements. THP-1 (Human monocytic leukemia cell line) and HMC-1 (Human mast cell line) were procured from NCCS, Pune, India. THP-1 cell line was cultured in RPMI-1640 medium containing 4.5 g/L glucose, 2 mM L-glutamine, and 10% fetal bovine serum. The HMC-1 cell line was cultured in Iscove's Modified Dulbecco's Medium (IMDM) containing, L-Glutamine but lacking alpha-thioglycerol and beta-mercaptoethanol. IMDM is a modified Dulbecco's media and is regarded as a more intricate and enriched media. The cell culture was maintained between 2 × 106 cells/mL at 37 °C and 5% CO2 and subcultured regularly using a fresh medium with fluid renewal done every 2 to 3 days. The cells under investigation were cultured in culture flasks in a humidified incubator at 37 °C with 5% CO2. After the THP-1 cell growth reached the expected level, the cell suspension was spun at 2000 rpm for 5 min and the cell pellets were resuspended in fresh RPMI-1640 medium. The cell count of the suspension was then adjusted to1 × 106 cells/mL. Various concentrations of NPs (1, 10, 20 and 50 μg/mL) were added respectively to the 96 well plate for MTT assay. The treated cells were subjected to overnight incubation with 5% CO2 at 37 °C. Following overnight incubation, the cell viabilities were examined by MTT (3-(4, 5-dimethyl-2-thiazol)-2, 5-diphenyl-2H-tetrazolium bromide) assay. MTT (5 mg/mL) was added to each well containing the sample and allowed to stand for 4 h. Dimethyl sulfoxide (DMSO) addition enabled the solubilisation of the formazan crystals. Absorbance readings were taken at 570 nm using a multimode reader (Biorad 680, USA). The percentage of cell viability was estimated as the relative absorbance of the number of dead and live cells in the sample. Cell suspension containing 1 × 106 cells/mL (THP-1 and HMC-1) were seeded in the 96 well plate and stimulated with LPS (E. coli). TiO2 NPs of different concentrations (1, 5, 10 μg/mL), suspended in a fresh medium, were added to the cells. The plate was incubated overnight with 5% CO2 at 37 °C. After incubation, the plate was spun at 1500 rpm for 5 min and the cell-free supernatants were collected and stored for cytokine assay. The levels of interleukins, IL-6 and IL-10 in the cell suspension were studied using Enzyme-linked Immunosorbent Assay (ELISA). The 96 well plates were layered overnight with the corresponding capture antibody. Then, the supernatant of cells treated with NPs containing cytokines was added and incubated at room temperature for 2 h. Cytokine detection was done by biotinylated antibody and streptavidinhorseradish peroxidase washing thoroughly with 0.5% Tween 20 in phosphate buffered saline (PBS) between steps. After incubating the sample with tetramethylbenzidine substrate for 15 min, the absorbance at 450 nm was measured using a multimode reader and the cytokine concentration was calculated from the standard curve. The production of cytokines and antibodies by immune cells through the interactions of the NPs with the immune system was shown in Fig. 1. TiO2 NPs triggers the cytokine production by activating the helper T-cells through APC (antigen presenting cell) which has antigen identified by their T-cell receptor. The activation of B-cells resulted in production of plasma cells which contains huge area of cytosol promotes the release of proteins called antibodies [25]. The antibodies are used to treat many diseases and also it designed specifically target some antigen present on cancer cells [26]. The X-ray diffraction pattern of the synthesized TiO2 NPs at three different molar concentrations of titanium precursor was shown in Fig. 2. The diffraction peaks at 2θ values of 25.55°, 38.127°, 47.947°, 54.54°, 55.28°, 63.05°, 69.72°, 70.24° and 75.407°, corresponding to Miller indices (101), (004), (200), (105), (211), (204), (116), (220) and (215), respectively were assigned to the pure anatase phase with a tetragonal crystal structure (JCPDS card no.21-1272). The average crystallite size of the synthesized NPs was estimated by Debye-Scherrer formula (1), Where D was the crystallite size (nm), λ was the wavelength of X-ray (1.5405 Å), β denoted the full width half maximum (FWHM) of the diffraction peak (radian) and θ, representing the diffraction angle (degree). For high-intensity peaks, the calculated average crystallite sizes were 10.5 nm (T1), 9 nm (T2), and 6.2 nm (T3), respectively. The lattice parameters a, b and c were evaluated for the synthesized tetragonal TiO2 NPs using the interplanar distance (d) value, derived from the Bragg's law Eq. (2), The calculated lattice constant a, b, c (Å) values were obtained from the following equations, where h, k, and l were the Miller indices, which were represented on the corresponding diffraction peaks of XRD. The dislocation density (δ) of the synthesized NPs was evaluated from the following equation, where δ was the dislocation density and D was the average crystallite size of the NPs. Most of the characteristics of the material were strongly determined by the presence of dislocations in the sample. The morphology index (MI) was estimated from the FWHM (fullwidth half maxima) of the diffraction peaks in XRD. MI of the synthesized particles was calculated from the following equation, where FWHMh was the highest FWHM value of the peak and FWHMp was the FWHM of the particular peak for which MI has been calculated. The calculated values of the crystallite size, lattice parameters, dislocation density, and morphology index of the synthesized TiO2 NPs are shown in Table 1. As the concentration of the precursor increased, the crystallite size of the NPs decreased and matched with the FWHM value. The dislocation densities were found to increase with a decrease in crystallite size. Larger dislocation density showed a higher hardness of the material [27]. In aqueous dispersion, the pure anatase TiO2 NPs were biologically more active than the rutile ones, especially with respect to cytotoxicity [28]. The results showed that the synthesized NPs were in a single phase (anatase) and indicating the absence of impurities. Ti-Oe mode and the band at 1106 cm−1 represented the COe stretching vibration [31]. The strong band around 655 and 605 cm−1 corresponded to the Ti-O-Ti vibrations and was associated with the characteristic mode of TiO2. The absence of a peak at 1020 cm−1 corresponding to acetic acid showed that the surface of TiO2 NPs was free from impurities, which could be attributed to the calcination process [32]. FESEM micrographs of the TiO2 NPs synthesized at three different concentrations by the sol-gel method are shown in Fig. 4. FESEM images of the samples T1 and T2 showed that the particles were highly agglomerated with rocklike structures and the sample T3 exhibited the formation of spherically shaped nanostructures, which were arranged in a uniform manner with particle sizes around 60 nm. The EDX graph attached with the FESEM images corresponds to sample T3 confirmed the presence of Ti and O components which revealed that the synthesized NPs were in pure form. The results obtained for the sample T3 were in good conformity with the XRD and FESEM results and T3 was taken for further analysis. In the biological environment, the particles in the nanometer range with high surface to volume ratios can easily bind to the surface of molecules [33]. Transmission Electron Microscope (TEM) was used to study the shape and size of the synthesized NPs as shown in Fig. 5a and b. The images revealed that the particles were monodispersed with spherical morphologies. The histogram obtained from TEM analysis (Fig. 5c) revealed that the average crystallite size of the synthesized TiO2 NPs of the sample T3 was around 30 nm with the size distribution between 23 to 38 nm. The viability of THP-1 cells following overnight incubation with particles was investigated by MTT assay (Fig. 8). The purple coloured formazan product occurred only in the active cells due to the reduction of MTT in the mitochondria. The viability of cells greatly depends on the quantity of formazan [40]. The percentage of cell viability was 90% for 1 μg/mL NPs. The cell viability decreased to approximately 60% for 10 μg/mL, showed that the TiO2 NPs were acting against THP-1 cells. As the concentration of NPs was increased to 20 and 50 μg/mL, the cell viability was about 52 and 45%, respectively. In general, the toxicity of the NPs are mainly depends on their size, crystal structure and the nature of the surface coating. TiO2 NPs have the potential to cross the cellular barriers owing to their smaller sizes and exhibit toxic effects through the generation of ROS on exposed cells [41,42]. The death of THP-1 cells was triggered by the TiO2 NPs due to the production of reactive oxygen species (ROS), which created oxidative stress in cells. The generation of ROS by pure TiO2 NPs was 50 times greater than the TiO2 with dopants like silica [43]. This showed that the cytotoxic effect of anatase TiO2 NPs on THP-1 cells was increased in a dose-dependent way. The viability of cells was counted by the ratio of optical density (OD) of the cells treated with different concentrations of NPs to that of control (cells without NPs), using the formula, The data obtained revealed that there were no eloquent differences in cell survival between the cells treated with TiO2 NPs of low doses. Only the highest concentration of TiO2 NPs exhibited a cytotoxic effect, which signified that the NPs would tend to accumulate in an aqueous medium, which would reduce the cell viability of treated cells. The expression of pro-inflammatory (IL-6) and anti-inflammatory cytokines (IL-10) in THP-1 and HMC-1 cells after overnight incubation with 1, 5 and 10 μg/mL of TiO2 NPs (T3) was studied by ELISA method. The concentration of inflammatory markers in THP-1 cell lines with LPS (Lipopolysaccharides) and various concentration of TiO2 NPs exposure are shown in Fig. 9a (IL-6) and 9b (IL-10). The level of inflammatory markers in HMC-1 cells stimulated with LPS and various concentrations of TiO2 NPs were shown in Fig. 10a (IL-6) and 10b (IL-10). The cytotoxic effects of TiO2 NPs seemed to be higher for the concentrations of more than 10 μg/mL. Consequently, for cytokine analysis, only low concentrations of NPs were taken to study the inflammatory responses on human cell lines. From the results, it was observed that the cells treated with LPS showed an increase in cytokine expression compared to the control group [44]. LPS are composed of polysaccharides with a lipid-A moiety found on the outer layer of gram-negative bacteria. It activates the inflammatory response in various cell types by secreting an inflammatory markers or small proteins called cytokines [45]. The nano-sized TiO2 particles facilitated the binding with LPS owing to the higher surface area of NPs. In THP-1 cells, the TiO2 NPs stimulated the production of Pro-inflammatory cytokine (IL-6). The IL-6 induced by 1 and 5 μg/mL of the TiO2 NPs was almost equal to the level of IL-6 secreted by LPS alone and lower induction of IL-6 was observed for the 10 μg/mL. This was possibly due to the size-dependent cellular uptake of NPs [46]. It showed that the anatase TiO2 NPs induced the pro-inflammatory cytokine in cancer cells. This cytokine may act as a B-cell stimulator, which in turn, produces antibodies [47]. Also, this cytokine enhances the antineoplastic functions due to its immunostimulating property, which inhibits the development of tumor [48]. When the NPs concentration increased, there was a gradual decrease in the concentration of IL-10, which could down regulate the pro-inflammatory cytokine production and inhibit the inflammatory response [49]. TiO2 NPs inhibited the production of anti-inflammatory (IL-10) cytokines at higher doses (10 μg/mL), which might be due to the cytotoxic effects of NPs. The immunosuppressive cytokine (IL-10) secreted by tumor cells, protects them from the immune surveillance [50–52]. The prominent decrease in the concentration of IL-6 and IL-10 was found in HMC-1 cells when treated with synthesized TiO2 NPs. This inhibition of cytokines with the increasing concentration of TiO2 NPs reduced the allergic inflammatory responses in human mast cells, triggered by the pro-inflammatory cytokines (IL-6) [53]. The secretory pathway of cytokines induced by NPs was cell specific and the phagocytic THP-1 and HMC-1 mast cells liberated the cytokines via the classical pathway [54]. The overexpression of cytokines or small proteins in normal human cells may cause autoimmune diseases or some cancers [55]. The observed immunomodulatory effects depended on both particle size and material [56]. The reduction of cytokine concentration in HMC-1 cell lines with increasing the amount of TiO2 NPs may suppress the immune system. TiO2 NPs triggered the inflammatory response in THP-1 cells by the secretion of signaling molecules called cytokines, which could activate the immune system to act against the invaders [57]. As the concentration of the TiO2 NPs gradually increased the cytokine stimulation was exacerbated with LPS in both THP-1 and HMC-1 cell lines. The results of the experiment showed that the cells treated with low concentration of TiO2 NPs induced the inflammatory activity through the secretion of cytokines. We suggest that the low concentrations of TiO2 NPs were compatible to the physiological conditions and showed less toxicity when exposed to human cells. Immunomodulators are those, which control the activity of immune response and stimulate or suppress the immune system, accordingly. From the results of XRD, UV, and FTIR, it has been confirmed that the synthesized NPs are pure anatase phase TiO2 NPs. The spherical structures of TiO2 NPs with a size of approximately 30 nm were analyzed using SEM and TEM. The particle size distribution and the stability of particles in the aqueous medium were studied using DLS and zeta potential. The cell viability of THP-1 cells was significantly increased with decreasing the amount of TiO2 NPs exposure. Cytokine analysis by ELISA showed the secretion of cytokines (IL-6 and IL-10) in THP-1 and HMC-1 cells treated with low concentrations of TiO2 NPs. The low concentration of TiO2 NPs showed immunomodulatory effects by activating the immune response on treated cells through cytokine secretion. Further, in vivo studies are required to analyze the complete mechanism of immunomodulatory effects of TiO2 NPs for biomedical applications. One of the authors V. Madhubala sincerely thanks SRM Institute of Science and Technology, Kattankulathur, Chennai, for the award of SRM Research Fellowship to carry out the work. dioxide (TiO2) nanoparticles in Chinese hamster lung fibroblast cells, Toxicol. Vitr. 27 (2) (2013) 864–873. inhibits mast cell-mediated allergic inflammation, Int. J. Mol. Med. 32 (3) (2013) 736–742. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 1.0, \"viab\": 87.472}, {\"h\": 24.0, \"mgL\": 10.0, \"viab\": 61.197}, {\"h\": 24.0, \"mgL\": 20.0, \"viab\": 51.774}, {\"h\": 24.0, \"mgL\": 50.0, \"viab\": 44.9}]}", "role": "assistant" } ]	13.json
[ { "content": "Extract sample context and measurement list. <doc> Journal of Trace Elements in Medicine and Biology xxx (xxxx) xxxx Contents lists available at ScienceDirect journal homepage: www.elsevier.com/locate/jtemb Sebastián Ariel Ferraroa,b , Mariela Gisele Domingoc,d , Analía Etcheverritoa , Daniel Gustavo Olmedob,c , Deborah Ruth Tasata,e,⁎ a Center of Studies in Health and Environment, School of Science and Technology, National University of San Martín, San Martín, Buenos Aires, Argentina b National Council of Scientific and Technical Research (CONICET), Buenos Aires, Argentina c Universidad de Buenos Aires, Facultad de Odontología, Cátedra de Anatomía Patológica, Buenos Aires, Argentina d Research Fellow of the University of Buenos Aires, Buenos Aires, Argentina e Universidad de Buenos Aires, Facultad de Odontología, Cátedra de Histología y Embriología, Buenos Aires, Argentina Keywords: Titanium Nanoparticles Oxidative stress Neurotoxicity Apoptosis Background: Titanium is widely used in biomedicine. Due to biotribocorrosion, titanium dioxide (TiO2) nanoparticles (NPs) can be released from the titanium implant surface, enter the systemic circulation, and migrate to various organs and tissues including the brain. A previous study showed that 5 nm TiO2 NPs reached the highest concentration in the brain. Even though TiO2 NPs are believed to possess low toxicity, little is known about their neurotoxic effects. The aim of the study was to evaluate in vitro the effects of 5 nm TiO2 NPs on a human neuroblastoma (SH-SY5Y) cell line. Methods: Cell cultures were divided into non-exposed and exposed to TiO2 NPs for 24 h. The following were evaluated: reactive oxygen species (ROS) generation, apoptosis, cellular antioxidant response, endoplasmic reticulum stress and autophagy. Results: Exposure to TiO2 NPs induced ROS generation in a dose dependent manner, with values reaching up to 10 fold those of controls (p < 0.001). Nrf2 nuclear localization and autophagy, also increased in a dose dependent manner. Apoptosis increased by 4- to 10-fold compared to the control group, depending on the dose employed. Conclusions: Our results show that TiO2 NPs cause ROS increase, induction of ER stress, Nrf2 cytoplasmic translocation to the nucleus and apoptosis. Thus, neuroblastoma cell response to TiO2 NPs may be associated with an imbalance of the oxidative metabolism where endoplasmic reticulum-mediated signal pathway seems to be the main neurotoxic mechanism. Nanoscience and nanotechnology are developing rapidly. Therefore, concerns with regard to human health risks and the potential toxicity of widely used nanoparticles (NPs) are on the rise [1–3]. On account of its excellent biocompatibility, titanium (Ti) is widely used in dental and orthopedic implants [4]. It is a highly reactive metal, which after exposure to air or fluids quickly develops a passivating layer of titanium dioxide (TiO2). However, as a result of electrochemical corrosion, frictional wear (tribology), or a synergistic combination of the two, ions/particles can be released into the bioenvironment [5–7]. The combination of tribology, corrosion, and the biological environment has been defined as 'biotribocorrosion' [8,9]. TiO2 NPs are widely used, and their entry into the body through inhalation, ingestion, or dermal contact is an exposure risk to humans [3]. Biotribocorrosion of metallic surfaces of titanium biomedical devices is yet another potential source of systemic contamination with TiO2 NPs [5,10]. Irrespective of the entry route into the body, the chemically active metallic NPs could bind to the surrounding tissues and be disseminated to distant organs via the vascular and lymphatic systems [9,11,12]. Previous works in experimental animal models conducted by our research team demonstrated the presence of particle deposits in different organs such as lung, liver, kidney, and blood cells after chronic ip https://doi.org/ Please cite this article as: Sebastián Ariel Ferraro, et al., Journal of Trace Elements in Medicine and Biology, https://doi.org/ Among cellular stressors, metallic particles and TiO2 particles could disturb the cell oxidative metabolism via different mechanisms involving redox reaction, most commonly increasing reactive oxygen species (ROS) generation in different tissues and organs [13,27,46,57,68]. Under normal conditions, ROS are easily neutralized by cellular antioxidant defenses. However, if ROS production increases and/or if the defense mechanisms are diminished, a pro-inflammatory response could occur. Several studies provide evidence that ROS generation can lead to the loss of mitochondrial membrane potential and the release of cytochrome C from the mitochondria cristae to the cytoplasm, which in turn can activate the caspase cascade ultimately triggering apoptotic cell death [28]. Many ROS sources are located in the mitochondria, and many others in the endoplasmic reticulum (ER); ROS is thought to act as local messenger between the ER and mitochondria. The role played by ER–mitochondria includes the regulation of a variety of processes ranging from mitochondrial fission to autophagy and inflammation. Endoplasmic reticulum stress occurs when proteins cannot reach their native folding state, leading to aggregation of unfolded proteins and consequently disturbing cell homeostasis [29,30]. ER stress has been associated with protein degradation via autophagy, which at unusual levels leads to cytotoxic processing or mechanisms such as apoptosis [31,32]. Nrf2 (nuclear factor erythroid 2- related factor) is a transcription factor that coordinates the cellular response against oxidative stress by regulating the expression of several genes that include antioxidant response elements in their promoters [33]. Although Nrf2 is present at low levels, in response to several stressors, including exogenous particles, its expression can be rapidly upregulated. In this regard, several studies have suggested that NP-toxicity should be tested in cultures of relevant target cells, by assessing ROS production [2,34] and its effect on oxidative metabolism. Particularly regarding TiO2 NPs, despite the extent of TiO2 nanotoxicity studies [35], little is known about the neurotoxic effects and underlying toxicity mechanisms of TiO2 NPs. Therefore, the aim of the present study was to evaluate in vitro the effects of 5 nm TiO2 NPs on a human neuroblastoma cell line (SH-SY5Y), used as a neurotoxicity model. Anatase TiO2 NPs were purchased from Nanostructured and Amorphous Materials, Los Alamos, NM. TiO2 NPs were previously characterized by our work group [13]. Briefly, morphological characterization by SEM confirmed average particle size to be 5 nm mostly lentil-shaped. Chemical analysis by EDS confirmed the presence of titanium in the NPs. Prior to use, all particles were sterilized by autoclave (1.5 atm, 20 min) in order to eliminate potential endotoxins. A TiO2 NPs stock solution was prepared by suspending sterile TiO2 particles in cultured media (1 mg/ml). To ensure a uniform distribution of the particles, TiO2 suspensions were prepared by sonicating the samples 5 times for 5 min (Astrason, Misonix) in culture media. To avoid overheating, TiO2 particle dispersions were sonicated in a cold-water bath kept at 10 °C. Neuroblastoma cells were exposed to 5, 10, 50 or 100 μg/ ml of TiO2 NPs for 24 h. Human neuroblastoma cells (SH-SY5Y) were culture following Ferraro et al (2016) [27]. SH-SY5Y were grown in a 1:1 mixture EMEMa: F-12 supplemented with FBS (10%), penicillin (100 I.U.) and streptomycin (100 mg/ml) and maintained at standard tissue culture conditions (5% CO2, 95% RH, 37 °C). Cells were plated at a density of 160.000 cells/ml. Immediately after incubation, 250 μl SDS (10%) were added to stop the MTT reaction and to solubilize the formazan precipitate. Optical density (OD) was measured at 570 nm in a spectrophotometer (Shimadzu UV-1201 V). Caspase-3 and Poly ADP ribose polymerase (PARP) were determined using Anti-Active Caspase-3 pAB (Promega, Madison, WI) and Anti-PARP p85 Fragment pAB (Promega, Madison, WI) antibodies. BiP and Light Chain 3B (LC3B) were determined using Anti-BiP (Cell Signaling, MA, USA) and Anti-LC3B (Cell Signaling, MA, USA) antibodies. Nrf2 transcription factor was assayed using anti-Nrf2 (Santa Cruz Biotechnology, Dallas, TX). Briefly, SH-SY5Y neuroblastoma cells were fixed with paraformaldehyde (4%) and permeabilized with Triton X-100 (0.2%). Fixed cells were blocked with Bovine Serum Albumin (BSA) (1%) for 1 h, and then incubated with the primary antibody for Caspase-3 (1:500), PARP (1:250), BiP (1:250), LC3B (1:250) or Nuclear factor erythroid 2-related factor 2 (Nrf2) (1:250) diluted in a solution of BSA (1%) and 0.3% Triton X100 in PBS 1X for 18 h. Cells were washed twice with PBS, and incubated with secondary antibody conjugated with Cy-3 (1:100) for 1 h. In order to identify cell nuclei, SH-SY5Y cells were counterstained with 100 nM DAPI (Sigma-Aldrich, St. Louis, MO) for 5 min. The cells were observed under a Nikon TE300 fluorescence microscope (λem = 615 nm for Cy3, and λem = 460 nm for DAPI). Results are expressed as the percentage of cells staining positive for Caspase-3, PARP, BiP, LC3B or nuclear Nrf2. Reactive oxygen species (ROS) was determined in control and TiO2 particle-exposed SH-SY5Y cultures using the 2,2-bis(4-nitrophenyl)-5,5 diphenyl-3,3-(3,3-dimethoxy-4,4-diphenylene) (NBT, Sigma–Aldrich) reduction test [38]. The intracellular release of ROS was evidenced by the amount of blue formazan precipitate in the cells after NBT reduction. Human neuroblastoma cells were treated with NBT in the presence and absence of 12-O-Tetradecanoylphorbol-13-acetate (TPA), a known inducer of O2 − production a predominant ROS. All the samples were incubated with NBT at 37 °C for 45 min. After incubation time, paraformaldehyde (0.4%) was added to stop the NBT reaction and fix the cells. The presence of a blue formazan precipitate was indicative of a reactive cell, whereas cells without the precipitate were considered non-reactive. At least 200 cells (reactive and non-reactive) per dish and 3 dishes per experimental condition were scored by light microscopy (Nikon Alphaphot-2 V52 microscope) as described elsewhere [39]. Results are expressed as percentage of reactive cells. Data are presented as the mean ± SD. Control and TiO2 exposed cultures were compared by one-way ANOVA in conjunction with Newman-Keuls or Bonferroni's post-test. Statistical significance was set at p < 0.001. SH-SY5Y cell cultures were exposed to TiO2 (5, 10, 50 and 100 μg/ ml) for 24 h. As shown by MTT assay, cell viability remained unchanged in 5 and 10 μg/ml exposed cultures, whereas 50 and 100 μg/ml of TiO2 NPs induced a marked increase (p < 0.001) in absorbance when compared to the OD levels reached by non-exposed cultured cells treated with tetrazolium salt MTT (Fig. 1). We investigate cell death using two known biomarkers of the apoptotic process. We found that exposure to increasing concentrations of TiO2 (5, 10, 50 and 100 μg/ml) induced a dose-dependent increase in Based on the known oxidative stress-inducing properties of metallic NPs, we evaluated the generation of ROS in SH-SY5Y cell cultures exposed to TiO2 (5, 10, 50 and 100 μg/ml) for 24 h. Fig. 3 shows ROS generation in all experimental groups, as shown by the NBT test. Most cells from control cultures were colorless, whereas TiO2 exposed cells exhibited dark blue granules as a result of formazan precipitation. A statistically significant (p < 0.001) dose dependent response was observed in cultures exposed to TiO2 at a concentration of 10 ug/ml or higher as compared to controls (Fig. 3). In response to either ROS endogenous or exogenous stress, Nrf2 translocates to the cell nuclei triggering antioxidant and phase II enzymes in order to restore redox homeostasis. Hence, we evaluated Nrf2 nuclear translocation using immunocytochemistry. Fig. 4(A) shows representative microphotographs of Nrf2 negative and positive cell nuclei translocation. Fig. 4(B) shows SH-SY5Y response to TiO2 NPs exposure. Increasing doses of TiO2 provoked a marked and significant dose dependent Nrf2 translocation from cytoplasm to nucleus. A large number of studies have established that induction of GRP78 (BiP) is a marker for ER stress. In order to determine ER stress response in cells following exposure to TiO2 NPs, we evaluated BiP expression using immunocytochemistry. Our results showed that TiO2 NPs induced ER stress and triggered transcription of BiP, altering ER homeostasis in human neuroblastoma cells (Fig. 5). To determine whether autophagy is induced in TiO2 exposed cells, the expression level of LC3B, one of the key regulators in autophagy, was examined by immunocytochemistry. As shown in Fig. 6, LC3B expression significantly increased in TiO2 exposed cells compared to controls. Moreover, TiO2 treatment increased both the number of LC3B fluorescent cells and fluorescence intensity. Taken together, our results clearly demonstrate that TiO2 NPs induced autophagy in human neuroblastoma cells. The wider range of nanomaterial applications will almost certainly result in increased potential hazards for living beings [40,41]. Their characterization has therefore gained interest among researchers, particularly in view of contradictory reports regarding their toxicity. TiO2 NPs have been shown to enter the central nervous system via the olfactory pathway or by crossing the blood-brain barrier, causing brain neuron damage, and/or neurodegenerative diseases [42,43]. Therefore, the primary task of the present research was to clarify the molecular mechanisms underlying the neurotoxicity of TiO2 NPs to humans. TiO2 doses generally used for toxicological assays in vitro, range from 0.2 to 500 mg/ml depending on the cell type employed [44–46]. Therefore, in this study we followed the doses reported in the literature. However, taking into perspective a comparison with real world conditions, it should be pointed out that in human studies the average amount of particles that are released due to tribocorrosion from First, we studied cell viability of human SH-SY5Y cells by evaluating the transformation of tetrazolium salts into formazan (MTT assay) after 24 h exposure to 5–100 μg/ml of 5 nm TiO2 NPs. It has been shown that some NPs interfere with formazan salts [Appendix A, 50]. Valdiglesias et al. (2013) [51] reported that TiO2 NPs, exposure within a range of concentrations (20–150 μg/ml) or exposure times (3, 6 and 24 h) showed no reduction on cell viability when assayed by MTT while they found a rise in apoptosis. In our study far from finding reduction on cell viability we found augmentation on MTT. Wolf et al. (2001) [52] showed that TiO2 and ZnO can absorb and scatter visible light possibly influencing the absorbance readouts of colorimetric assays. Particularly, Dunford R et al. (1997) [53] informed that TiO2 can absorb approximately 70% of incident UV. In this sense, we hypothesize that MTT value could be the result of TiO2 NPs particle interference when using spectrophotometry. In addition, other research groups [54] have found that the presence of metallic NPs in cell cultures conceal their potential cytotoxic effect due to particle interference with OD readings. Nevertheless, studies evaluating the effect of TiO2 NPs in other cell types (primary hippocampal neurons or PC12) have shown a decrease in cellular metabolism [55,56]. These controversial in vitro toxicological results could be due to the use of different cell types, particle sizes, exposure concentrations, and length of time in culture. Oxidative stress as a result of an imbalance in the production of reactive oxygen species (ROS) and/or altered antioxidant protection mechanisms has been linked to metallic particles [58]. Since ER regulates oxidative stress sensitivity [59] and the relation between TiO2 NPs and neural cell death has not been explored, we first examined intracellular ROS changes in the ER of human neuroblastoma cells exposed to nano-sized TiO2 particles. Our results showed that TiO2 NPs were able to activate immediate generation of ROS, which can potentially damage proteins, lipids, and DNA. ROS can be produced in the cytosol and in several organelles, including the ER and mitochondria, which are known to be morphologically and biochemically closely associated [60]. Altered redox homeostasis in the ER can be sufficient to cause ER stress, which in turn, could induce the production of ROS [59,61,62]. Current research indicates that in a variety of physiological and pathophysiological conditions, oxidative stress and ER stress are closely linked events. We found an increase not only in ROS production but also in ER stress in response to TiO2 NPs (Figs. 3 and 5). Given the possible harmful impact of oxidative stress, cells have evolved anti-oxidative stress responses to restore cellular redox homeostasis. We used immunocytochemistry to evaluate Nrf2 transcription factor, which is considered a main regulator of ROS-scavenging enzymes protecting against oxidative stress. As shown in this in vitro experimental model, TiO2 NPs stimulated Nrf2 translocation to the cell nuclei in order to restore redox homeostasis (Fig. 4). It has recently been shown that Nrf2 can also be activated after autophagic degradation of Keap1 [63]. In line with these observations, we found a dosedependent increase in both LC3B autophagic biomarker and Nrf2 antioxidant transcription factor in TiO2 NPs exposed neuroblastoma cell cultures. Therefore, the induction of both autophagy and Nrf2 expression in response to stress observed in the present study could be considered pro-survival mechanisms. However, excessive autophagy can contribute to cell death. A number of studies have reported the occurrence of both oxidative stress and autophagy in response to cell death stimuli [64]. In fact, autophagy has been determined to be a potential mechanism of nanotoxicity [65]. It is important to point out that under a number of conditions different cell death mechanisms can be activated simultaneously, and inhibition of one specific pathway may \"switch\" to another. Therefore, autophagy and apoptosis could be potential cell death mechanisms underlying TiO2 NPs-induced neurotoxicity. showed NPs four times smaller than those used by Wu et al. (2010) [68] to cause neurotoxicity and cell death. More studies are needed to further investigate the relationship between brain damage and TiO2 NPs neurotoxicity, by studying the biological response to TiO2 NPs using co-cultures of SH-SY5Y and human microglial cells, a cell type that is responsive to xenobiotics, and is pivotal in oxidative stress. Our results show that TiO2 NPs cause ROS increase, induction of ER stress, Nrf2 cytoplasmic translocation to the nucleus and apoptosis. Thus, neuroblastoma cell response to TiO2 NPs may be associated with an imbalance of the oxidative metabolism where endoplasmic reticulum-mediated signal pathway seems to be the main neurotoxic mechanism. We wish to confirm that there are no known conflicts of interest associated with this publication and there has been no significant financial support for this work that could have influenced its outcome. This work was supported by the University of Buenos Aires [grant number: UBACyT 20020150100032BA]; the National Council of Scientific and Technical Research, Argentina [grant number: PIP 11220130100091]; the School of Dentistry of the University of Buenos Aires [FOUBA Res. (CD) N°330/19-01] and the National Agency for the Promotion of Science and Technology [PICT-2017-1309]. Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/. nanoparticle size influences trace concentration levels in skin appendages, Toxicol. Pathol. 45 (2017) 624–632, https://doi.org/. strategy, Part. Fibre Toxicol. 2 (2005) 8, https://doi.org/-8977-2-8. [35] B. Sha, W. Gao, X. Cui, L. Wang, F. Xu, The potential health challenges of TiO2 endoplasmic reticulum oxidoreductases, mitochondrial electron transport, and NADPH oxidase, Antioxid. Redox Signal. 11 (2009) 2409–2427, https://doi.org/10. 1089/ARS.2009.2625. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 5.0, \"viab\": 99.234}, {\"h\": 24.0, \"mgL\": 10.0, \"viab\": 104.44}, {\"h\": 24.0, \"mgL\": 50.0, \"viab\": 136.91}, {\"h\": 24.0, \"mgL\": 100.0, \"viab\": 162.02}]}", "role": "assistant" } ]	12.json
[ { "content": "Extract sample context and measurement list. <doc> Contents lists available at ScienceDirect Journal of Photochemistry & Photobiology, B: Biology journal homepage: www.elsevier.com/locate/jphotobiol Avishek Chakrabortya,1 , Ankita Dasa,1 , Sreyan Rahab , Ananya Baruia,⁎ a Centre for Healthcare Science and Technology, Indian Institute of Engineering Science and Technology (IIEST) Shibpur, Howrah 711103, India b Department of Physics, Main Campus, Bose Institute, Kolkata 700009, India Keywords: Gold nanoparticles Size dependent cytotoxicity Apoptosis Surface enhanced Raman scattering MG63 cells In the last decade, gold nanoparticles have emerged as promising agents for in vitro bio-sensing and in vivo cancer theranostics. However, different investigations have reported widely varying cytotoxicity and uptake efficiency of gold nanoparticles depending upon their size. Therefore, more extensive studies are needed to standardize these biological effects as a function of size on a particular cell line. In addition, to obtain robust confirmation on the correlation of a size to biological effect, thorough mechanistic study must also be performed. In this study, the size dependent biological activities of gold nanoparticles on osteosarcoma cells is investigated towards exploring their potential theranostic application in bone cancer, for which very scarce literature reports are available. Tris-assisted citrate based method was optimized to synthesize stable gold naoparticles of 40–60 nm sizes. Nanoparticles were characterized through UV–Vis spectroscopy, field emission scanning electron microscope (FESEM) and dynamic light scattering (DLS). Increasing concentrations of gold nanoparticles (AuNPs) of 46 nm size, enhanced the rate of reactive oxygen species (ROS)-induced apoptosis in MG63 cells by disrupting their mitochondrial membrane potential. Considerably higher cell death was observed for 46 and 60 nm AuNPs compared to 38 nm at all concentrations of 200, 400 and 800 ng/mL. Further, molecular signatures of cellular apoptosis under nanoparticle treatment were optically assessed through surface enhanced Raman scattering (SERS). A significant Raman enhancement in cancer cells under treatment of larger gold nanoparticles (46 and 60 nm) at fixed wavelength of 785 nm and laser power of 8.0 mW was evident. In corroboration with molecular biology techniques, SERS observation confirmed the size-dependent apoptotic phenomena in osteosarcoma cells under treatment of gold nanoparticles. Study demonstrates a facile, non-active targeting approach for detection of size-dependent AuNP-induced apoptosis in osteosarcoma cells through labelfree SERS method. Despite significant advancements in understanding of molecular mechanisms behind cancer, absolute cure of the disease or the management of metastatic disease still remains a prodigious mission [1]. The quest for solution to this most challenging human health problem of the present generation, have seen exploration of biophysical and colloidal agents for treatment in addition to the traditional molecular medicinal approach. Specifically, application of nanomaterials has shown promise for a paradigm shift in the field of cancer therapy [2]. Among them, gold nanoparticles (AuNP) are gaining increased attention over other nanomaterials due to their relative biological inertness, strong surface plasmon resonance, and easily controlled size, shape and surface functionalization [3–5]. However, now it is known that AuNP can exhibit cytotoxicity in vitro and in vivo, while more clarifications are necessary to ascertain biodistribution and pharmacokinetics in vivo. Moreover, AuNP show size-dependent internalization and toxicity with these biological properties for a particular size varying from one cell to another. In addition, different sizes of AuNP show toxicity at different concentrations [6,7]. For example, 45 nm AuNP have shown toxicity at 10 μg/mL whereas 13 nm induced toxicity at much higher concentration (75 μg/mL) [8]. Other studies have shown that spherical AuNP are most efficiently internalized by the cells and 50 nm spherical AuNP are considered as the optimum size for higher cellular uptake [9–11]. Similarly, Stojiljkovi'c et al. have compared intercellular uptake of AuNP of three different sizes (15, 40 and 80 nm) by phagocytic neural and non-phagocytic microglial cells [12]. Study showed that the phagocytic cells preferred 15 and 40 nm of AuNP whereas non-phagocytic cells ⁎ Corresponding author. https://doi.org/ Received 17 January 2019; Received in revised form 29 August 2019; Accepted 3 January 2020 Available online 07 January 2020 1011-1344/ 2020 Published by Elsevier B.V. Rc/a: Molar ratio of citrate to HAuCl4. mostly internalized 80 nm particles. In normal cell lines, AuNP of diameter < 10 nm have shown increased cytotoxicity compared to AuNP with diameter in the range of 10–100 nm [16]. However, HeLa cells showed higher uptake efficiency of 50 nm citrate-capped spherical AuNP [17]. It thus becomes clear that any batch-to-batch variation in size of AuNP can have significant impact on the results observed for a particular cellular system of investigation. On the other hand, several researchers have found that the conventional cytotoxic assays like MTT, LDH or fluorescence-based assays are inadequate to provide reliable assessment of AuNP toxicity. Ostensiosly, the mechanism of toxicity can vary between normal and cancer cells. These shortcomings strongly advocate need for generating in-depth correlations between observed toxicity with molecular effects for monodispersed AuNP of different size ranges. One of the most common mechanism of AuNP-induced cytotoxicity is over production of reactive oxygen species (ROS) leading to oxidative stress generation in the target cells [13]. AuNP-associated oxidative stress has been studied in several cancer cell lines including HeLa [14], HepG2 and PMBC cells [15] and such increased ROS burden is reported to induce apoptosis. However, very limited number of investigations has been performed on AuNP interaction with osteosarcoma cells. Some limited understanding is made evident from the study of Tsai et al., who reported that sub-10 nm citrate-capped AuNP cause low long-term cytotoxicity in osteosarcoma cancer cells at concentrations of 1.0 ppm [18]. However, the overall mechanism of such cell death with AuNP treatment is yet not clear. It can be expected that elucidation of endogenous ROS accumulation and mitochondria-mediated apoptotic pathway or membrane potential will provide further insight on the mechanism. Apoptosis via DNA fragmentation is another approach, which occur at par with the consequences associated with mitochondria-mediated apoptotic pathway. In the present study, we hypothesized that the SERS properties of AuNP can be leveraged to obtain comprehensive insights into the mechanism of size-dependent toxicity of these materials on osteosarcoma cells. SERS enables facile ultrasensitive detection of various biomolecules [25–27] and real-time monitoring of living cell organelles [28] while the detection of cellular apoptosis using SERS has potential application for both cancer diagnosis and therapy [29–32]. It is evident that toxicity assessment of diseased cells is predominantly performed by coating the nanoparticles to target specific organelles through complex and time consuming processes [19–21]. In the present study, the size-dependent cytotoxicity of bare AuNP on osteosarcoma cells has been investigated. Apoptosis was analyzed by change in mitochondrial membrane potential and expression of Caspase-3, a pro-apoptotic marker. These events were then correlated with SERS finger-prints. Elucidation of response of the cells to different sizes of AuNP without organelle-specific targeting provides a facile method for standardization of nanomaterial toxicity and its application in bone cancer theranostics. Chloroauric acid (anhydrous HAuCl4, 49% metal basis, 339.79 g/ mol) was obtained from Spectrochem Pvt. Ltd. (India). Tri‐sodium One-pot size-controlled synthesis of AuNP was performed by using the modified Frens method as described by Xia et al. [1]. In this citratebased reduction process, AgNO3 was used as catalyst for rapid seed formation at room temperature, whereas Tris was used as pH buffer of the reactants. Premixed solution containing tri‐sodium citrate, HAuCl4 and AgNO3 were rapidly injected to 47.5 mL of boiling water in a threenecked round bottom flask under reflux and heated at 98.0 ± 1.0 °C under vigorous stirring for 30 min. Solution was cooled to room temperature and stored at 4 °C. The parameters are described in Table 1. Absorbance spectra of AuNPs were characterized using Thermo Scientific Multiskan GO microplate reader (Vantaa, Finland). Wavelength from 300 to 900 nm was tested with step-wise precision of 1 nm. Peak at incidence wavelength showed surface Plasmon resonance (SPR) effect of AuNP of particular size. In case of AuNPs, red shift of SPR peak represents increase in particle size. Size dispersion of AuNPs can be evaluated based on the width of the SPR peak, i.e. broader peak indicate high polydispersity of AuNPs. AuNPs were drop casted on Formvar/Carbon coated Copper Grid by EMSTM and desiccated without vacuum for 24 h prior to electron microscopy. The sample grids were observed using Zeiss Merlin FESEM (Germany) with Gemini II column at 20.0 kV operating voltage and 200.0 kX magnifications. 0.5 mL of nanoparticles was centrifuged, supernatant was removed, and the re-dispersed in Type I water. Hydrodynamic diameters of nanoparticles were investigated using MalvernTM Zetasizer Nano ZS (United Kingdom) equipped 4.0 mW red laser (633 nm) and a detection angle of 173°. Quartz cuvette was used as sample container. For measurement, solvent parameters were dynamic viscosity and refractive index of water, i.e. 0.89 cP and 1.330 respectively. High glucose Dulbecco's modified Eagle medium (DMEM) from (ThermoFisher Scientific; USA), MTT assay kit (Himedia, India), Live/ Dead Double Staining Kit (Merck, USA); DCFDA - cellular reactive oxygen species assay kit from Abcam, USA; FragelTM DNA Fragmentation detection kit from Merck CalbiochemTM, Mitochondrial Membrane Potential Assay Kit(II) from Cell Signaling Technology were purchased. MG63 cell line was obtained from NCCS, Pune, India. Prior to each assay, nanoparticles were centrifuged and re-dispersed in incomplete high glucose DMEM media. MG63 cells were seeded in culture plates and grown in 37 °C and 5% CO2 till they reached 70–80% confluency. Cell media were removed from each well and the cells were washed three times using 1× PBS before treatment with AuNPs. Cells were incubated with 200, 400 and 800 ng/mL of AuNP1, AuNP2 and AuNP3 in serum-free cell culture media for 24 h. After incubation, cells were further washed three times. For live/dead assay, the staining solution was prepared by mixing 1.0 μL of 1.0 mM propidium iodide and 1.0 μL of 2.5 mM Cyto-dye in 1.0 mL of staining buffer. Cells were treated with 100 μL of staining solution and incubated in dark at 37 °C and 5% CO2 for 10 min. Based on the observations from MTT and Live/Dead assay, 800 ng/mL of AuNP was optimized for further experiments. Cells were treated with AuNPs of different sizes (AuNP1, AuNP2 and AuNP3) and incubated for 24 h. Cells were further washed three times using 1× PBS after removal of media and incubated with incomplete high glucose DMEM and DCF-DA at a concentration of 20 μM at 37 °C for 30 min. temperature. Cell permeabilization was performed in 100 μl of 20 μg/ mL proteinase K for 5 min at room temperature. Permeabilized cells were incubated with 100 μl of 1× TdT equilibration buffer for 20 min at room temperature. 60 μl of TdT Labeling Reaction Mixture was added immediately after carefully blotting the TdT equilibration buffer and incubated at 37 °C for 60 min in the dark. The cells suspended in 1× TBS were analyzed using flow cytometry (Becton Dickinson FACS Calibur, India). Cells cultured in a 48 well plate treated with AuNP1, AuNP2 and AuNP3 was first fixed with 4% paraformaldehyde for 15 min and then permeabilized with 0.1% Triton X-100 at room temperature. Cells were incubated with anti-caspase3 primary antibody (Merck Millipore, India) overnight at 4 °C after blocking with goat serum. After thorough washing, cells were incubated with goat anti-mouse IgG conjugated with Alexa Fluor 488 (1:500) (Santa Cruz Technology, Shanghai) secondary antibody for 2 h at room temperature. Immunostained cells were counterstained with DAPI (Sigma Aldrich, Saint Louis, MO, USA). Mounting media (Sigma Aldrich, USA) was used to mount the cells on the slide after thorough washing of the cells with 1× PBS. Images were acquired under inverted fluorescence microscope (Nikon eclipse Tί U, Japan) equipped with 20× objective. Three different fluorescence filters (λex340–380 nm and λem435–485 nm, λex465–495 nm and λem515–555 nm, λex512–552 nm and λem565–615 nm for blue, green and red emission respectively) were used for imaging. Fluorescence intensity of the images was measured using ImageJ software (version 1.44p). Dimension of fluorescence images were 1280 × 1024 pixel. Cells were treated with AuNP1, AuNP2 and AuNP3 and incubated for 24 h at 37 °C and 5% CO2. Cells were trypsinized and washed three times using 1× PBS. Cells were smeared on aluminium foil coated glass slide to avoid photoluminescence background noise in Raman spectra. Raman spectral acquisition was performed using LabRAM HR spectrometer under laser excitation of 785 nm wavelength and ~8.0 mW power. In this study, live cells without any aldehyde-based fixation were observed with hydration provided by 1× PBS. Acquisition time of 15–20s was required in case of AuNP1 for spectral acquisition, whereas Raman spectra with high spectral intensity were achieved in case of AuNP2 and AuNP2 with acquisition time of 5–10s only. In the present study, gold nanoparicles of 38, 46 and 60 nm were synthesized using modified Frens method. UV–vis spectroscopy (Fig. S1 I) of AuNPs showed absorption peaks at 523, 525 and 537 nm for the above mentioned nanoparticle sizes respectively. Further, the size distribution of nanoparticles was confirmed through FESEM and DLS studies (Fig. S1 II and Fig. S2) and the result summarized in Table 2. FESEM images showed the spherical shape of AuNP and the average diameters in aqueous solution were 38 ± 3.1 for 40 nm, 46 ± 2.3 for 48 nm and 60 ± 1.8 for 60 nm of AuNPs (Table 2). The AuNPs maintained spherical shape but slight aggregation after 24 h of incubation in solution was observed by DLS study (Fig. S1 II). DLS showed the average hydrodynamic diameters of nanoparticles in cell free media (Table 2) to be 44 nm for 38 nm, 48 for 46 nm and 64 for 60 nm of AuNPs. Expected size ranges: AuNP1: ~38 nm; AuNP2: ~46 nm; AuNP3: ~60 nm The live/dead assay (Fig. 2 A-J) of MG63 cells after treatment with AuNP showed corroboration with the outcome of MTT assay. In live/ dead assay, numbers of viable cells was significantly (p < .05) reduced at 800 ng/mL in comparison to 200 ng/mL and 400 ng/mL for all nanoparticle sizes. For AuNP1, higher percentage (~90%) of dead cells was observed at concentrations of 800 ng/mL while at the same concentration, percentage of dead cells for AuNP2 and AuNP3 were ~94% and 92% respectively (Fig. 2 K-M). 800 ng/mL concentrations of AuNPs was selected for further studies. Therefore, due to the highest toxicity observed at this concentration, further understanding of toxicity was performed using 800 ng/mL concentrations for each group. It is well known that excessive endogenous ROS production can lead to disruption of membrane potential. The membrane potential of AuNP treated MG63 cells are shown in Fig. 5 (A-D). The intensity plot of membrane potential (Fig. 5 E) showed average red intensity of 15.49 a.u., 9.51 a.u., 7.31 a.u., and 8.98 a.u in MG63 cells of control group (Fig. 5 A) and nanoparticle treated groups (AuNP 1, 2, 3) (Fig. 5 B-D) respectively. The membrane potential was significantly (p < .001) reduced in cells treated with AuNP1, AuNP2 and AuNP3 in comparison to the untreated group. In between treated groups, significant (p < .001) change in membrane potential was observed between AuNP1, AuNP2 and AuNP3. SERS spectra of MG63 cells after treatment of different size of AuNP are shown in Fig. S3 and the pre-assigned peaks are summarized in Table 3. Higher spectral enhancement at lower acquisition time of 5–10 s was observed with AuNP2 and AuNP3 compared to AuNP1 showing low spectral intensity at 20 s of acquisition at fixed wavelength (785 nm) and power (8.0 mW). Spectral changes were evident among the osteosarcoma cells treated with different sizes of AuNPs. Major peaks associated with cellular apoptosis (viz protein, DNA, lipids and amino acids) and their alterations under NP treatment could be identified. Major spectral changes occurred in the osteosarcoma cells treated with AuNP2. In AuNP2 treated cells, the increased intensity of peaks associated with proteins (at 600, 650, 1032, 1130, 1380–90 and 1450 cm−1), DNA (at 712, 850, 870, 1157, 1230–40, 1340, 1380–90, 1470 and 1580 cm−1) and lipids (at 1157 and 1450 cm−1) were observed. On the other hand, decrease in the intensity of peaks associated with DNA at 1080 and 1180–1190 cm−1 were observed in AuNP2 treated group and the results are related to the active intracellular apoptotic process. Correlation of cytotoxic effects of AuNPs with their size on different cancer cells has remained inconclusive, with little attention paid to standardizing the effect of size with molecular mechanisms of cell death. For example, several studies have reported the nontoxic effect of AuNPs on different cell lines [2,3], whereas other research groups reported cytotoxicity of AuNPs in dose-dependent and size-dependent manner [4–6]. Effects of gold nanoparticle size on cellular viability and proliferation have been studied in different cell lines previously. Chen et al. studied the inhibitory effect of 20, 40 and 80 nm AuNPs in human periodontal ligament cells (hPDLC) and human periodontal ligament stem cell (hPDLSC) proliferation [7]. Effect of capping material of gold nanoparticle also plays a role in cellular viability and proliferation. 30 and 50 nm of polyethylene glycol (PEG) coated gold nanoparticles reduced cell viability in human lymphocytes [8] while 8, 15, 40 nm size citratecoated gold nanoparticle have been found to induce apoptosis in HT29, SPEV and A549 cells [9,10]. Citrate coated gold nanoparticles of 40 nm size has also been studied in Sparus aurata with enhanced expression of apoptotic proteins [11]. Gold nanoparticle in the range of 10–40 and 50–60 nm size also induced apoptosis in Vero cells and MCF-7 cells [12]. Reports of gold nanoparticle with sizes higher than 70 nm has also been published in MCF-7 cells [13] and human neutrophils [14]. The size dependent cyto-toxicity of AuNPs has also observed in present study. Lowest viability of MG63 cells has been observed under treatment of 46 nm (AuNP2) gold nanoparticles. In addition to the size, concentration of AuNPs also plays deciding role in cellular uptake as well as cytotoxicity [15]. For instance, citrate-decorated 2 nM of 13 nm AuNPs showed cytotoxicity and induced apoptosis in rabbit chondrocytes whereas the same nanoparticles of 2 nM of 3 mm and 0.02 nM of 45 mm did not affect the cellular viability [16,17]. In other study, it was observed that the gold nanoparticles of different sizes can induce the cytotoxicity only at the concentration > 500μM [18]. Similar to previous observations, our study also reveals that increasing the concentration of AuNP1, AuNP2 and AuNP3 from 200 to 800 ng/mL decreased the viability of MG63 cells. Among the different sizes of monodispersed AuNPs, AuNP2 showed lowest viability of MG63 cells at concentration of 800 ng/mL after 24 h of treatment (Fig. 1). Actually the size dependent uptake and cytotoxic effect of AuNPs is under constant evolution. Several reports show the AuNPs with smaller size ranges have more cytotoxicity than the larger size range. However, recent studies reveal the AuNPs with 50 nm diameter are preferentially uptaken by the cells. In this regard, a study by Chithrani et al. [19] showed the uptake effectiveness of different sizes of citrate capped AuNPs (14, 30, 50, 74, and 100 nm) through [ICP-AES] and results indicated maximum uptake occurred for the 50 nm AuNPs [20]. Another study on cancer cell line showed similar trend, in which [64] AuNPs with intermediate size (~50 nm) when exposed within 30 min were found to be the most abundant cellular AuNPs in the a system [21]. Such interesting phenomenon of cellular uptake was investigated further. A recent study investigates the relation between AuNP size with uptake kinetics and the study showed that intermediate size (~50 nm) NPs were taken up by mammalian cells at faster rate and higher concentration than other sizes. Actually the uptake of NPs is controlled by several factors including ratio of adhesion and membrane stretching, the membrane's bending energy and such parameters affected the cell membrane \"wrapping time\". [22,23] Moreover the cellular uptake is governed by the competition between a) thermodynamic driving force which indicates the free energy required for wrapping the NPs inside cells and b) receptor diffusion kinetics referrers the kinetics of receptor attachment at the binding sites. A pioneering study by Gao et al. has suggested that the wrapping time is dependent on particle size. [24] According to their experiment, NPs with ~50 nm diameter showed the fastest wrapping time as well as receptor-ligand interaction which provides sufficient free energy for cellular internalization and thus enhanced the accumulation of NPs inside cells. However NPs lower than 50 nm do not produce sufficient free energy required to completely wrap NPs on the membrane surface. Hence for smaller NPs, clustering of particles is required for efficient uptake. For example, single particle with 50 nm diameter can enter the cell whereas, for 14 nm, cluster of atleast 6 NPs are required for cellular uptake. On the other hand, NPs larger than 50 nm shows slower receptor-ligand kinetics and thus slow wrapping time. This led to a smaller number of NPs taken up. [25] In the fuzzy based classification of NPs with different hydrodynamic sizes through k-clustering method, a cluster boundary was reported at ~50 nm size. Experimental result indicates this intermediate size has specific propensity to optimum cellular uptake. Another interesting dimension revealed from this study that uptake efficiency of adherent cells and suspended cells are different. The NPs uptake by suspended cells is governed by passive translocation and uptake efficiency is reduced with increase in NPs diameters [26]. After observing this effect with a homogenously distributed AuNP sample, the size-dependent toxicity was correlated with SERS-based molecular fingerprints. In addition to these evidences about the optimum uptake efficiency of mammalian cells at 50 nm size of NPs, present study also revealed the alterations in major biomolecules in NP-treated cells through Raman spectroscopy study. Cells treated with AuNP2 treated cells showed the alterations in specific peaks related to cellular apoptosis like chromatin condensation, inter-nucelosomal DNA cleavage (1375 cm − 1, 780–850 cm − 1) or abnormal lipid or protein concentration etc. Further pathway-based analysis may facilitate in revealing the exact molecular mechanism for the cytotoxic effect of AuNP2 treated cells. Apoptosis levels are affected by the size, shape, and surface chemistry of AuNPs as well as depend on target cell types. AuNPs primarily induce apoptosis through production of excess ROS which alter the cellular redox status, mitochondrial pathways and makes cancer cells more vulnerable [27]. Treating cancer cells with exogenous ROS producing agents like AuNPs result in accumulation of ROS beyond certain threshold level, which cannot be compensated by the defense mechanism of cancer cells generating excess oxidative stress [28]. Interestingly, increased ROS level causes depolarization of mitochondrial membrane potential which further initiates apoptotic signaling cascades [29]. In agreement to the above observations, our study has also demonstrated the size-dependent increase in the ROS load in MG63 cells treated with AuNPs. Gold nanoparticle of 46 nm size showed highest ROS generation along with increased mitochondrial depolarization. Mitochondrial dysfunction with excess oxidative stress can be further correlated to the damage of treated cells. From flowcytometry and IF studies (Fig. 3) it was evident that DNA fragmentation and Caspase 3 expressions are higher in AuNP2 treated groups in comparison to the other treated one. By virtue of surface plasmon resonance, AuNPs provide an Table 3: +: highest intensity;#: second highest intensity; *: least intensity, blank cell: peak not found, str: Stretch;sym: Symmetric;bend: Bending;asym: Asymmetric;twist: Twisting;def: Deformations; Tyr: Tyrosine; Phe: Phenylalanine; Pro: Proline. opportunity for detection molecular alteration in apoptotic cells using Raman spectroscopy [30]. Elsewhere, AuNPs with SERS signatures are used extensively in biomedical applications due to their inert biocompatible properties and high sensitivity in imaging application [31]. Localized SPR and SERS are due to the resonance of free electrons present in the AuNPs [32]. In case of SERS study of MG63 cells, higher Raman spectral enhancement at lower acquisition time of 5–10 s was observed with AuNP2 and AuNP3 compared to AuNP1, which provided low spectral intensity at longer (20 s) acquisition time at a fixed laser irradiation (785 nm, 8.0 mW). From the Raman spectra, significant peaks at different wavenumbers were observed representing vibrational modes of DNA/RNA, proteins and lipids. These spectral alternations can be utilized to monitor cellular dynamics in order to understand cell growth, differentiation and apoptosis [33,34]. Structural and chemical changes in DNA during their interaction with ROS are vital in understanding the physiological role of ROS on DNA damage and cell death. ROS induced DNA damage was confirmed by observing new Raman spectral bands at 640 cm−1, 715–738 cm−1 and 800 cm−1 compared to 660 cm−1 and 740 cm−1 without ROS-DNA interaction. Decrease in intensity in peak at 1075 cm−1, 1159 cm−1, 1179 cm−1, 1334 cm−1, 1438 cm−1, 1502 cm−1 and increase in intensity at 1387 cm−1 was observed due to change in DNA bases and DNA backbone as a result of ROS induced damage [35]. Earlier, it has been reported that treatment of TiO2 nanoparticles on A549 cell line increases triglycerides and cholesterol levels due to oxidative stress accumulation. These changes were observed from the intensity ratio of triglyceride (1071 cm−1) and cholesterol (710 cm−1) peaks. Moreover, the shift of metabolic activity to fatty acid biosynthesis due to mitochondrial dysfunction was also concluded in previous studies by examining the linear increase of the triglyceride/glycogen ratio (1071/ 1018 cm−1) [36]. On similar lines, early apoptosis in Saos-2 cells were reported to exhibit characteristic Raman peak at 1375 cm−1 with increased prominence at late apoptotic stage [37]. Relating to the present work, the increase in Raman signal at 1375 cm−1 was observed in AuNP2 treated cells. This peak is Overall our study reveals that gold nanoparticles posses size and concentration dependent cellular activity. Highest cytotoxicity was observed for AuNP with 46 nm size and 800 ng/mL concentrations. Similar to cytotoxicity, both DNA fragmentation and Caspase 3 studies confirmed that AuNP2 showed highest apoptotic activity in MG63 cells. Gold nanoparticles promoted ROS-dependent cell death by accumulating ROS and disturbing the mitochondrial function. Reduced mitochondrial membrane potential in AuNP treated cells also confirmed the initiation of cellular apoptosis. Further the SERS spectra of AuNP-treated cells exhibited the molecular signatures related to cellular apoptosis. The vibration assignments for apoptotic events have been made from previously reported vibration assignments. Spectral bands specifically provide evidences of on-set of cellular apoptosis in terms of molecular events including – i) protein denaturation/degradation ii) DNA fragmentation and breakage of disulphide bonds iii) alteration in hydrophobic amino acids and loss of tertiary structure of proteins iv) increase in lipid and v) condensation of chromatin structure. Altered band intensity among all different groups exhibited highest apoptotic activity under treatment of AuNP2. Therefore, SERS corroborates the results obtained by molecular biology techniques. Further, SERS spectra showed the opportunity for real time monitoring of the multimolecular complex biological events. Declared none. Supplementary data to this article can be found online at https:// doi.org/. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": 0.2, \"viab\": 98.827}, {\"h\": 24.0, \"mgL\": 0.4, \"viab\": 91.134}, {\"h\": 24.0, \"mgL\": 0.8, \"viab\": 79.531}, {\"h\": 24.0, \"mgL\": 0.2, \"viab\": 91.134}, {\"h\": 24.0, \"mgL\": 0.4, \"viab\": 87.223}, {\"h\": 24.0, \"mgL\": 0.8, \"viab\": 47.849}, {\"h\": 24.0, \"mgL\": 0.2, \"viab\": 91.134}, {\"h\": 24.0, \"mgL\": 0.4, \"viab\": 87.353}, {\"h\": 24.0, \"mgL\": 0.8, \"viab\": 56.845}]}", "role": "assistant" } ]	11.json
[ { "content": "Extract sample context and measurement list. <doc> Cell cultures: Human ovarian cancer cells A2780 and CP70 were cultured in DMEM and RPMI 1640 medium respectively. Human bronchial epithelial cells (BECs) and airway smooth muscle (ASM) cells were enzymatically isolated using previously described techniques1, 2 from 3rd generation or higher bronchi excised from lung samples incidental to patient surgery. BECs and ASM cells were maintained (up to 3 passages) in serum-free bronchial epithelial growth medium (BEGM) 3 or phenol red-free DMEM/F12, respectively. Phenotypes were verified by expression of cytokeratins and CD31 (BECs) vs. smooth muscle myosin or actin (ASM). Experiments were performed in cells at ~80% confluence. ratiometric Ca2+ indicator) and simultaneously visualization using appropriate filters. 3 H-Thymidine incorporation assay for cellular proliferation: Cells (2x104 ) in 24-well culture plates were treated with + AuNPs for 30 min, rinsed with phosphate buffered saline (PBS) and exposed to cell-appropriate fresh media. Thymidine incorporation assay was performed as previously published 5 . In brief, after 24h, 1 μCi/ml [3H]thymidine was added, and 4h later, cells washed with ice-cold PBS, fixed with 100% cold methanol, washed again, and lysed with 250 μl 0.1N NaOH. [3H]thymidine incorporation was measured in scintillation solution. Cellular apoptosis assay: Cells seeded in 60 mm dishes at 3 x 105 and at ~80% confluence were treated with 0 (vehicle), 0.4 or 0.8 μM +AuNPs for 30 min. They were then rinsed with PBS, replenished with fresh media and maintained in culture overnight. Annexin FITC-propidium iodide staining flow cytometry was performed per manufacturer's protocol (BioVision, CA). Cellular viability assay: Cells in 96-well plates at ~80% confluence (10,000– 20,000 cells) were exposed for 30 min to + AuNPs (0 (vehicle), 0.4, 0.8, or 1.2 μM), washed, and then maintained for 24h. Subsequently, 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) was added for 30 min, and absorbance at 490 nm measured. Nanoparticle Fabrication: Gold nanoparticles were synthesized according to reported procedures 6 . Dynamic light scattering DLS and zeta potential measurements were carried out using a Nano series Zetasizer (Malvern Instruments). Quantification of Gold Uptake (Instrumental Neutron Activation Analysis; INAA): Samples were analyzed by INAA as previously described 7 . Cells were grown in 60 mm dishes. Once 80% confluent, the cells were incubated with 0.4 μM +AuNPs for 30 min and rinsed in PBS five times. After being trypsinized for 10 minutes, the cells were collected into eppendorf tubes and pelleted (10 min, 1400 rpm, 4°C). They were subsequently transferred and weighed into precleaned, high-density polyethylene irradiation vials and lyophilized to constant dry weight. Samples were then reconstituted with 100 μL sample solution, loaded in polyethylene transfer \"rabbits\" and irradiated for 90 s in a thermal flux density of ~5×1013 ncm-2 s-1 . Samples were allowed to decay for 24-48 h and counted in real-time on a high-purity germanium detector for 3600 s at a sample-to-detector distance of ~5 cm. Gold mass was quantified by measuring the 411.8 keV gamma ray from β decay of 198Au (t1/2=2.7 days), and calibrated using certified gold standard solutions as described previously. TEM Imaging: Cells grown on Aclar film in 24 well plates were exposed to 0.4 μM + AuNPs for fixed durations (5 min, 2h, 6h), thoroughly rinsed with PBS, and fixed with Trumps fixative for microtome sectioning and TEM microscopy 8 . 1. Fulcher, M. L.; Gabriel, S.; Burns, K. A.; Yankaskas, J. R.; Randell, S. H. Methods Mol Med 2005, 107, 183-206. 2. Prakash, Y. S.; Thompson, M. A.; Pabelick, C. M. Am J Respir Cell Mol Biol 2009, 41, (5), 603-11. 3. Fulcher, C.; Kaukinen, C. AIDS Care 2005, 17, (3), 386-96. 4. Grynkiewicz, G.; Poenie, M.; Tsien, R. Y. J Biol Chem 1985, 260, (6), 3440-50. 5. Patra, C. R.; Bhattacharya, R.; Wang, E.; Katarya, A.; Lau, J. S.; Dutta, S.; Muders, M.; Wang, S.; Buhrow, S. A.; Safgren, S. L.; Yaszemski, M. J.; Reid, J. M.; Ames, M. M.; Mukherjee, P.; Mukhopadhyay, D. Cancer Res 2008, 68, (6), 1970-8. 6. You, C. C. M., O.R. Gider, B. Ghosh, S. P. Kim, I. Erdogan, B. Krovi, S. A. Bunz, U. H. F. Rotello, V. M. Nature Nanotechnology 2007, 2, 318-323. 7. Kattumuri, V.; Katti, K.; Bhaskaran, S.; Boote, E. J.; Casteel, S. W.; Fent, G. M.; Robertson, D. J.; Chandrasekhar, M.; Kannan, R.; Katti, K. V. Small 2007, 3, (2), 333-341. 8. Bhattacharya, R.; Patra, C. R.; Earl, A.; Wang, S.; Katarya, A.; Lu, L.; Kizhakkedathu, J. N.; Yaszemski, M. J.; Greipp, P. R.; Mukhopadhyay, D.; Mukherjee, P. Nanomed 2007, 3, 224-238. approximately 3 min. The extent of depolarization was greater in the ovarian cancer cells and in epithelium, compared to airway smooth muscle cells. In comparison to positively charged AuNPs, those with other charges had negligible effects on membrane potential in any cell type. In control experiments, each of these cell types were exposed to 40 mM KCl (black arrow also indicates time of KCl application), which produced fairly immediate depolarization across cell types (see Figure 1 of main text for summary). mM KCl (black arrow also indicates time of KCl application), which produced [Ca2+]i elevations across cell types albeit with different time delays and profiles (see Figure 3 of main text for summary). smaller, especially in BECs. Similarly, KCl by itself did induce some degree of apoptosis in BECs and ASM, but again not to the substantial extent induced by + AuNPs (Figure 4). Values are means + SE. * indicates significant KCl effect (p<0.05). From the SelectedWorks of Vincent Rotello June 2010 Effect of Nanoparticle Surface Charge at the Plasma Membrane and Beyond Contact Author Start Your Own SelectedWorks Notify Me of New Work Available at: http://works.bepress.com/vincent_rotello/21 †Department of Biochemistry and Molecular Biology, ‡Department of Anesthesiology, §Department of Physiology and Biomedical Engineering, and \| Mayo Clinic Cancer Center, Mayo Clinic College of Medicine, Rochester, Minnesota 55905, ⊥Department of Chemistry, University of Massachusetts, Amherst, Massachusetts 01003, and #Department of Chemistry and University of Missouri Research Reactor, University of Missouri, Columbia, Missouri 65211 ABSTRACT Herein, we demonstrate that the surface charge of gold nanoparticles (AuNPs) plays a critical role in modulating membrane potential of different malignant and nonmalignant cell types and subsequent downstream intracellular events. The findings presented here describe a novel mechanism for cell-nanoparticle interactions and AuNP uptake: modulation of membrane potential and its effect on intracellular events. These studies will help understand the biology of cell-nanoparticle interactions and facilitate the engineering of nanoparticles for specific intracellular targets. KEYWORDS Gold nanoparticles, surface properties, cancer, membrane potential, calcium, apoptosis I n recent years, significant effort has been devoted to develop nanotechnology for the delivery of small molecular weight drugs, as well as macromolecules such as proteins, peptides, or genes into cells and tissue.1-7 Targeted nanoparticle-mediated drug delivery may be used to direct the particles to specific tissues (minimizing toxicity), improve oral bioavailability, sustain drug/gene effect in the target tissue, solubilize drugs for intravascular delivery, and/or improve the stability of therapeutic agents against enzymatic degradation.8 Despite the fantastic potential for nanoparticle use in medicine, fundamental studies to understand the molecular interactions of nanoparticles with their target cells (normal as well as malignant) remain largely unexplored. One such mechanism of action may be ionic interactions; the negative membrane potential of most cells likely interacts differently with nanoparticles of a positive versus negative charge density. These interactions could, in turn, determine intracellular uptake and localization of the nanoparticles and their biological functions. Understanding such interactions between cells and nanoparticles with different surface properties is important not only for engineering of nanoparticles that exhibit selective intracellular uptake (to subsequently modulate cellular processes of interest) but also for determining the relative cytotoxicity of nanoparticles. All living cells have an inherent membrane potential that is determined by ionic permeability and modulated by A2780, BEC,15 and ASM16 cells loaded with the fluorescent, fast-response membrane potential-sensitive dye RH414. The baseline plasma membrane potential ranged between -75 and -55 mV depending on cell type. With images taken at 1-2 frames/s, fluorescence levels remained stable for at least 5 min in vehicle controls.16 Among the four species of AuNPs, only +AuNPs induced membrane depolarization across different cell types (Figure 1B). In comparison, membrane depolarization induced by -AuNP, 0AuNP or (AuNP was negligible (Figure 1B, also see Figure S1). The extent of membrane depolarization was found to be dependent on +AuNP concentration (Figure 1C; p < 0.05 compared to vehicle control) with minimal depolarization at 10 nM, and substantial depolarization at 1.2 μM +AuNP in less than 10 s with maximum depolarization reached in ∼5 min across cell types ( Figure S1). Among cell types, the extent of depolarization was greatest in ovarian cancer cells (CP70, A2780) and comparable to that achieved with 40 mM KCl (which produces a depolarization to ∼-25 mV) (Figure 1B, and Figure S1). We verified lack of fluorescence quenching by examining the effect of AuNPs on RH414 fluorescence in an in vitro acellular preparation (not shown). Next, we wanted to investigate the factors that determined intracellular uptake of AuNPs, focusing on membrane potential. In CP70, A2780, BEC, and ASM cells, uptake of +AuNPs (as determined by INAA17) was significantly higher than AuNPs of other charges (Figure 2A; p < 0.05). However, prior depolarization of the plasma membrane using 40 or 80 mM KCl (which changed membrane potential to ∼-25 and ∼-8 mV, respectively) resulted in significant reduction in the extent of +AuNP uptake in all cell types (Figure 2B; p < 0.05). Furthermore, in cells pre-exposed to KCl, the extent of membrane depolarization induced by 1.2 μM +AuNPs was significantly smaller, confirming the inability of these particles to depolarize the membrane under these conditions (Figure 2C; p < 0.05). In summary, these data clearly demonstrate a key role for membrane potential in intracellular uptake of AuNPs. Furthermore, by altering membrane potential, AuNPs may modulate their own uptake. A2780, BEC, and ASM cells loaded with the ratiometric fluorescent Ca2+-sensitive dye fura-2/AM, baseline [Ca2+]i ranged between 75 and 120 nM (depending on cell type). In conformity with changes in membrane potential, addition of 1.2 μM +AuNP to CP70 and A2780 cells resulted in immediate and sustained increases in [Ca2+]i while in BEC and ASM cells, the increase in [Ca2+]i was slightly delayed. In all cell types, [Ca2+]i levels increased rapidly to a plateau level ( Figure S2), with maximum [Ca2+]i reached in ∼5 min (Figure 3A). Some cells displayed an initial higher [Ca2+], followed by a decay to a lower level above baseline ( Figure S2). Addition of AuNPs of other surface charges produced negligible changes in [Ca2+]i levels (Figure 3A,B and Figure S2). In control experiments, each of these cell types were exposed to 40 mM KCl, which produced [Ca2+]i elevations across cell types albeit with different time delays and profiles ( Figure S2). The extent of change in [Ca2+]i was concentration-dependent with significant changes observed even at 10 nM +AuNPs (Figure 3C; p < 0.05). As with RH414, lack of fura-2 quenching by AuNPs was verified using the cell-impermeant pentapotassium form of fura-2 (not shown). Finally we wanted to investigate whether +AuNP modulation of membrane potential and +AuNP uptake affects cellular proliferation or viability. +AuNPs completely inhib- ited proliferation (determined by 3H-thymidine incorporation4 ) of BECs, whereas proliferation of CP70 and A2780 cells remained largely unaffected (Figure 4A; p > 0.05). Furthermore, apoptosis (determined using annexin-propidium iodide assay) was only slightly increased in CP70 cells following +AuNP exposure (Figure 4B). In contrast, BEC cells displayed substantial apoptosis (Figure 4B; p < 0.05). Indeed, cellular viability (determined by an MTS assay) of normal BEC and ASM cells was substantially reduced by +AuNP exposure (Figure 4C; p < 0.05). To determine whether these changes in cellular proliferation and apoptosis were a result of +AuNP-induced membrane depolarization and [Ca2+]i elevation, we performed control studies where 40 or 80 mM KCl was used to induced membrane potential and [Ca2+]i changes. Cells were exposed to KCl only for 5 or 30 min (to transiently induce [Ca2+]i changes) to mimic +AuNP exposure. Compared to +AuNPs, KCl induced substantially lesser apoptosis and affected proliferation of BECs to a lesser extent ( Figure S3, compare to Figure 4). Furthermore, unlike +AuNPs, KCl had negligible effects on apoptosis of ASM cells. In all of these experiments, it must be noted that the duration of AuNP (or KCl) exposure was brief (minutes), while apoptosis or proliferation was evaluated after ∼24 h (overnight). Accordingly, these changes are unlikely to reflect short-term cell death resulting from cytotoxicity of AuNPs. These novel data highlight several characteristics of +AuNPs: (1) uptake of +AuNPs results in substantial inhibition of proliferation and decreased viability of normal cells, but not of cancer cells, even though comparable membrane depolarization and increased [Ca2+]i occurs across cell types. These differential effects of +AuNPs on normal versus malignant cells, and their potential relevance to nanoparticle design and applications are interesting and require further investigation; (2) the fact that within a cell type (e.g., BECs), +AuNP effects on proliferation, apoptosis, or viability are greater than that of KCl only (in spite of comparable depolarization or [Ca2+]i elevation) indicates that +AuNP effects on cells are mediated not only via altered membrane potential and [Ca2+]i, but additional effects on signaling pathways. Accordingly, an important aspect of understanding AuNP action may be identifying different signaling mechanismsthatmaybetargetedbyAuNPs,withnormalversus cancer cells being differently sensitive to alterations in these mechanisms (especially relating to apoptosis and proliferation). In conclusion, we have demonstrated that cellular membrane potential plays a prominent role in intracellular uptake of AuNPs. Perturbation of the membrane potential is dependent on surface charge of the nanoparticles; positively charged nanoparticles depolarize the membrane to the greatest extent with nanoparticles of other charges having negligible effect. Such membrane potential perturbations result in increased [Ca2+]i, which in turn inhibits the proliferation of normal cells whereas malignant cells remain unaffected. The mechanisms by which positively charged nanoparticles interact with the plasma membrane need to be further investigated. Such interactions may involve AuNPs binding to the plasma membrane. Indeed, this was found using transmission electron microscopy25 (TEM) where nanoparticles were clearly seen to be bound to the cell membrane (Figures 5A, left panels are the low magnification images, right panels being the higher magnification images of the corresponding left panels). Once bound to the plasma membrane, an obvious question is whether AuNPs disrupt the membrane, potentially resulting in depolarization and Ca2+ influx. However, TEM studies did not demonstrate any membrane disruption (Figure 5A). One plausible mechanism for AuNP action is the flipping of membrane areas by these particles. Uptake may also involve lipid rafts, pinocytosis and other plasma membrane mechanisms. Indeed, previous studies have found that modulation of nanoparticle surface properties can influence the mechanism of intracellular uptake (i.e., endosomal, passive diffusion).26-28 However, the extremely fast membrane depolarization and rapid uptake of AuNPs that was observed in our study need to be reconciled with the relatively slow rate of such uptake processes. Regardless, the findings of the present study will help to better define the biology of cell-nanoparticle interactions and help engineer nanoparticles to modulate cellular functions of interest. For example, varying surface charge density or combining positive and negative charges on the same nanoparticle may allow for graduated cellular uptake, targeting toward specific intracellular organelles, as well as control of the extent of change in [Ca2+]i and other effects, thus balanced unintended cytotoxicity versus targeting mechanisms of interest. Acknowledgment.SupportedbyNIHCA135011,CA136494, and UTMD-1 Grants (P.M.) and by NIH Grants HL090595 (C.M.P.), GM077173 (V.M.R.), and HL088029 (Y.S.P.). Available. Materials and methods; additional data. This material is available free of charge via the Internet at http://pubs.acs.org. </doc> ###", "role": "user" }, { "content": "{\"context\": {\"CellAge\": null, \"CellSource\": null, \"CellTissue\": null, \"CellType\": null, \"CoatFunctionalGroup\": null, \"HumanAnimal\": null, \"Material\": null, \"Shape\": null, \"SizeInMediumNm\": null, \"SurfaceCharge\": null}, \"measurements\": [{\"h\": 24.0, \"mgL\": null, \"viab\": 28.349}, {\"h\": 24.0, \"mgL\": null, \"viab\": 24.611}, {\"h\": 24.0, \"mgL\": null, \"viab\": 21.028}]}", "role": "assistant" } ]	5.json

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