Reaction of monocytes to polystyrene and silica nanoparticles in short-term and long-term exposures.

Volume 3 Number 2 March 2014 Pages 69–154

Toxicology Research www.rsc.org/toxicology

ISSN 2045-452X

PAPER Eleonore Fröhlich et al. Reaction of monocytes to polystyrene and silica nanoparticles in short-term and long-term exposures

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Reaction of monocytes to polystyrene and silica nanoparticles in short-term and long-term exposures† Maria Mrakovcic,a Claudia Meindl,a Eva Robleggb and Eleonore Fröhlich*a Nanoparticles (NPs) are increasingly used in industrial, health and consumer products. In addition to the intended effects, NPs may also cause cell damage. Typical cytotoxicity assays assess short-term effects in adherent cells but do not evaluate longer exposure times and do not focus on cells in suspension. Since NPs are not removed easily from the organism, non-biodegradable NPs may persist in the systemic circulation and affect monocyte function at low concentrations. To mimic this situation, THP-1 monocytes were exposed to low concentrations of plain polystyrene particles (PPP) in different sizes for short (24 h) and prolonged (16 d) time periods. CELLine CL350, a small two-chamber bioreactor, and sub-culturing in flasks were compared regarding prolonged cytotoxicity testing. Uptake rates of the particles, cytotoxicity screening assays, and interleukin secretion were used for the identification of adverse effects. After 24 h, 50 µg ml−1 20 nm PPP did not affect cellular viability and interleukin secretion, while at higher concentrations the cytotoxicity of PPP (20 nm–500 nm) was correlated to surface area. After 16 d of exposure at 50 µg ml−1 20 nm PPP, the decrease in cell number and the increase in interleukins were significant. 200 nm PPP, by contrast, caused only minimal effects. Due to lower reproducibility, CELLine proved to

Received 27th December 2013, Accepted 8th January 2014

be less suitable for the assessment as compared to sub-culturing in flasks. After prolonged exposure,

DOI: 10.1039/c3tx50112d

silica Aerosil OX50 particles also were more cytotoxic towards THP-1 monocytes. The data suggest that prolonged exposure to NPs leads to cytotoxicity at low doses and that induction of cell death may be

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involved in the observed pro-inflammatory action of NPs.

Introduction Nanoparticles (NPs) are used in a variety of industrial, consumer, and medical products. Although the quantities of NP that are taken up into the organism might be small, a reduced potential for elimination could lead to accumulation and potential damage of cells and organs. Animal experiments showed that repeated exposure to non-biodegradable NPs, such as gold NPs and magnetic NPs, caused histo-pathological changes in various organs, weight loss and changes in blood count.1–3 Clearance of NPs from the organism is low4,5 and it cannot be excluded that exposure over a long period to low concentrations of NPs may interfere with cell function. Environmental studies show that people exposed to higher

a Center for Medical Research, Medical University of Graz, Stiftingtalstr. 24, Graz, Austria. E-mail: [email protected]; Fax: +43 316 385 73009; Tel: +43 316 385 73011 b Institute of Pharmaceutical Sciences, Department of Pharmaceutical Technology, Karl-Franzens-University of Graz, Helmholtzstr. 46, Graz, Austria † Electronic supplementary information (ESI) available. See DOI: 10.1039/ c3tx50112d

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levels of particulate matter for a prolonged time show a greater tendency to develop atherosclerotic lesions (e.g. ref. 6 and 7). Animal experiments suggest an important role of monocytes in this pro-atherosclerotic effect.8 Studies on long-term toxicity of NPs are mostly performed in vivo in order to overcome the limitations of cell cultures, such as the study of only one cell type, lack of interactions between cell types, lack of metabolisation and excretion, etc. On the other hand, cellular mechanisms can be better studied in vitro because the system is less complex and parameters can be better controlled. For cytotoxicity screening, usually adherent cells are exposed for periods up to 72 h. This exposure scenario, however, might not be relevant for prolonged contact with NPs. In such a situation, cell damage can be more severe than acute exposure due to accumulation of adverse events. On the other hand, cytotoxicity could also be lower when cellular repair mechanisms are activated. Previous experiments on the endothelial cell lines EAhy926 showed a time-dependent decrease in cell viability to 50% of the control cells after 28 d in the presence of plain polystyrene particles (PPP) in a bioreactor. Multi-walled carbon nanotubes decreased cell numbers dose-dependently to 75–40% at day 7 followed by cell

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recovery to 90–70% of the control cells.9 This suggests that cells are able to adapt to the exposure with NPs. When sub-culturing of cells was performed, the observed effect was much smaller and cell numbers decreased to maximally 80–70% of untreated control cells. Using also sub-culturing, Thurnherr et al. demonstrated the absence of cytotoxicity for multi-walled carbon nanotubes in the adherent cell line A549 and in the non-adherent T-cell line Jurkat-6 for up to 3 months.5 This technique might be less sensitive for the detection of cytotoxicity in adherent cells because cells may compensate for cell loss by proliferation, while contact inhibition in non-treated control cells may decrease proliferation. For the assessment of cytotoxicity in blood, hemolysis and testing of peripheral blood monocytic cells can be used. While erythrocytes cannot be cultured for longer time periods, different protocols allow the maintenance of monocytes for several weeks.10 These prolonged cultures are used to induce differentiation into mesenchymal cells, particularly osteoblasts.11 The use of monocytes from peripheral blood has the disadvantage of inter-donor variations. In addition, culture of monocytes and lymphocytes leads to variable cell loss and a variable degree of differentiation to macrophages and to plasma cells, respectively. Due to the higher degree of reproducibility, cell lines are used to study cytotoxicity in leukocytes. Cytotoxicity testing over prolonged periods of exposure is further complicated by the fact that no established methods are available. To mimic the prolonged contact of NPs with leukocytes, a system where medium could be changed without interfering with the cell–NP interaction would be desirable. In this study, CELLine, a small-size disposable bioreactor, was evaluated for its suitability for chronic cytotoxicity testing of non-adherent cells. The CELLine system has been used for antibody production by hybridoma cells12–15 and for antitrypsin production by transgenic rice cells16 and provided an optimal environment for these cells. To identify the method for the assessment of NP effects on monocytes after prolonged exposure, PPP were used as model particles. Data obtained using CELLine were compared with a more conventional mode of testing, where cells were grown in the presence of NPs and transferred to larger vessels when the

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controls reached densities indicated for sub-culturing. In order to find out whether the increased sensitivity to long-term exposure is also valid for other NPs, silica NPs were assessed with the more reliable of the two culture systems, the sub-culturing technique. Aerosil® fumed silica NPs are used in a variety of food applications, such as anti-caking agents in spices, seasonings, egg- or milk powder, and cappuccino, and may represent an example of chronic exposure by the oral route.17 Remains of silica NPs from other industrial applications (paint, anti-blocking agents, dental composites, PET-films, etc.) in the environment may also cause effects by long-term exposure to these particles.

Materials and methods Particles For short-term and long-term exposures, plain polystyrene particles (PPP) of 20 and 200 nm (Thermo Scientific) were used. Particle uptake was investigated using red fluorescently labelled PPP of 25 nm, 200 nm, and 500 nm (FluoroMax red, Thermo Scientific). Silica particles Aerosil OX50 were obtained from Degussa. PPP and Aerosil OX50 particles were applied to cells suspended in cell culture medium (DMEM + 10% FBS) after sonication in an Elmasonic S40 water bath (ultrasonic frequency: 37 kHz, 40 W, Elma) for 20 min. The CELLine bioreactor The two compartments of the CELLine bioreactor CL350 (Integra Biosciences) (Fig. 1) are separated by a semipermeable membrane that enables exchange of nutrients from the upper medium compartment and waste products from the lower cell compartment. A gas-permeable bottom membrane allows the exchange of O2 and CO2 in the cell compartment. The cell compartment contains 10 ml of medium, the medium compartment 200 ml. Physicochemical characterization Particles were characterized in terms of size and zeta potential using a ZetaSizer Nano-ZS (Malvern Instruments). Particles

Fig. 1 Schematic drawing of the bioreactor CELLine. Cells, proteins, and nanoparticles are retained in the cell compartment, while small molecules, such as glucose or lactate, pass through the 10 000 MW dialysis membrane to the medium compartment. Gas exchange takes place through the gas permeable silicone membrane at the bottom of the flask.


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were diluted in DMEM + 10% FBS to a concentration of 200 µg ml−1. After equilibration of the sample solution to 25 °C, hydrodynamic sizes were measured with a 532 nm laser and a detection angle of 173°. Dynamic fluctuations of light scattering intensity caused by Brownian motion of the particles were evaluated. The zeta potential was measured by Laser Doppler Velocimetry (scattering angle of 17°) coupled with Photon Correlation Spectroscopy and was calculated out of the electrophoretic mobility by applying the Henry equation. Time-dependent particle changes Changes in particle size and surface charge were recorded in suspensions of particles (PPP: 50 µg ml−1; Aerosil OX50: 12.5– 100 µg ml−1) in DMEM + 10% FBS. After sonification the solutions were kept for various periods of time at 37 °C. Size and surface charge were measured again on a ZetaSizer Nano-ZS using the same protocol. Leakage of dye To identify a potential leakage of the indicator dye from the particles, suspensions of particles (20 µg ml−1) in DMEM + 10% FBS were prepared. After sonification they were kept for various periods of time at 37 °C. Subsequently, suspensions were centrifuged at 16 000g for 30 min in a Juan KR25i centrifuge. Fluorescence was read on a fluorescence plate reader (FLUOstar Optima, BMG Labortechnik) at 544 nm/612 nm and the obtained value was related to the fluorescence of the freshly prepared sonicated solution as 100%. Detection of endotoxin A PYROGENT Ultra (sensitivity = 0.06 EU ml−1, Lonza) was used for the endotoxin testing. Each sample dilution was tested in duplicate, and the different endotoxin standards with E. coli strain 055:B5 in triplicates. The assay was performed first as a yes/no test and the samples with positive endotoxin detection were further tested via a dilution series to quantify free endotoxin. The assay was performed according to the instructions given in the manual. A positive reaction is characterized by the formation of a firm gel that remains intact momentarily when the tube is inverted (vertical rotation of 180°). A negative reaction is characterized by the absence of a solid clot after inversion. Determination of cellular dose THP-1 cells were exposed to 20 µg ml−1 fluorescently labelled PPP for 24 h. Medium height was 3 mm. Cells were centrifuged at 800g for 10 min, washed and centrifuged again. Fluorescence of the cells and of serial dilutions of the exposure solution was measured on a fluorescence plate reader (FLUOstar Optima) at 544 nm/612 nm. The exposure solution was diluted with cell suspensions instead of medium alone to account for autofluorescence or quenching effects caused by the cells. The amount of fluorescence in the cells was related to total fluorescence added. For the calculation of ingested particle numbers, data provided by the manufacturer (http://www.distrilab-particles.com/filters/pdf/Bul120%27A%27%20FluorPoly.pdf) were used.

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Short-term exposure THP-1 cells (Cell Line Services) were cultured in RPMI 1640 growth medium, supplemented with 10% FBS, 200 mM L-glutamine, and 1% penicillin/streptomycin. For cytotoxicity screening, cells were plated at a density of 5 × 104 cells per 96-well plate and were immediately exposed to nanoparticles suspended in DMEM + 10% FBS. For the determination of interleukins, 5 × 105 THP-1 cells were seeded per well of a 24-well plate. Long-term culturing in CELLine The initial cell density suitable for culturing over a time-period of 4 weeks was determined in a pilot experiment. THP-1 cells were inoculated with 10 × 105 cells per 10 ml, 5 × 105 cells per 10 ml, and 2.5 × 105 cells per 10 ml in the cell compartment of the CELLine bioreactor CL-350 and were cultured in a conventional incubator (HeraSafe). The medium compartments were filled with 100 ml growth medium. For the actual experiments, 2.5 × 105 cells in 10 ml were inoculated. The medium in the cell chamber was immediately supplemented with NPs at final concentrations of 20 µg ml−1 and 50 µg ml−1 of PPP20, 50 µg ml−1 of PPP200, as well as with 2% EtOH. An additional 100 ml of growth medium was added to the medium chamber. All cultures were studied for at least 3 weeks. Once a week, the cells were counted and the medium from the medium compartment was exchanged. The medium from the cell compartment was not changed at any time-point. The cellular densities and viability were assessed on an automated cell counter based on electrochemical sensing.18 An aliquot of the cells was removed from the cell compartment and was placed in a 50 ml tube. An aliquot of this solution was used for counting in the CasyTT (Inovatis). The following cursor setting was used: calculation cursor left/right (10.00/ 48.88), normalization cursor left/right (5.86/48.88), and dilution factor 1 : 100. Conventional long-term culture (sub-culturing) THP-1 cells were seeded in 6 well plates at densities of 5 × 105 cells per well. The NPs were added immediately at the final concentration of 20 µg ml−1 and 50 µg ml−1 for PPP and 12.5, 25, 50 and 100 µg ml−1 for Aerosil OX50. Cells not exposed to NPs and cells treated with 2% EtOH were used as negative and positive controls, respectively. At each time point, the cells were counted and all cells were transferred into a larger cell culture flask (25 cm2, 75 cm2, and 175 cm2), where the treatment schedule was continued. The experiment was stopped after 16 days when proliferation of the control cells in the 175 cm2 flasks declined. Cytotoxicity by formazan bioreduction The CellTiter 96® Aqueous Non-Radioactive Cell Proliferation Assay (Promega) was used according to the manufacturer’s instructions. 200 µg ml−1 20 nm and 200 nm carboxyl polystyrene nanoparticles (Invitrogen) suspended in DMEM and


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known to decrease viability to 10 ± 5% and 96 ± 9% of controls were used as positive and negative controls, respectively. 20 µl of the combined MTS–PMS solution was added to 100 µl of each well and plates were incubated for 2 hours at 37 °C in the cell incubator. Absorbance was read at 490 nm on a plate reader (SPECTRA MAX plus 384, Molecular Devices).


Lactate dehydrogenase (LDH) release Samples of the entire observation period were assessed together. To make sure that storage of the samples did not cause a decrease in enzyme activity, stability tests were performed with commercial LDH. For a time-period of four weeks, LDH from bovine heart (Sigma Aldrich) was diluted to 0.002–0.2 U ml−1 in DMEM + 10% FBS. To one half of the samples a 2% BSA solution was added to improve the storage conditions. The samples were stored at −20 °C and −80 °C.19 Storage at −80 °C for 4 weeks retained the activity at 100% and 98% in the presence and absence of BSA, respectively. Storage at −20 °C caused a loss of activity to 65% in the presence of BSA and to 59% in the absence of BSA. Therefore, −80 °C and addition of BSA were chosen for the storage of the samples. After four weeks, the frozen samples were thawed on ice and then all samples were pre-warmed to RT. The stability of the enzyme was assessed by the CytoTox-ONE™ Homogeneous Membrane Integrity Assay (Promega). 25 µl of the supernatant was transferred into a 96 well plate suitable for fluorescence detection. Then, an equal volume of the substrate mix was added. The plate was incubated for 10 min at RT protected from light. The reaction was stopped and fluorescence was read on a FLUOstar Optima fluorescence photometer (BMG Labortechnik) with an excitation/emission wavelength of 544/ 590 nm. Lysis was performed by addition of 0.1% Triton X-100 and 70% ethanol for 10 min at RT. Caspase 3/7 activation The Caspase-Glo® 3/7 Assay (Promega) was used according to the instructions given by the provider. An equal volume of assay reagent was added to the medium and the plate was then incubated at RT for 30 min protected from light. Subsequently, the supernatant was transferred to a 96 well plate suitable for luminescence detection and was measured on a LUMIstar Optima (BMG Labortechnik). Apoptosis was induced by 1 µM Staurosporine (Sigma-Aldrich) as positive control. Interleukin secretion in THP-1 cells Untreated cells and LPS treated cells (100 and 1000 ng ml−1) were used as negative and positive controls, respectively. After incubation, the cells were centrifuged at 1000g for 4 min. The supernatant was then used for further investigations. Secretion of interleukins (IL) 1β, 6, and 8 was assessed by an ELISA assay (BD Biosciences). The assay was performed according to the provided instructions. First, ELISA plates were coated with the respective capture antibody and were incubated overnight at 4 °C. Thereafter, the plates were washed 3 times with >300 µl 1× wash buffer at RT and were blocked with 100 µl assay diluent for 60 min at RT. Subsequently, plates


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were washed 3 times. Meanwhile, the standards were prepared, to range from 0 pg ml−1 to either 200 pg ml−1 (IL-8), or 250 pg ml−1 (IL-1β), or 300 pg ml−1 (IL-6). 100 µl standards and samples were incubated for 2 h at RT. After five rinses, 100 µl detection antibody coupled to horseradish peroxidase was added to each well and incubated for 60 min at RT. The last washing step was repeated 7 times. Next, 100 µl of a substrate solution was added to each well and incubated for 30 min at RT protected from light. Upon addition of 50 µl stop reagent, the blue coloured substrate solution turned yellow. Then, the OD was determined on a SPECTRA MAX plus 384 at a wavelength of 450 nm. Statistics Data were analyzed with one-way analysis of variance (ANOVA) followed by the Tukey-HSD post hoc test for multiple comparisons (SPSS 19 software). p < 0.05 was regarded as significant. Trend analysis was performed using Excel 2010.

Results Physico-chemical characterization of PPP Sizes of PPP when suspended in water were very similar to the values indicated by the producer. When dispersed in DMEM + 10% FBS, however, the size of 20 nm PPP increased markedly to 74 nm, while 200 nm and 500 nm PPPs showed only small increases in size (Table 1). Surface charge of all PPPs was strongly negative in water and slightly negative in medium + 10% FBS. At a concentration of 100 µg ml−1 all PPP endotoxin levels were below the detection limit. Changes in size and surface charge were recorded at certain time points over 14 d to identify changes of particles in the medium. Surface charge was very constant at −12.6 ± 0.9 mV for 20 nm PPP, −12.9 ± 0.7 mV for 200 nm PPP, and −13.3 ± 0.6 mV for 500 nm PPP. Time-dependent changes in size were more prominent. On the first day of incubation, sizes of 46–62 nm were recorded for 20 nm PPP. From the second day onward, sizes varied between 101 and 140 nm. Sizes for 200 nm PPP varied between 232 and 240 nm on the first day. Subsequently, sizes of 197–387 nm were recorded. 500 nm PPP measured 530–621 nm on the first day and 652–879 nm the following days. No trend for a timedependent increase in size was identified. The stability of the fluorescent labelling was studied by monitoring leakage of the dye. Only 200 nm and 500 nm PPP can be pelleted under the given conditions, 20 nm PPP would need 3 × 106 g to settle 10 cm h−1 (http://www.bangslabs.com/ sites/default/files/bangs/docs/pdf/Particle%20Dr%20Q%20&% 20A.pdf ). Fluorescence in the supernatant related to the noncentrifuged solution was 4–3% for the 200 nm PPP and 5–4% for the 500 nm PPP after 1–3 days of exposure at 37 °C. As fluorescent dyes show self-quenching when close together (inside the polystyrene bead) the values obtained in the solutions most likely overestimate the amount of leaked dye.20 Aerosil OX50 particles were 40 nm large according to information provided by the manufacturer (http://www.aerosil.com/

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Table 1 Sizes and surface charges of plain polystyrene particles (PPP). Sizes given by the producer and parameters when suspended in water and in DMEM + 10% FBS according to the photon correlation spectroscopy and laser Doppler velocity are indicated

Non-fluorescent labeled PPP

Fluorescent labeled

Particle

Size (nm), producera

Size (nm), water, PCS

ζ-Potential (mV), water, LDV

Size (nm), medium, PCS

ζ-Potential (mV), medium, LDV

Size (nm), producerb

Size (nm), medium, PCS

ζ-Potential (mV), medium, LDV

PPP20 PPP200 PPP500

21 200 500

23 212 523

−37.1 −57.6 −58.7

74 224 568

−11.7 −8.08 −10.3

28 200 490

58 230 584

−10.3 −13.5 −13.5


a

Source: http://www.polysciences.com/SiteData/docs/Microparti/dbdc94c1bcbdf576/Microparticles%20Guide.pdf. www.distrilabparticles.com/filters/pdf/Bul120%27A%27%20FluorPoly.pdf.

www2/uploads_all/text/TI_1271_AE_DIN_A4_e_Apr05.pdf, #1606). Suspended in DMEM + 10% FBS, two peaks at sizes of 35–50 nm and of 522–731 nm on the first day of incubation were observed. The relationship between the two peaks was concentration-dependent: at 12.5 µg ml−1 40% of the particles were detected in the first peak and a smaller fraction was present as aggregates, at 100 µg ml−1 only 10% of the particles were present as single particles and the majority as aggregates. Irrespective of the particle concentration, surface charge ranges between −9.6 mV and −11.6 mV. Time-dependent changes in the size distribution were in the same order of magnitude (35–56 nm and 435–748 nm) and surface charge varied between −8.5 mV and −12.3 mV. Short-term effects: uptake, cytotoxicity, and interleukin secretion of PPP Fluorescence measurements after 24 h of exposure showed that all THP-1 cells contained fluorescently labelled PPP of all sizes (Fig. 2a–c). Quantification by fluorometry showed that between 7 ± 1.6% and 1.8 ± 0.6% of the particles added to the cultures were ingested (Table 2). In a viability dose–response curve with all PPP, different effects of the particles were seen at 100 µg ml−1. This concentration could be used to correlate uptake to viability loss, while at higher doses viability in PPP20 exposed cells was zero and at lower doses no differences were seen between 200 nm and 500 nm PPP. Viability at 100 µg ml−1 was related more closely to ingested particle number than to ingested particle amount (Fig. 2e and d). The coefficient of determination (R2 value) was similar for the correlation of viability with particle sizes determined by photon correlation spectroscopy in medium (R2 = 0.96) and with particle size as given by the manufacturer (R2 = 0.96) (Fig. 2f and g). 20 µg ml−1 and 50 µg ml−1 20 nm PPP caused no significant decrease in the viability according to the MTS assay (Fig. 3a). At higher concentrations (200 µg ml−1) of 20 nm PPP, viability decreased to 12% (data not shown). Caspase 3/7 activation and LDH-release, however, were already significantly increased at 50 µg ml−1 20 nm PPP (Fig. 3b and c). The maximum amount of IL-6 released from treated cells was 10 times lower than the release of IL-8 (15 pg ml−1 vs. 223 pg ml−1 after stimulation with LPS). At 20 µg ml−1 and 50 µg ml−1 of 20 nm and 200 nm PPP, no increase in IL-8 secretion

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b

Data source FluoMax: http://

was seen (Fig. 3d). Similarly, IL-6 levels were not changed by particle exposure (ESI, Fig. S3†). Based on the absence of significant loss of viability, 20 µg ml−1 and 50 µg ml−1 were used for long-term exposures. Since 200 nm and 500 nm PPP both showed no cytotoxicity in short-term exposures, only 20 nm and 200 nm PPP were assessed in long-term exposures. Establishment of CELLine cultures To determine which cell number allowed culturing over 4 weeks, CELLine bioreactors were inoculated with different numbers of THP-1 cells. The maximal culturing period of 28 days was reached in cultures where 2.5 × 106 cells were inoculated in the 10 ml cell compartment. The initial application of 5 × 105 cells resulted in a reduced culturing period by approximately 2 weeks. Inoculation of 1 × 106 cells yielded the shortest culturing period of 7 days (data not shown). Variability of the growth rates was also linked to cell density. If 2.5 × 105 cells were inoculated, a total of 15 ± 15.8 × 106 cells were recorded after 7 d. After inoculation of 1 × 106 cells 381 ± 76 × 106 cells were counted in CELLine at this time point. Relative variation in the cell number was 110% for the low cell number inoculation and 20% for the higher cell number inoculation. After the first maximum of cell density was achieved cell densities fluctuated. Long-term exposure in CELLine: uptake, cell numbers, and interleukin secretion of PPP Data of 7 independent experiments showed that long-term culturing of THP-1 cells in CELLine bioreactors was highly variable. The variability was mostly due to differences in growth curves of untreated cells between the experiments (Fig. 4). Maximal cell densities between 3.9 × 107 and 6.5 × 107 cells per ml were obtained after 21–24 d, and after 24 d of culture, cell numbers decreased. Therefore, cultures were evaluated up to 21 d. While after 24 h most THP-1 cells contained PPP (Fig. 2a–c), only a few cells contained PPP at later time points and too low for quantification (data not shown). Reduction in cell numbers was highest for 2% EtOH ( positive control), followed by reductions caused by 50 µg ml−1 20 nm PPP, 20 µg ml−1 20 nm PPP and 50 µg ml−1 200 nm PPP (Fig. 5a).


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Fig. 2 Uptake and correlation to viability of fluorescently labelled 20 nm, 200 nm and 500 nm PPP after 24 h of exposure. Intracellular localization of 20 nm (a), 200 nm (b), and 500 nm (c) PPP. Correlation of cytotoxicity at 100 µg ml−1 with the amount of ingested particles in µg (d), particle number (e) and surface area of primary particles according to data (f ) from the producer and as characterized by PCS in the exposure medium (g).

Table 2

Uptake of fluorescent PPP after 24 h of exposure

Fluorescent particles

Cellular dose (% applied)

Cellular dose (µg)

Cellular dose (particles)

PPP20 PPP200 PPP500

7 ± 1.6 2.2 ± 0.8 1.8 ± 0.6

1.4 ± 0.7 0.4 ± 0.2 0.4 ± 0.1

2.3 ± 1.1 × 1012 1.9 ± 0.7 × 109 1.1 ± 0.3 × 108

The highest amount of released LDH was noted at day 7 with significant increases caused by 2% EtOH, followed by non-significant increases caused by 50 µg ml−1 PPP20 (Fig. 5b). Overall release of IL-8 was higher than that for IL-6. Here too, the highest amounts were detected 7 days after exposure. 2% EtOH increased IL-8 secretion about 50-fold, while 200 nm PPP had no impact at any time-point. Although 20 nm PPP increased IL-8 levels markedly, these changes were not statistically significant (Fig. 5c). Release of IL-6 by untreated and treated cells, in


general, was very low. The highest rates were detected 7 days after inoculation. Exposure to all PPP caused no significant changes; 50 µg ml−1 20 nm PPP induced a 2.7-fold increase and 200 nm PPP a 1.8-fold increase, while no increase was noted after exposure to 20 µg ml−1 20 nm PPP (Fig. S5, ESI†). Long-term exposure in sub-culture testing: uptake, cell numbers, and interleukin secretion of PPP Similar to exposures in CELLine, the fraction of cells containing PPP was markedly decreased compared to the 24 h exposures. Cell densities of approximately 140 × 106 cells per culture vessel were reached after 16 days. A slight reduction in cell numbers by approximately 15% was detected in cells exposed to 200 nm PPP, while cell numbers after exposure to 50 µg ml−1 20 nm PPP and 2% EtOH were reduced by 92% and 95%, respectively. No changes were observed upon exposure to 20 µg ml−1 of 20 nm PPP (Fig. 6a).

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Fig. 4 Variation of total cell numbers of baseline conditions (untreated THP-1 cultures) when inoculated with 0.25 × 105 cells between different experiments (n = 6).

Fig. 3 Short-term (24 h) effects of 20 µg ml−1 and 50 µg ml−1 20 nm PPP (abbreviated as 20/20 and 50/20) and 50 µg ml−1 200 nm PPP (abbreviated as 50/200). (a) Viability (formazan bioreduction) indicated as optical density (OD), CPS200: 200 µg ml−1 carboxyl polystyrene particles are used as a positive control. (b) Membrane integrity (LDHrelease) indicated as relative fluorescence units (RFU), lysis: 0.1% Triton X-100 + 70% ethanol is used as a positive control. (c) Apoptosis (caspase 3/7 activation) indicated as relative chemiluminescence units (RCU); Stau: 1 µM staurosporine is used as a positive control. (d) Interleukin 8 secretion. Abbreviation: co: control; LPS: 100 ng ml−1 lipopolysaccharide is used as a positive control. Significant changes are marked by asterisks.

Highest amounts of released LDH were detected with EtOH-treated cells and upon exposure to 50 µg ml−1 20 nm PPP. Releases by the other NP exposures were not significantly changed (Fig. 6b). The peak amount in released IL-8 of untreated cells was detected 5 days after inoculation. IL-8 secretion was increased upon exposure to 50 µg ml−1 20 nm PPP but no significant

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Fig. 5 Changes in cell number (a), LDH release (b), and IL-8 secretion (c) of THP-1 cells, treated with 20 µg ml−1 and 50 µg ml−1 20 nm PPP (abbreviated as 20/20 and 50/20), and 50 µg ml−1 200 nm PPP (abbreviated as 50/200) over time in CELLine culture. 2% Ethanol (EtOH) serves as a positive control. Abbreviation: RFU: relative fluorescence units. Significant changes are marked by asterisk.

increases in IL-8 levels caused by all PPP exposure were noted (Fig. 6c). Increases in IL-1β were parallel to the changes observed for IL-8 (Fig. S6, ESI†). IL-6 levels showed similar changes with only significant increases after exposure to 50 µg ml−1 20 nm PPP and 2% EtOH (Fig. S6, in ESI†).


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For experiments performed with CELLine, where great variations between the experiments were seen, changes of more than 50% of the control values were regarded as relevant. According to this definition, 50 µg ml−1 20 nm PPP caused cytotoxicity, necrosis and inflammation in both systems. In addition to that, after 7–8 d of exposure, 20 µg ml−1 20 nm PPP decreased cell number and increased LDH-release in CELLine, not in sub-culturing exposure. LDH-release and increased secretion of IL-6 were identified for 50 µg ml−1 200 nm PPP in CELLine. Long-term exposure in sub-culture testing: cell numbers and interleukin secretion of silica (Aerosil OX50) particles With the better protocol (sub-culturing), the effect of Aerosil OX50 particles was assessed. Cytotoxicity after 24 h of exposure was performed to identify not acutely cytotoxic concentrations. Significant reduction of viability was observed after exposure to 200 µg ml−1 Aerosil OX50 (Fig. 8). In contrast to PPP, also at higher concentrations, viability did not decrease below 50% (data not shown). Upon longer exposure times (≥8 days), 100 µg ml−1 Aerosil OX50 particles decreased cell numbers significantly (Fig. 9a).

Fig. 6 Changes in cell number (a), LDH release (b), and IL-8 secretion (c) of THP-1 cells, treated with 20 µg ml−1 and 50 µg ml−1 20 nm PPP (abbreviated as 20/20 and 50/20), and 50 µg ml−1 200 nm PPP (abbreviated as 50/200) over time in sub-culture testing. 2% Ethanol (EtOH) serves as a positive control. Abbreviation: RFU: relative fluorescence units. Significant changes are marked by asterisks.

Data comparison of PPP exposures by CELLine and sub-culturing In order to identify common trends in both systems, significant changes were highlighted for the sub-culturing (Fig. 7).

Fig. 8 Short-term cytotoxicity (24 h) of Aerosil OX50 particles in THP-1 cells. Abbreviation: PC: positive control (200 µg ml−1 carboxyl polystyrene particles are used as a positive control). Significant changes are marked by asterisks.

Fig. 7 Overview of time-dependent changes of cell number, LDH-release and IL-6 secretion induced by 20 µg ml−1 and 50 µg ml−1 20 nm PPP (PPP20), and 50 µg ml−1 200 nm PPP (PPP200) in CELLine culture and in sub-culture testing. Changes >50% of the control values for CELLine experiments and statistically significant changes in sub-culture testing are highlighted.


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Fig. 9 Changes in cell number (a), LDH release (b), and IL-8 secretion (c) of THP-1 cells, treated with 25 µg ml−1, 50 µg ml−1, and 100 µg ml−1 Aerosil OX50 over time in sub-culture testing. 2% Ethanol (EtOH) serves as a positive control. Abbreviation: RFU: relative fluorescence units. Significant decreases are indicated by asterisks and significant increases by hatch.

LDH-release was increased after 5 d only for exposures to 100 µg ml−1 Aerosil OX50, at 8 d for ≥25 µg ml−1, and at later time points for ≥50 µg ml−1 Aerosil OX50 (Fig. 9b). IL-8 secretion was increased at all time points after exposure to ≥50 µg ml−1 Aerosil OX50 particles (Fig. 9c). Changes in IL-1β levels were similar to those of IL-8 levels (Fig. S9, in ESI†), while IL-6 values were below the detection threshold for all exposures (data not shown). THP-1 cultures exposed to 2% EtOH showed such dramatic decreases in cell numbers that LDH-release and cytokine secretion were also decreased (Fig. 9).

Discussion Although similar trends were identified in both systems, CELLine proved to be less reliable in the identification of these effects than testing by sub-culturing. After 24 h of exposure, 50 µg ml−1 of all PPP did not act significantly cytotoxic in THP-1. At this concentration, however, markedly decreased cell numbers and increased LDH-release were recorded after longer exposure. This suggests that cell death by necrosis has occurred and that THP-1 cells reacted to

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prolonged contact with 20 nm PPP in a similar way to adherent cells.9 A similar trend was also observed after exposure to Aerosil OX50 particles. 24 h cytotoxicity of the PPP was most strongly linked to the surface of ingested intracellular particles, followed by particle number and much less to weight. According to in vivo experiments, surface area was recommended as a good measure for inhalation effects by several authors.21–23 Other authors did not report a dependence of toxicological effects from particle size and particle surface.24 Pulmonary toxicity of ZnO NPs correlated quite well with surface area, while particle number did not.25 Surface area of differently sized silica particles was also correlated to changes in metabolomics to a greater extent than particle number upon intravenous injection into mice.26 When iron oxide particles of 10 nm and 50 nm were compared in A3 lymphocytes, the small ones were more toxic based on weight and the larger ones more toxic based on surface area.27 In general, as stated by Wittmaack,28 surface area appears to be a good indicator of toxicity when similar materials are compared. Surprisingly, the correlation of cytotoxicity with particle surface area in this study was similar with data indicated by the producer and with measurements obtained by characterization in medium. One potential explanation might be that PPP are taken up as single particles after the protein coat is digested by membrane-associated and secreted serine proteases of THP-1 cells. THP-1 cells possess relatively high levels of metalloproteinases 1, 2 and 9 and of matriptase,29–31 which play an important role in monocyte migration.30 Alternatively, the presence of agglomerates occurring in protein-containing media could lead to greater reported particle sizes in the PCS measurements. This is because a single detection element collects light from all particles simultaneously, meaning that the estimate of particle size and size distribution is biased to a larger particle size.32 Several algorithms have been validated for the calculation of cellular doses with polystyrene particles, silica particles, and iron oxide particles (e.g. ref. 33 and 34) in adherent cells. Such algorithms have not been validated for phagocytising and nonphagocytising cells in suspension since suspension cells are rarely used for cytotoxicity screening. The capability for phagocytosis, however, is relevant because adherently growing phagocytising cells ingest larger amounts of particles than epithelial cells.35,36 Similarly, the THP-1 monocytes used in this study ingested about 7% of the 20 nm PPP, while nonphagocytising lymphocytes TK-6, under the same conditions, took up less than 1% (unpublished results). Ideally, cellular doses should be determined in parallel with the biological assays. Long-term exposure could decrease particle cytotoxicity by particle agglomeration. In this study, however, we did not observe such an effect. Despite an increase in hydrodynamic size of 20 nm PPP from ∼50 nm to ∼150 nm, these particles showed long-term cytotoxicity at concentrations where no short-term cytotoxicity was observed. The strong correlation of particle surface to cytotoxic effect identified in the short-term studies could explain the profound effect in the long-term exposures. The missing decrease of the cytotoxic effect despite


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an increase of hydrodynamic size, together with the finding that short-term cytotoxicity was closer correlated to nominal size than to hydrodynamic size, appears to suggest that THP-1 cells also came into contact with particles in their nominal size. In the present study, CELLine CL350 was assessed for its suitability for chronic cytotoxicity testing of cells in suspension. This bioreactor had been designed to allow fast proliferation of suspension cells with a high yield of antibody.14 The protocol recommends an inoculation of 1.5 × 106 cells for optimal growth in CELLine CL350, meaning 6 times the amount used in this study. On the other hand, the CELLine CL 6-well bioreactor was reported to be suitable for the toxicological assessment of adherent cells.37 In this study, highly variable results with THP-1 cells were obtained. The main reason for the limited use of CELLine of long-term cytotoxicity testing in this study was the short observation time when inoculated at conventional densities. When cells reached the maximum cell density, they could not be further evaluated. The theoretical smallest distance between two cells in suspension is 2 µm. In addition to that, the maximum cell number is determined by cell diameter. Maximum reported cell densities for suspension cells in static bioreactors, in general, range between 106 and 107 cells per ml.38 After optimization of the culture medium, the maximum cell density of 3 × 106 cells per ml HL-60 monocytes could increase to 6–7 × 107 cells per ml.39 Even higher maximum concentrations of more than 2 × 108 cells per ml CHO cells were reached in the WAVE Bioreactor™ system 20/50 in perfusion mode with disposable Cellbag™ bioreactors.40 In this study maximum densities between 4.5 and 7.5 × 107 cells per ml in a static bioreactor were recorded. Since THP-1 cells are larger than CHO cells, these cell densities can be regarded as very high, demonstrating that CELLine provided optimal growth conditions for these cells. To prolong the observation time, cell numbers for inoculation were decreased in this study. The inoculation number, however, also has a lower limit because cell death increases due to lack of survival factors. This cellular behaviour has been described for lymphocytes and for monocytes several times.41 Ma et al. identified 2 × 105 cells per ml as the lower threshold for normal survival of B cell chronic leukemia cells. In this study, inoculated cell numbers (2.5 × 105 cells in 10 ml cell compartment) were ten times lower. A variable rate of cell death at these low densities, therefore, could explain the high variations in the non-treated cultures. Since initial cell densities in the sub-culturing experiments were about 10 times higher than in CELLine and higher seeding densities in CELLine also produced lower variations in cell densities, more reproducible growth was obtained by sub-culturing. Unfortunately, seeding numbers for reproducible growth in CELLine studies cannot be employed for long-term studies because maximal cell densities were reached already after 7 d. After this time, cell numbers of untreated cells fluctuate (cycles of decrease and increase) and, therefore, cannot be used as a reference for the treated cells any more. Cytotoxicity testing in subculture in this study produced more reproducible results than studies in CELLine. However,


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this method cannot be generally recommended for chronic cytotoxicity testing because growth inhibition in untreated cultures may be overlooked. In this case, the sensitivity of the testing is dramatically decreased. Interleukin secretion of THP-1 cells in this study was used to identify potential pro-inflammatory effects. Increased secretion of pro-inflammatory interleukins by peripheral blood monocytic cells (PBMCs) has been reported upon short-term exposure to 5 nm and 28 nm Ag NPs (1L-1β42), PGLA NPs, and ZnO NPs (TNF-α and IL-843,44). In THP-1 cells, increases of IL-1β and TNF-α in THP-1 cells were higher upon exposure to SiO2 than to TiO2, ZrO2, and Co NPs.45 While in 24 h exposure to 20 µg ml−1 and 50 µg ml−1 of both PPP, no increase in interleukin secretion was noted, long-term exposures led to increases in IL-6 and IL-8 release after >5 d of culture (overview, Fig. 6). For most exposures, LDH-release and IL-8 release, induced by PPP and Aerosil OX50 particles, were increased in parallel (either as a trend or significantly). It may be suspected, therefore, that the detected pro-inflammatory effects of NPs are, in part, caused by cell death acting as an inflammatory stimulus. Cell death as a pro-inflammatory signal has been reported in the presence and absence of microbial infection.46 This immune activation is believed to serve as protection against infection, for elimination of transformed cells, and as a stimulus for faster tissue repair. Activation of the immune system by minimally increased levels of dying cells could cause adverse effects on chronic exposure to NPs. Similar to PPP, Aerosil OX50 particles also showed greater cytotoxicity after prolonged exposure. For these particles, however, both dose-dependent decreases in viability after 24 h of incubation and differences between 24 h and longer incubations were less pronounced. This might be due to the presence of agglomerates; at higher concentrations (≥50 µg ml−1), Aerosil OX50 particles were predominantly present as agglomerates (435–748 nm), while 20 nm PPP also at 200 µg ml−1 were around 75 nm and increased with time only to 101–140 nm.

Conclusions The bioreactor CELLine CL350 was not suitable for long-term toxicity testing of THP-1 cells because the low inoculation densities led to high variations in cell counts. If, on the other hand, cell numbers for reproducible growth were seeded, observation time was too short for long-term evaluation. The strong correlation of surface area to cytotoxic effect identified in the short-term testing may indicate that, although not acutely cytotoxic at relevant concentrations, particles with large surface area also bear a higher risk for long-term cytotoxicity. Activation of the immune system as a reaction to minimal cytotoxicity could play a role in chronic toxicity of NP. Such processes could also explain the observed promotion of atherosclerosis after long-term exposure of humans to inhaled nanoparticles.47,48 The higher toxicity of 20 nm PPP and of Aerosil OX50 particles at long-term than at short-term exposure suggests potential cellular accumulation and

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highlights the importance of long-term exposure testing. It appears that increased long-term cytotoxicity is inversely correlated to the extent to which the particles form aggregates.

Acknowledgements


This work was supported by the Austrian Science Fund grant P 22576-B18. We thank Ramona Baumgartner for particle characterization.

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