- Nano Express
- Open Access
Serum Proteins Enhance Dispersion Stability and Influence the Cytotoxicity and Dosimetry of ZnO Nanoparticles in Suspension and Adherent Cancer Cell Models
© Anders et al. 2015
- Received: 27 July 2015
- Accepted: 5 November 2015
- Published: 17 November 2015
Agglomeration and sedimentation of nanoparticles (NPs) within biological solutions is a major limitation in their use in many downstream applications. It has been proposed that serum proteins associate with the NP surface to form a protein corona that limits agglomeration and sedimentation. Here, we investigate the effect of fetal bovine serum (FBS) proteins on the dispersion stability, dosimetry, and NP-induced cytotoxicity of cationic zinc oxide nanoparticles (nZnO) synthesized via forced hydrolysis with a core size of 10 nm. Two different in vitro cell culture models, suspension and adherent, were evaluated by comparing a phosphate buffered saline (PBS) nZnO dispersion (nZnO/PBS) and an FBS-stabilized PBS nZnO dispersion (nZnO – FBS/PBS). Surface interactions of FBS on nZnO were analyzed via spectroscopic and optical techniques. Fourier transformed infrared spectroscopy (FTIR) confirmed the adsorption of negatively charged protein components on the cationic nZnO surface through the disappearance of surfaced-adsorbed carboxyl functional groups and the subsequent detection of vibrational modes associated with the protein backbone of FBS-associated proteins. Further confirmation of these interactions was noted in the isoelectric point shift of the nZnO from the characteristic pH of 9.5 to a pH of 6.1. In nZnO – FBS/PBS dispersions, the FBS reduced agglomeration and sedimentation behaviors to impart long-term improvements (>24 h) to the nZnO dispersion stability. Furthermore, mathematical dosimetry models indicate that nZnO – FBS/PBS dispersions had consistent NP deposition patterns over time unlike unstable nZnO/PBS dispersions. In suspension cell models, the stable nZnO – FBS/PBS dispersion resulted in a ~33 % increase in the NP-induced cytotoxicity for both Jurkat leukemic and Hut-78 lymphoma cancer cells. In contrast, the nZnO – FBS/PBS dispersion resulted in 49 and 71 % reductions in the cytotoxicity observed towards the adherent breast (T-47D) and prostate (LNCaP) cancer cell lines, respectively. Presence of FBS in the NP dispersions also increased the reactive oxygen species generation. These observations indicate that the improved dispersion stability leads to increased NP bioavailability for suspension cell models and reduced NP sedimentation onto adherent cell layers resulting in more accurate in vitro toxicity assessments.
- Zinc oxide nanoparticles
- Dispersion stability
When prepared in nanostructured form, many materials develop novel and unique physicochemical properties leading to their use in nanotechnology applications such as quantum dots and MRI contrast agents for imaging , biosensors that use surface plasmon resonance , and nanomedicinal approaches that employ hyperthermia and photothermal therapy . Efficient use of NPs and nanocarriers in biomedical applications, such as delivery of therapeutic and imaging agents into tumor sites and genetic materials into cells, has been successfully demonstrated in recent years [4–6]. Interestingly, cytotoxicity is a property that is inherent in many of these commonly used materials when they are synthesized with their crystallite size in the nanoscale range [7–9]. This has raised significant scientific and public concerns on the environmental and health effects of every nanomaterial before employing them in nanotechnology applications. It is therefore necessary to evaluate the cytotoxic response of nanomaterials and the underlying mechanisms of nanomaterial toxicity to accurately address the growing concerns about the impact of the unwanted cytotoxicity of nanomaterials.
Numerous studies reported in the literature have shown that NP-induced toxicity depends on the nanomaterial properties such as crystallite size [10, 11], shape , charge , and chemical phase . Some studies have also shown that the toxic response of a given nanomaterial might vary with the specific cell/organism type such as prokaryotes versus eukaryotes , gram-positive bacteria versus gram-negative bacteria, and normal cells versus cancerous cells . These observations suggest that the role of organism-specific factors such as surface structure of cells [17, 18] and the proliferation status [10, 16] also influence the toxic response. Although the cytotoxicity of nanomaterials is an unfavorable feature for many applications, it could become very useful for certain therapeutic applications, if the toxicity is cell- or organism-specific. The cytotoxicity reported for most nanomaterials does not display a cell-specific or organism-specific behavior. However, many groups [16, 19, 20], including the authors, have demonstrated that certain ZnO NP formulations show a strong preferential cytotoxicity to cancer cells of identical lineage. For example, while resting primary human immune cells (T lymphocytes) showed no appreciable effect when treated with ≤5 mM concentrations of ZnO NPs, cancer cells of the same lineage showed significantly increased NP-induced toxicity resulting in approximately 28–35-fold increases in NP sensitivity [10, 16]. Thus, toxicity studies are extremely important both to assess the environmental and health impacts of NPs and to modify the nanomaterial design to make them safer, as well as to develop novel nanomedicinal approaches utilizing cell-specific cytotoxicity.
Considering the large number and types of new nanomaterials being developed and coming to the market, in vitro studies are essential in assessing the toxicity in high-throughput systems as they provide rapid and cost-effective screening. In most in vitro cytotoxicity experiments, cells are cultured in a culture medium to which a prepared dispersion of the NPs in a suitable solution is introduced. In contrast, in vivo toxicology studies frequently employ NPs dispersed in a suitable medium that are subsequently injected into the blood vessels or tissues of the animal model. If successful, the dispersed NPs will reach tumor sites and achieve deep penetration and entrapment in the cancerous tissue by means of the enhanced permeability and retention (EPR) effect, characterized by increased vascular permeability and compressed lymphatic networks in solid tumors . However, making highly stable and efficient dispersions of nanoparticles to achieve efficient nanoparticle delivery for both in vitro and in vivo applications has been a challenging task. A major difference that NPs possess compared to the well-studied conventional toxic chemicals (which are well soluble molecules and ions) is the difficulty in dispersing them in suitable solutions and culture media and to maintain them as stable dispersions for sufficient durations . For most in vitro studies, NP suspensions should remain stable at least for 24 h while in vivo studies require stability for much longer durations . Formation of large agglomerates and their sedimentation over longer periods often prevents a significant fraction of the NPs from participating in efficient interactions with the test cells and the resulting cytotoxic response.
Nanoparticle dispersion stability is a complex interplay between the agglomerate size of NP complexes in dispersion (hydrodynamic size) and their sedimentation behavior over time . Factors that affect NP stability include NP size [25, 26], surface structure [18, 27], capping agents , pH , ionic strength , organic and protein components [26, 27, 30], and free ions in the solution system . Although it is now known that the physicochemical properties of the nanomaterial, the specific cell/organism type, and the membrane properties [10, 17, 18] of the test organism play a crucial role in the resulting cytotoxic response, very few studies have focused on the important role of NP dispersion characteristics and dispersion stability and the effects of NP agglomeration and sedimentation in the resulting toxicity [32, 33]. Furthermore, Cho et al.  have recently shown that the cellular uptake of gold NP in breast cancer cells measured using conventional upright in vitro cell culture configuration was significantly higher than that obtained when an inverted setup was used, suggesting the effect of NP sedimentation and diffusion in the media play a significant role in the NP-cell interaction. In addition, suspension stability has been shown to directly influence effective administered dose of NPs to adherent cells grown in vitro [22, 32, 33].
When NPs are introduced to biological fluids such as human plasma, a biomolecular corona consisting of proteins and small molecules rapidly forms on the NP surface [27, 35–37]. Since the blood circulatory system is the most probable treatment administration option for NPs into the human body, it is important to investigate how adsorption of blood proteins on NPs will affect the cellular response, and more importantly, the differential cytotoxicity of ZnO NPs between cancerous and normal cells. The effect of having serum in culture medium and/or adsorption of the serum proteins on some NP systems such as silica , carbon nanotubes , graphene oxide , and ZnO [41, 42] has been studied recently. In all these cases, the primary effects of serum addition included improved dispersion stability and a clear reduction of the NP-induced cytotoxicity. Interestingly, the abovementioned studies were conducted using adherent cell models; therefore, comprehensive studies are needed to determine if similar responses are present for suspension cell models as well. Hypothetically, increases in the NP-induced toxicity in suspension cell models would be observed as NP sedimentation decreases, thereby improving the bioavailability of the NPs to the cells.
In this study, we investigated the use of FBS as a ZnO NP coating agent and the effects of this FBS coating the surface structure of the uncoated NPs, the dispersive stability of the NPs in solution, and the effective dosimetry of the NPs in cellular toxicity assays. Since NP dispersion stability and dosimetry is highly dependent upon a balance of NP diffusion and sedimentation velocities within a solution [32–34, 43, 44], the use of multiple suspension and adherent cell models was necessary to fully characterize any observations attributable to the addition of the FBS. A complete NP characterization profile was employed to fully interrogate NP surface changes as a result of the FBS coating. In addition, we examined the agglomeration and sedimentation behavior of the FBS-coated ZnO NPs (nZnO – FBS) compared to uncoated ZnO NP (nZnO) in biologically relevant solutions. Finally, we investigated the effects that FBS coating would have on the solubility of the nZnO, the NP-induced toxicity, and nZnO-induced reactive oxygen species production. As a whole, this work provides a comprehensive look at effects of FBS as a dispersion stabilizing agent in nanotoxicology studies.
Synthesis and Characterization of nZnO
The nZnO sample used in this study was produced using the forced hydrolysis of Zn acetate dehydrate precursor in diethylene glycol (DEG). The solution was held at 150 °C for 90 min. Nanopure water was added to the solution at 80 °C to obtain the desired crystallite size. This synthesis method allows for excellent control of the crystallite size by carefully regulating the temperature parameters and the hydrolysis ratio of the reaction media . Once cooled to room temperature, the nZnO were separated from solution via centrifuging at 20,000 rpm and subsequent washings with ethanol. A dried FBS-coated (nZnO – FBSdried) sample was prepared by sonicating the nZnO with FBS for 10 min. The resulting suspension was then centrifuged, and the resulting pellet rinsed several times with nanopure water to remove any unbound FBS. The pellet was dried overnight at 60 °C to create a powdered nZnO – FBSdried form for material characterization. Both NP samples were thoroughly characterized and investigated in detail using X-ray diffraction (XRD), transmission electron microscopy (TEM), zeta potential measurements, UV-vis spectrophotometry, and Fourier transformed infrared spectroscopy (FTIR). X-ray diffraction (XRD) spectra were recorded at room temperature on a Philips X’Pert X-ray diffractometer with a Cu Kα source (λ = 1.5418 Å) in Bragg-Brentano geometry. The loose powder samples were leveled in the sample holder to ensure a smooth surface and mounted on a fixed horizontal sample plane. Lattice parameters and crystal size were analyzed with Rietveld refinement using Materials Analysis Using Diffraction (MAUD) software, corrected for instrumental broadening . TEM and high-resolution TEM analysis was carried out using an FEI Tecnai and FEI Titan, respectively. The operating voltage of the Tecnai microscope was 120 and 300 kV on the Titan. Image processing was carried out using the Digital Micrograph software from Gatan (Pleasant, CA, USA). Room-temperature optical spectra in the ultraviolet and visible light wavelength ranges were collected using a CARY 5000 spectrophotometer. Zeta potentials of the powdered samples of nZnO were measured in nanopure water as a function of pH with a Malvern Zetasizer NanoZS. The temperature was equilibrated to 25 °C, and the pH was varied in the 6 to 12 range using 1.0 N HCl and 1.0 N NaOH prior to collecting the data. At least six data collections per run were performed on three separate aliquots of the ZnO suspension for each sample. The Zetasizer unit was also used to measure the hydrodynamic size of nZnO aggregates dispersed in selected media. Several solution systems were used for NP stock preparation and analyzed during this experiment.
NP Stock Preparation
To achieve the most stable dispersion for each solution, the critical material-specific delivered sonication energy (DSEcr) for the ZnO NPs was predetermined for the calorimetrically calibrated JSP Super-sonic bath sonication device employed for this study. A DSEcr of 181 J/ml was established through established protocols outlined by Taurozzi et al. (2011)  and employed in multiple dosimetry studies [33, 47]. To achieve the needed DSEcr, ZnO NPs were suspended at concentrations of 25 mM and sonicated in the bath sonicator delivering a power of 1.05 W for a total 30 min. Several biologically relevant solutions were prepared per manufacturer’s instructions unless otherwise defined. They include nanopure water, PBS, FBS, Roswell Park Memorial Institute (RPMI) medium (RPMI 1640 buffered with sodium bicarbonate and Hepes, pH adjusted to 7.3), RPMI-based cellular medium (RPMI 1640 medium supplemented with 10 % FBS and 2 mM l-glutamine, 1.5 g/l sodium bicarbonate, 4.5 g/l glucose, 10 mM HEPES, 1.0 mM sodium pyruvate and penicillin-streptomycin), and DMEM-based cellular medium.
In addition to single solution NP dispersions, several NP stock dispersions were prepared with FBS. Based on most of the reported protocols found in the literature and our own preliminary data, 10 % FBS addition was found to provide optimal results and was used to prepare these stock dispersions. The stock dispersions involved first suspending the desired quantity of NPs in a volume of FBS equivalent to 10 % of the final solution volume and sonicating at 1.05 W for 10 min. After sonication, the FBS suspension was diluted to the calculated volume with the desired medium (90 % of the total volume) and re-sonicated at 1.05 W for an additional 20 min. These stock dispersions are referred to as either nZnO – FBS/water or nZnO – FBS/PBS for stocks prepared in nanopure water or PBS, respectively. Additionally, two nZnO – FBS formulations were created by removing the excess FBS prior to re-suspension and sonication in the appropriate solution. One formulation involved centrifuging the nZnO/FBS suspension after the initial sonication period followed by several washes with nanopure water to remove any unbound FBS. The remaining pellet was then re-suspended in the desired solution to the appropriate concentration to create nZnO – FBSwashed/water or nZnO – FBSwashed/PBS for nanopure water and PBS dispersions, respectively. Finally, stock dispersions for the dried sample, described in the previous section, are designated as nZnO – FBSdried/water and nZnO – FBSdried/PBS for nanopure and PBS dispersions, respectively.
For extracellular dissolution experiments, nZnO stock dispersions were prepared as previously described at 6 mM concentrations. To simulate cellular assay conditions, an aliquot of each stock dispersion was added to cellular media at a final nZnO concentration of 0.6 mM. All dispersions were stirred continuously to prevent nZnO sedimentation. At desired time points, an aliquot of the sample was removed and centrifuged at 5000×g for 20 min using an Amicon™ Ultra-4 Centrifugal Filter Unit with a 3-kD molecular weight cutoff (0.1 nm pore size) to remove any undissolved nZnO while allowing free zinc ions to pass through. Quantitative analysis of the dissolved Zn2+ ions was conducted on a Thermo X Series 2 quadrupole inductively coupled plasma mass spectrometer (ICP-MS) under normal operating conditions (i.e., no CCT) with the Xt cone set. Instrument performance was evaluated and optimized for each run. The instrument was calibrated using multi-element calibration standards containing Zn, Cr, Mn, Fe, Co, and Ni in 2 % HNO3 at concentrations of approximately 1, 10, 100, and 1000 ppb. Instrument drift was monitored and corrected using 20 ppb indium as an internal standard introduced online.
Dosimetry Analysis and Empirical Deposition Fractions
Since NPs suspended in solution often form loosely packed agglomerates consisting of NP clusters and entrapped media, two empirical methods, the Harvard Volume Centrifugation Method (VCM) [32, 33] and the in vitro sedimentation, diffusion, and dosimetry (ISDD) computational model , were employed to determine the effective NP density of the agglomerates (ρ e, g/cm3), agglomerate porosity parameter (ε agg, unitless), diffusion rate (D, m2/s), sedimentation velocity (V, m/s), deposition factor, (α, h−1), and the deposition fraction curve [f D (t)]. Harvard VCM, which utilizes experimental methods to measure the effective volume of NP pellets (V pellet), was employed to estimate the ρ e of the NP agglomerates. To measure the V pellet, NP suspensions were prepared at a concentration of 100 μg/ml in nanopure water and sonicated with 1.05 W of power for 30 min. The resulting stock dispersions were introduced to RPMI-based cellular media at a concentration of 1.25 mM and then centrifuged at 2000×g for 1 h in TPP (Techno Plastic Products, Trasadingen, Switzerland) packed cell volume tubes, and the resulting NP pellets were measured utilizing the TPP “easy read” measuring device by the same manufacturer.
Cell Culture and Cytotoxicity Studies
For cell cytotoxicity assays, Jurkat T cell leukemia, Hut-78 T cell lymphoma, T-47D epithelial mammary gland carcinoma, and the LNCaP epithelial prostate cancer cell line (ATCC, Rockville, MD) were cultured in cellular medium. Additionally, the T-47D media was supplemented with 0.2 units/ml bovine insulin. Cells were maintained in log phase at 37 °C and 5 % CO2 and seeded at a concentration of 5 × 105 cells/well in 96-well plates for Jurkat and Hut-78 cells. For the T-47D and LNCaP cells, the cells were first trypsinized using 0.25 % trypsin/0.53 mM EDTA (MP Biomedicals, LLC; Santa Ana, CA), re-suspended in fresh media, and then seeded at a concentration of 2.5 × 105 cells/well in 24-well plates. The cells were then incubated overnight to allow the cells to re-adhere to the plate. Prior to treatment, the growth medium was gently aspirated from each well and replaced with 400 μL of fresh medium. Cells were subsequently treated with freshly sonicated (30-min) NPs reconstituted in the desired solution. Jurkat cells and Hut-78 cells were treated with NPs immediately after plating. For example, for a 2 mg/ml ZnO NP stock dispersion, 3.2 mg of NP was re-suspended in 1.58 ml of solution medium and sonicated at 50 W for 30 min. Then working dilutions were prepared from the NP stock dispersion and added to 200 μl of cell suspensions in 96-well plates or 400 μl of cell suspensions in 24-well plates and cultured for 24 h. After 24 h, the effects of the various NP stock dispersions on cell viability were determined using two different viability assays, alamar blue and flow cytometry. The alamar blue metabolic assay was employed to assess viability based on the fluorogenic redox indicator dye resazurin being converted to the fluorescent resofurin upon reduction by mitochondrial enzymes in metabolically active cells. In this assay, cell populations were seeded into the desired plate at the concentrations indicated above, treated with NPs for 20 h, and then incubated with alamar blue (10 % of total volume) for an additional 4 h. Changes in fluorescence were evaluated spectrophotometrically using excitation/emission at 530/590 nm in a Biotek Synergy MX® plate reader (Winooski, VT).
Flow cytometry was employed as an alternate viability assay to verify experimental results. NP-treated cells were dually stained with a fluorescein isothiocyanate-labeled anti-HLA ABC antibody and 50 μg/ml propidium iodide (PI) to monitor losses in membrane integrity. Fluorescent CountBright counting beads (Invitrogen, Carlsbad, CA) were added to samples to enable determinations of absolute cell numbers, and changes in PI staining were used to quantify cell death. Nanoparticles were excluded from analysis based on the absence of fluorescence signal and light forward scatter (FS) and side scatter (SSC) characteristics, and samples were analyzed using a BD FACS Caliber flow cytometer.
For CD4+ T cell cytotoxicity experiments, written informed consent was obtained from all blood donors and the University Institutional Review Board approved this study. Peripheral blood mononuclear cells were obtained from healthy human blood samples via Ficoll-Hypaque density centrifugation (Histopaque-1077, Sigma, St. Louis, MO) and CD4+ T cells isolated by negative immunomagnetic selection as previously reported . Purified CD4+ T cells (>95 % purity) were cultured in RPMI/10 % FBS and suspended at a final concentration of 1 × 106 cells/ml. CD4+ T cells were then incubated for 24 h with increasing amounts of each of the nZnO suspensions (24–800 μg/ml) in order to establish an IC50. CD4+ T cells were then stained and viability analyzed by flow cytometry.
Reactive Oxygen Species (ROS) Detection
Mitochondrial superoxide levels were determined by flow cytometry using MitoSOX Red (Invitrogen, Eugene, OR) staining. MitoSOX Red is a cell permeable fluorescent dye that becomes oxidized by interacting with mitochondrial superoxide . The resulting fluorescent signal occurs upon binding of the oxidized dye to nucleic acids. As a positive control, samples were treated with 0.2 nM of the mitochondrial electron transport inhibitor antimycin-A to ensure that the cells were capable of producing ROS. Jurkat cells were incubated with 0.4 mM nZnO or nZnO-FBS for 6, 18, and 24 h and then dually stained with CD3 antibody and 2.5 μM MitoSOX Red for 30 min. Mitochondrial superoxide levels were determined by flow cytometry.
Statistical analyses were performed using Jump Pro 10 software (SAS; Cary, NC). The cytotoxicity data were analyzed using repeated measures of variance with post hoc comparisons and significance levels defined as p < 0.05 to determine statistical differences between the means and allow within-sample variation to be separated from between-sample variation. Data for Figs. 10, 11, and 12 were analyzed using a two-way analysis of variance (ANOVA) to test for statistical significance of the model, and post hoc comparisons were used to test for statistically significant effects of treatment on cell viability (p < 0.05) with an asterisk denoting statistical significance in the indicated figures.
ZnO Nanoparticle Synthesis and Characterization
The FTIR spectra of nZnO and nZnO – FBSdried are shown in Fig. 2b. Both samples contain a strong band characteristic of Zn-O vibrations  at 456 cm−1 (nZnO) and 451 cm−1 (nZnO – FBSdried). The nZnO sample exhibits two bands commonly associated with the carboxyl functional group, specifically 1412 cm−1 [νs(COO−)] and 1597 cm−1 [νas(COO−)] [53, 54]. Additionally, weaker bands consistent with τ(CH2) and ν(C–OH) at 903 and 1076 cm−1, respectively, indicate the presence of surface-adsorbed carboxyl groups originating from the zinc acetate precursor or the DEG solvent [52–54]. Furthermore, the absence of the ν(C=OOH) band at ~1720 cm−1 suggests that the carboxyl groups are in an expected deprotonated form as a result of the alkaline synthesis procedure . Noticeably absent in the nZnO – FBSdried samples are the strong symmetric and asymmetric stretching modes of the carboxyl group with new bands appearing in the spectrum at 1646 and 1533 cm−1 representing the amide I and amide II regions that arise from vibrations of the peptide bonds of the protein backbone [55, 56]. The amide I region, often used for secondary structure prediction, exhibits a maximum at 1650–1656 cm−1 when α-helices predominate in the protein secondary structure . At 1646 cm−1, the amide I band of the nZnO – FBSdried particles is close to this wavenumber maximum and suggestive of the presence of a protein with the α-helical structure components found in BSA. The slight shift of this maximum is consistent with a more random secondary structure (1644–1648 cm−1) and has been observed in protein adsorption and binding studies with albumin . Taken together, these findings clearly establish the displacement of adsorbed carboxyl groups and strong binding of the FBS protein components [58, 59] on the nanoparticle surface.
Agglomeration and Sedimentation Profiles
As previously discussed, determining the best media components and dispersion conditions for the preparation of stable nZnO dispersions was a major goal of this study. Reported cellular toxicity studies using Jurkat and Hut-78 cells have frequently employed a variety of different biological solutions used to disperse NP powder samples prior to treatment. Considerations in choosing stock solution components include maintaining a viable environment for the cells as well as creating a stable NP dispersion for treatment conditions. Commonly used stock solutions consist of nanopure water, PBS, or un-supplemented cellular media. In addition, various capping agents  or protein components are often utilized to reduce steric attractions between charged particles and solution components and reduce NP agglomeration [51, 60, 61]. Solutions such as PBS or un-supplemented media are generally preferred as they do not result in unwanted pH changes during the toxicity study or affect cellular osmolality. Likewise, media proteins such as BSA and FBS are frequently used as stabilizing agents.
Reactive Oxygen Species
Cellular Toxicity Studies
Our previous work has shown that ZnO NPs display some selectivity in their toxic response against cancerous T cells as the IC50 values observed for primary quiescent T cells under similar conditions are considerably higher. Similar toxicity studies using nZnO – FBS/PBS dispersions also showed stronger toxic responses (p = 0.0303) to resting CD4+ T lymphocytes (Fig. 11d). Here, the IC50 values decreased from 4.69 ± 0.85 mM when treated with nZnO/PBS dispersion to 2.43 ± 0.0.24 mM when nZnO – FBS/PBS dispersions were used. Thus, the effect of coating NP with serum proteins is evident in primary cells as well. While these results are consistent with our previously reported preferential cytotoxicity of nZnO to cancer cells, the therapeutic index of Hut cancer cells decreased from 22.6 for nZnO/PBS dispersions to 17.0 for the nZnO – FBS/PBS dispersion. Even with the modest decrease noted in the therapeutic index for the nZnO – FBS/PBS dispersion, the resulting therapeutic efficiency of the nZnO appears better than some commonly recognized traditional chemotherapeutic agents using comparable in vitro assays . Thus, the protein interaction with nZnO increases their cytotoxicity in the suspension cell models evaluated, yet maintains some of the cancer cell selectivity previously observed.
In comparison to the suspension cell models, FBS coating leads to opposite effects in adherent cells. Suspension cells interact with fewer NPs in experimental conditions consisting of even minimally unstable dispersions since a portion of the original NP concentration sediments and may not be bioavailable for cellular interactions. This will skew toxicity assay results to suggest lower toxicity for suspension cells than expected. When the stability of the NP dispersion is improved by suitable molecular additives such as 10 % FBS (Fig. 11), the cytotoxicity increases significantly for both Jurkat and Hut cancer cells as well as for the primary T cell lymphocytes. More ZnO NPs are freely available in the stable NP dispersions to interact with more cells, thus resulting in the higher cytotoxicity observed in suspension cell models. The stability-induced differences between suspension and adherent cells underscore the importance of this work. The results from suspension cell studies alone would seem surprising considering other reports employing similar serum coatings describe decreases in NP-induced cytotoxicity in adherent cell lines suggesting that NP coating could be a potential means to control unwanted cytotoxicity . One such study, Hsiao and Huang , demonstrated that 5–10 % FBS coating on ZnO NPs increased dispersion stability and Zn2+ ion release, and decreased the NP-induced cytotoxicity. They hypothesized that this reduction in cytotoxicity in the serum-coated NPs was a result of serum-induced increases in cell growth and resulting decreases in nZnO dosage-per-cell ratios. Additionally, these authors  showed larger Zn2+ release in their experiments with FBS-coated ZnO thus negating Zn2+ release as a primary contributing mechanism to nZnO cytotoxicity. In this present study, FBS coating appears to decrease the Zn2+ release while significantly increasing NP-induced cytotoxicity for the suspension cell model when compared to uncoated nZnO. With the addition of the 10 % FBS to our phosphate-rich stock dispersion, it is plausible that decreases in formation of toxic insoluble Zn3(PO4)2 precipitates  could translate to perceptible decreases in the NP-induced cytotoxicity observed for nZnO – FBS/PBS treatment. However, decreases in Zn3(PO4)2 formation would also result in measured increases in Zn2+ release which was not observed. Therefore, extracellular Zn2+ dissolution is unlikely to be a primary factor for the increased cytotoxicity noted for our nZnO – FBS NPs. It is clear from FTIR and zeta potential studies that FBS molecules bind to the ZnO NP surface. This modification of the NP surface influences their interaction with other NPs (and on agglomerate formation and sedimentation effects) as well as with cells.
Modeling and Dosimetry Studies
Given the toxicity trend differences noted for the differing cell models, mathematical models were investigated as a potential method to confirm the observed trends and to model the dosimetry kinetics for both the investigated stock dispersions (nZnO/PBS versus nZnO – FBS/PBS) and the different cellular models. All of the investigated experimental parameters of dispersion stability can be applied to the in vitro sedimentation, diffusion, and dosimetry (ISDD) computational models, put forth by Hinderliter et al. (2010), to predict the transport properties of the NPs in various experimental conditions . The NP and experimental parameters which most influence the NP transport properties include hydrodynamic size (d H), agglomeration state, effective particle density (ρ e), and the media height in the cell well (L) . Central to the effective dosimetry determination in the ISDD model is the calculation of a NP deposition factor α (h−1), Eq. 2 in “Experimental Details”, which interrelates the diffusion rate (D), sedimentation velocity (V), and media height [32, 33, 43, 44]. While the diffusion rate is primarily dependent upon the hydrodynamic size, the sedimentation velocity depends upon both the hydrodynamic size and the effective particle density occupied by NP agglomerates [43, 44]. The effective particle density (ρ e) can be approximated in the ISDD model (Eq. 7) using the hydrodynamic size-dependent agglomerate porosity parameter (ε agg)  or can be determined experimentally (Eq. 1) as a part of the Harvard Volume Centrifugation Method (VCM) [33, 43]. The Harvard VCM employs an experimental determination of NP agglomerate volumes to directly determine effective particle density. Furthermore, the VCM has been modified to incorporate mass lost due to the dissolution of soluble or partially soluble NPs such as ZnO [32, 33].
Delivered dose metrics
Stock dispersion in complete RPMI
Zeta potential (mV)
d H (nm)
ρ e (g/cm3)
(L = 6.6 mm)
(L = 2.25 mm)
t 90 (h)
t 90 (h)
−16.1 ± 1.45
−15.1 ± 0.81
−13.9 ± 0.40
−8.1 ± 1.93
−9.8 ± 2.08
−14.7 ± 1.46
−13.6 ± 2.63
The influence of FBS on the dosimetry kinetics is evident in an analysis of the experimentally stable water, FBS/water, and FBS stock dispersions introduced to the RPMI-based cellular media. Given the dependence of agglomerate behavior and effective density (ρ e) on the hydrodynamic size of NPs in dispersion, the water (182.5 nm), FBS/water (185.1 nm), and FBS (185.5 nm) stock dispersions should exhibit similar dosimetry kinetics. However, the effective density (ρ e) of the NPs in the three dispersions decreased with increasing amounts of FBS in the stock preparation thus increasing the calculated deposition factor. These disparities are a direct result of measured increases in the effective volume of the NP pellet with increasing amounts of FBS in solution. So while the similarities in the hydrodynamic diameters of these three dispersions imply consistent NP diffusion rates (Eq. 9), the differing effective volume measurements suggest different NP sedimentation velocities (Eq. 8) . Therefore, even though all three dispersions demonstrate dispersion stability experimentally, the effective volume occupied by the resulting protein corona in the NP agglomerates decreases the sedimentation velocities of the NPs predicted by the model, thus increasing the deposition factor and shifting the dosimetry kinetics.
Interestingly, the dose metrics obtained for the unstable stock dispersions of PBS and RPMI suggest dosimetry trends opposite to what would be expected considering the experimentally determined agglomeration and sedimentation profiles. However, the VCM dosimetry model assumes that the dispersions used to obtain the experimental α values are stable across the time frames defined by the model. As this is not the case with the PBS and RPMI stock dispersions, these obtained dosimetry curves cannot be used to accurately model the dosimetry kinetics under these conditions. To model potentially unstable dispersions, it is necessary to calculate α values using experimentally determined stability parameters such as hydrodynamic size at multiple time points. For this, we employed the ISDD model. This model approximates effective particle density utilizing the hydrodynamic size-dependent parameter ε agg. While the approximation of the ρ e employed in this model lacks the experimental accuracy achieved for ρ e measurements afforded by VCM, the ISDD model does allow for time-resolved approximation of NP transport parameters within unstable dispersions. Employing the ISDD model, α values were obtained using time-resolved d H values and the dosimetry curves for both the nZnO/PBS and nZnO – FBS/PBS stock dispersions in RPMI-based cellular media were modeled for several time points from 0 to 24 h. These time-resolved points were compiled to generate ISDD curves which were then fit to both the f D (t) and f A (t) mathematical models to obtain α values for each representative graph (Fig. 13c, d).
Studies reported in this work show that ZnO nanoparticles form very large agglomerates with hydrodynamic sizes in the 300 to 20,000 nm range which and depend primarily on the solution composition used. Presence of such agglomerates leads to sedimentation of a significant portion of the nanoparticles which increases in time. For adherent cells, this will increase the effective NP concentration interacting with the cells at very early time points and may cause unusually large and potentially inaccurate in vitro toxicity measurements. This was demonstrated using two adherent epithelial cell lines (T-47D and LNCaP) with both cell types experiencing significant decreases in NP-mediated cell death with the more stable NP stock dispersions. Likewise, this sedimentation process will reduce the effective NP concentration remaining in the solution medium. Subsequent investigations using suspension cells showed that this phenomenon results in an opposite trend due to the reduction in the number of nanocrystals interacting with the suspension cells and, consequently, a lower than actual toxic response is obtained. Using a 10 % FBS/PBS stock to disperse the same nZnO, the IC50 values for Jurkat (Hut) cancer cells decreased from 0.39 mM (0.21 mM) for nZnO/PBS-treated cells to 0.27 mM (0.14 mM) for the nZnO – FBS/PBS dispersion. Their normal counterparts (CD4+ T lymphocytes) also showed a reduction in IC50 with 10 % FBS addition. Its IC50 decreased from 4.69 mM for nZnO/PBS dispersions to 2.43 mM when nZnO – FBS/PBS stock was used. On the other hand, treating the adherent cells T-47D and LNCaP using the nZnO – FBS/PBS dispersion resulted in 48.7 and 71.4 % reductions, respectively, in the NP-induced toxic response compared to nZnO/PBS dispersion. Surface modification of these nZnO using serum proteins seems to be a suitable approach to obtain stable dispersions and accurate cytotoxicity values, thus improving their utility in therapeutic applications.
This study has shown the important role of serum proteins in modifying the ZnO nanocrystal surface resulting in the formation of considerably smaller agglomerates and stable NP dispersions. Furthermore, these FBS surface coatings influenced Zn2+ release from the nanocrystals and induced changes in the mitochondrial function of the cells as evidenced by the increased ROS production in the nZnO – FBS/PBS-treated cells. Given these findings, it is readily apparent that the addition of serum proteins adds a level of complexity to our experimental understanding of NP-cell interactions. Further studies are needed to examine how FBS coatings change NP-cellular surface interactions, influence cellular uptake mechanisms, and modify molecular interactions with other extracellular matrix components. Additionally, these studies should be extended to include newer 3D cellular culture models which often employ complex biomolecular solution systems or molecular scaffolds to better mimic in vivo-like environments for mechanistic biological assays [71, 72]. These complex culture systems will influence the formation of the NP biomolecular corona and change the diffusion and sedimentation rates for NP dispersions especially those containing additional serum proteins. Furthermore, 3D culture systems alter cellular growth patterns, metabolism, genetic expression, and phenotype organization changing NP-induced cellular responses which could be further influenced by the addition of NP coating materials [72, 73]. Finally, these studies should be extended into other cell types. Molecular coatings are often considered as a means to achieve safer nZnO structures, yet little is known about how these molecular coatings will influence NP-cellular interactions with endothelial cells [18, 27, 74] and other primary immune system cells .
This work was supported in part by NSF CBET 1134468, NSF EAGER DMR-1137419, ARO W911NF-09-1-0051, and NIH 1R15CA141358 grants. The authors thank Dr. Marion Lytle and the Biotrace Lab at Boise State University for the ICP-MS use. Additionally, we thank Prof. Ben McMorran and CAMCOR for TEM images, CAMCOR’s Titan and Tecnai are supported by W.M. Keck Foundation, ONAMI, the Air Force Research Laboratory (agreement number FA8650-05-1-5041), and University of Oregon Office of Research.
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