The Influences of Cell Type and ZnO Nanoparticle Size on Immune Cell Cytotoxicity and Cytokine Induction
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Nanotechnology represents a new and enabling platform that promises to provide a range of innovative technologies for biological applications. ZnO nanoparticles of controlled size were synthesized, and their cytotoxicity toward different human immune cells evaluated. A differential cytotoxic response between human immune cell subsets was observed, with lymphocytes being the most resistant and monocytes being the most susceptible to ZnO nanoparticle-induced toxicity. Significant differences were also observed between previously activated memory lymphocytes and naive lymphocytes, indicating a relationship between cell-cycle potential and nanoparticle susceptibility. Mechanisms of toxicity involve the generation of reactive oxygen species, with monocytes displaying the highest levels, and the degree of cytotoxicity dependent on the extent of nanoparticle interactions with cellular membranes. An inverse relationship between nanoparticle size and cytotoxicity, as well as nanoparticle size and reactive oxygen species production was observed. In addition, ZnO nanoparticles induce the production of the proinflammatory cytokines, IFN-γ, TNF-α, and IL-12, at concentrations below those causing appreciable cell death. Collectively, these results underscore the need for careful evaluation of ZnO nanoparticle effects across a spectrum of relevant cell types when considering their use for potential new nanotechnology-based biological applications.
KeywordsNanoparticle ZnO T lymphocyte Monocytes Cytokine Immunity Nanotoxicity
Nanotechnology permits the manipulation of matter at the nanometer scale, which enables precision engineering to control nanomaterial physiochemical properties as well as their interactions with biological systems. It is the alterations in electrical, magnetic, structural, morphological, chemical, and physical properties of nanomaterials, which are comparable in size to naturally occurring biomolecules, that makes them particularly attractive for pioneering applications in technological and biological applications [1–4]. Nanomaterials comprised of ZnO have received considerable attention in recent years because of their potential use in electronic and industrial applications arising from their wide band gap (3.36 eV) semiconductor properties . From a biological perspective, nanoscale ZnO materials are already being used in the cosmetic and sunscreen industry due to their transparency and ability to reflect, scatter, and absorb UV radiation, and as food additives [5, 6]. ZnO nanomaterials are also being considered for use in next-generation biological applications including antimicrobial agents, drug delivery, bioimaging probes, and cancer treatment [7–10]. Although bulk (micron-sized) ZnO is generally recognized as a GRAS substance by the FDA [5, 6], many benign materials can exhibit appreciable cellular toxicity when reduced to the nanoscale . Recent toxicological studies using certain engineered nanoparticles (NPs) have confirmed the potentially harmful effects due to their high surface area, unique physiochemical properties, and increased reactivity of the material’s surface [11, 12]. This has led to the term, nanotoxicology, which describes the relationship between nanoparticle physiochemical properties and toxicological effects on living cells and biological systems.
In this regard, several recent reports have demonstrated the toxicity of metal-oxide NPs to both prokaryotic and eukaryotic cellular systems [7, 10, 13–16], while the bulk micron-sized materials remain nontoxic. The majority of eukaryotic studies specific to ZnO NPs, however, have relied heavily on the use of immortalized cells lines, which are recognized to display altered sensitivities to foreign materials/chemicals due to alterations in metabolic processes and major genetic instabilities. To date, only limited studies have evaluated the toxicity of ZnO NPs on normal primary human cells and their potential immunomodulatory effects. Even more challenging is the fact that the cytotoxic response is very different between cell types and nanomaterial systems, making it difficult to develop predictive models without detailed and systematic investigations. Recent reports demonstrate that ZnO NPs display differential toxicity towards primary human cells depending upon their proliferation potential, with normal T lymphocytes that are stimulated to divide by signaling through the T cell receptor displaying significantly greater toxicity than quiescent nonproliferating cells of identical lineage . When these studies were extended to immortalized T leukemic and lymphoma cells using identical ZnO NPs, an even greater sensitivity to NP-induced toxicity was observed (~33-fold) . Thus, susceptibility to NP-induced cytotoxicity appears to be related to the proliferative capacity of the cell and may also be affected by other physiologically relevant parameters, including cell-NP electrostatic interactions and inherent differences in cellular endocytic/phagocytic processes that facilitate NP uptake. These variations in cytotoxic response indicate that a more detailed and carefully controlled evaluation of ZnO NP effects on multiple types of normal human cells is needed to better understand the biological consequences of NP exposure. In addition, it is increasingly being recognized that toxicity depends on nanomaterial characteristics including size, shape, and electrostatic charge, as well as materials composition [7, 10, 17, 18]. In this study, the evaluation of size-controlled ZnO NPs and their resulting toxicity on different cells comprising the human immune system is explored. In addition, mechanisms underlying the differential toxicity response are investigated, as well as effects of ZnO NPs to induce proinflammatory cytokine expression.
Preparation and Characterization of ZnO Nanoparticles
ZnO NPs utilized in all experiments were synthesized in diethylene glycol (DEG) via forced hydrolysis of zinc acetate . In brief, zinc acetate was dissolved in DEG, and then nanopure water was added under magnetic stirring. Subsequently, the system was heated at 160 °C under reflux for 90 min. After cooling, the resulting product was removed from DEG via centrifugation, and washed with ethanol several times before drying for 24 h at 50 °C, resulting in a powder sample. The sample crystal phase, crystallite size, and morphology were characterized via X-ray diffraction (XRD) and transmission electron microscopy (TEM) as previously described [13, 14]. The NPs were then weighed and reconstituted in phosphate buffered saline (PBS) solution to the desired stock concentration. After reconstitution, NPs were sonicated for 10 min and immediately vortexed prior to addition to cell cultures.
Isolation of Peripheral Blood Mononuclear Cells
For isolation of PBMC (peripheral blood mononuclear cells) and immune cell subsets, written informed consent was obtained from all blood donors. The University Institutional Review Board approved this study. PBMC were obtained via Ficoll-Hypaque (Histopaque-1077, Sigma, St Louis, MO) gradient centrifugation using heparinized phlebotomy samples . After removal of the leukocyte layer, cells were washed three times with Hank’s buffer (Sigma, St. Louis, MO) and resuspended at a final concentration of 1 × 106 cells/mL in RPMI-1640 (Sigma), containing 10% fetal bovine serum (FBS) and cultured at 37° C and 5% CO2.
Isolation and Culture of Primary CD14+Monocytes and CD4+T cells
PBMC were obtained by Ficoll-Hypaque density centrifugation as described above, and CD14+cells isolated from the mixed cell suspension. Negative immunomagnetic selection was performed according to the manufacturer’s protocol using a cocktail of antibodies directed against CD2, CD3, CD16, CD19, CD20, CD56, CD66b, CD123, and glycophorin A (StemCell Technologies, Vancouver, BC). For optimal cell recovery, PBMC were first blocked with anti-human CD32 (Fcγ RII) blocker reagent before labeling with the antibody cocktail. Purified CD14+monocytes populations were typically of >92% purity and >97% viable as assessed by flow cytometry and cultured in RPMI/10% FBS at 5 × 105cells/mL in 96-well plates at 37°C and 5% CO2.
CD4+T cells were purified from PBMC using negative immunomagnetic selection per manufacturer’s instructions, and a cocktail of antibodies directed against CD8, CD14, CD16, CD19, CD56, and glycophorin A surface markers (StemCell Technologies). Unlabeled T cells were collected (typically >97% purity and >95% viable as assessed by flow cytometry) and subsequently cultured in RPMI/10% FBS at a final concentration of 1 × 106cells/mL.
Methods of immunofluorescent staining and flow cytometric analysis were performed as previously described  using a four-color Epics XL flow cytometer (Beckman Coulter, Miami, FL). Cells were stained with fluorescently labeled antibodies (Beckman Coulter) for 30 min at 4 °C, washed two times, and immediately analyzed. Ten thousand events gated on the parameter of size (forward scatter, or FS) and granularity (side scatter, or SSC) were analyzed, and expression of the percentage of positively staining cells and the mean fluorescence intensity (MFI) determined by comparisons to isotype controls. Appropriate concentrations of each antibody were determined by titration for optimal staining prior to experimental use. In PBMC cultures, individual cell types were distinguished from one another based on differential antibody staining and on FS and SSC properties. T cells were defined as CD3+; T helper cells defined as CD3+, CD4+; naive T cells defined as CD3+, CD45RA+; memory T cells defined as CD3+, CD45RO; B cells defined as CD19+, CD3; NK cells defined as CD56+, CD16+, CD3−; and monocytes defined as CD14+, CD3−. To prevent indiscriminate antibody staining of monocytes via Fc receptors, 100 μL of heat-inactivated human AB serum was added to experimental samples immediately prior to staining.
Cell Viability Assays
Cell viability following NP treatment was assessed using two different assays. In the first assay, cell subsets were identified using fluorescently labeled antibodies, and viability was determined by staining with 50 μg/mL of propidium iodide (PI) to monitor losses in cell membrane integrity. Fluorescent CountBright counting beads (Invitrogen, Carlsbad, CA) were added to samples to enable determinations of absolute cell numbers, and flow cytometry was used to evaluate changes in PI staining and to quantify cell death. NPs were excluded from the analysis based on the absence of fluorescence signal and light FS and SSC characteristics.
The second viability assay employed the fluorogenic redox indicator dye, Alamar Blue. This dye becomes fluorescent upon reduction by mitochondrial enzymes in metabolically active cells. Cell populations were seeded into 96-well plates at 5 × 105cells/mL, treated with ZnO NP for 18 h, and 20 μL of Alamar Blue added to cultures for an additional 6 h. Changes in fluorescence were evaluated spectrophotometrically using excitation/emission at 530/590 nm.
To assay for NP-induced reactive oxygen species (ROS) production, cells were first obtained from whole blood treated with an ammonium chloride lysing solution (1.5 M NH4Cl, 0.1 M NaHCO3, 0.01 EDTA) to lyse red blood cells, and centrifuged for 10 min at 4 °C to remove erythrocytic debris. The white blood cells were then resuspended in phenol red-free RPMI to a final concentration of 1 × 106 cells/mL and incubated with ZnO NPs for 6 to 20 h. Cells were subsequently loaded with 5 μM of the oxidation-sensitive dye, 2′,7′-dichlorofluorescein diacetate (DCFH-DA, Invitrogen, Carlsbad, CA) for 20 min. Production of the oxidation product was evaluated using flow cytometry as previously described . White blood cell populations (i.e., CD3+ T cells and CD14+, CD3− monocytes) present in the samples were simultaneously evaluated based on FS and SSC gating and staining with appropriate fluorescently labeled antibodies. As a positive control for ROS production, control cells were loaded with DCFH-DA dye and activated with PMA (25 ng/mL) for 1 h. For studies described in Fig. 8, ROS production was evaluated in PBMC treated with differently sized ZnO NPs using a fluorescent microplate reader as previously described [21, 22]. Experimental methodology, including the use of the DCFH-DA dye to indicate ROS generation, was identical as described above.
To determine the role of ROS in NP-induced cell death, purified primary CD4+T cells and monocytes were seeded in a 96-well plate at 5 × 105cells/mL and pretreated with 5 mM of the ROS scavenger,N-acetyl cysteine (NAC, Sigma) for 4–6 h. Cultures were then treated with various concentrations of ZnO NP for 24 h, and viability was determined using the Alamar Blue cytotoxicity assay and a fluorescent microplate reader.
To investigate the effect of ZnO NPs on cytokine production, an ELISA (enzyme linked immunosorbent assay) was used. For determination of IFN-γ and TNF-α production, freshly isolated PBMC were cultured at 1 × 106cells/mL and treated with varying concentrations of 8nm ZnO NPs (0.05, 0.1 and 0.2 mM) for 38 h. For determination of IL-12 levels, PBMC cultures were either left untreated or pretreated with 1,000 U/mL of IFN-γ (Peprotech, Rocky Hill, NJ), to prime for the production of this cytokine, and then cultured with ZnO NPs for 24 h. After NP treatment, cell-free supernatants were harvested via successive 10-min centrifugations (2,000 rpm, 7,000 rpm and 13,000 rpm) and stored at −80 °C until analysis. ELISA was performed by the UMAB Cytokine Core Laboratory (Baltimore, MD), with all samples analyzed in triplicate.
All data were analyzed using SAS, Inc. software (Cary, NC). Data for Figs. 3,4, and6were analyzed using repeated measures of variance withpost hoc comparisons to allow within-subject variation to be separated from between-subject variation. Data for Figs. 5,7and8were analyzed using a two-way analysis of variance (ANOVA). In all cases, significance levels were defined asp < 0.05.
Results and Discussion
Synthesis and Size Control of ZnO Nanoparticles
T and B Lymphocytes are More Resistant to NP Toxicity Compared to Monocytes and NK Cells
Because adherent monocytes appear considerably more susceptible to NP-induced cytotoxicity compared to other immune cell subsets, additional experiments were performed using purified monocytes and lower concentrations of NP, to more accurately determine the IC50 value. In these experiments, monocytes were treated with varying concentrations of ZnO NPs, and viability was evaluated using the fluorogenic redox Alamar Blue cytotoxicity assay. In agreement with monocyte data obtained from PBMC cultures, an IC50 of ~0.30 mM was observed (Fig. 3b). The Alamar Blue cytotoxicity assay was also used to confirm the IC50 using purified CD4+ T cells, and a similar IC50 value of ~5.4 was observed (data not shown). As previously reported by our laboratory [13, 14, 23], control experiments using bulk micron-sized ZnO powder showed no appreciable toxicity effect at any of the concentrations tested (e.g., viability with bulk ZnO: 96 ± 3% at 1 mM, 93 ± 3% at 10 mM), demonstrating that toxicity is limited to nanoscale ZnO. In addition, no appreciable toxicity was observed using NP-free supernatants (e.g., 98% viability with NP-free supernatant equivalent to 1–10 mM), indicating that the toxicity is likely not due to dissolved Zn ions from NP preparations.
Although these results indicate that monocytes are considerably more susceptible to NP-induced cytotoxicity than other immune cell types tested, it is important to note that these differences may be related to distinctions in cell-culture conditions. While T cells, B cells, and NK cells grow as suspension cultures, cultured monocytes grow as an adherent monolayer. These differences in growth characteristics may act to increase the effective ZnO NP concentration in adherent cultures. It is also plausible that the inherently greater capacity of adherent monocytes to phagocytose foreign materials, including NPs, may underlie their greater sensitivity, and the inherent cytolytic activity of NK cells against foreign pathogens, and altered self-cells may contribute to their greater sensitivity compared to lymphocyte populations. To address these possibilities, future experiments involving three-dimensional cell culture systems and those evaluating the extent to which phagocytic/endocytic mechanisms contribute to cell-type differences are needed. Nevertheless, given the variable susceptibilities of different immune cell types to ZnO NPs, it seems clear that careful analysis of in vitro cellular systems, followed by appropriate in vivo studies, is necessary to provide thorough and possibly predictive screening data regarding the relative toxicity and immunomodulatory effects of ZnO NP.
Memory and Naive T cells Differ in Cytotoxic Response to ZnO NP
Our previous findings that ZnO NP toxicity is dependent on the cell activation status, with quiescent T cells being more resistant to ZnO NP-induced cytotoxicity than identical cells activated to divide via stimulation through the T cell receptor , led us to evaluate whether “memory” T cells display greater sensitivity to ZnO NPs than “naive” T cells. During an immune response, the activation of T cells to a specific antigen found on a pathogen results in a cascade of intracellular signaling events, and to the differentiation of naive T cells into memory cells. Once memory cells have formed, they can become activated to proliferate much more readily upon subsequent exposure to the original antigen . This occurs because memory T cells require lower activation signals/thresholds to proliferate, which is due, at least in part, to alterations in intracellular calcium mobilization and calcium-dependent signaling processes [25, 26].
ZnO NP Induce ROS Production in Monocytes and T Cells
ROS Quenchers Rescue Primary Monocytes and T Cells from ZnO NP-Induced Cytotoxicity
ZnO NP Preferentially Associate with Monocytes Compared to Lymphocytes
NP association with immune cell subsets
+ FITC-NP (%/MFI)b
1.79% ± 3.1%/1.15 ± 1.27
84.2% ± 4.1%/9.84 ± 3.23
1.61% ± 2.6%/1.23 ± 2.98
82.5% ± 5.9%/14.1 ± 7.21
2.0% ± 1.9%/2.85 ± 4.39
78.1% ± 3.1%/9.61 ± 5.11
1.7% ± 3.4%/5.80 ± 4.82
98.0% ± 2.5%/131.2 ± 20.11
Effect of ZnO NP Size on Cytotoxicity and ROS Production
The increased nanotoxicity with decreasing NP size may be due in part to, the larger surface area/volume ratio of smaller NPs, which provides them with a greater area to associate with cellular membranes and proteins, as well as greater surface reactivity. In addition, ZnO particles prepared in a nonaqueous medium may have oxygen deficient/zinc rich surface chemistries  that exhibit strong electrostatic interactions with the negatively charged cell membrane [35, 36], with smaller particles predicted to have a greater positive surface charge to volume ratio. Thus, greater initial cell membrane-NP association would be expected for smaller NP, leading to potentially greater intracellular uptake.
All of the various ROS molecules produced in this fashion can trigger redox-cycling cascades in the cell, or on adjacent cell membranes. This can then lead to the depletion of endogenous cellular reserves of antioxidants such that irreparable oxidative damage occurs to cellular biomolecules and eventually results in cell death.
ZnO NP Induce Proinflammatory Cytokine Production in Primary Human Immune Cells
The ability of ZnO NPs to induce proinflammatory cytokine expression in human primary immune cells is consistent with the recognized relationship between oxidative stress and inflammation, which is partially mediated by induction of the NF-κB transcription factor . To date, only limited studies have evaluated the effects of ZnO NPs on cytokine production, and most of these studies have been conducted in nonhematological cell types or in immortalized cell lines, which frequently display alterations in signal transduction pathways, leading to unpredictable changes in protein expression. In one report, ZnO NPs were shown to increase IL-8 and MCP-1 cytokine mRNA expression in human aortic endothelial cells, although no information was provided regarding changes in corresponding protein levels . In two other studies conducted in immortalized rodent lung epithelial cells, alveolar macrophage cell lines and primary alveolar macrophages, ZnO NPs fail to induce TNF-α at concentrations exceeding those used in this study [40, 44]. In addition, no changes in other cytokines and chemokines including IL-6, G-CSF, MIP-2, CXCL10, and CCL2 were detected. The ability of ZnO NPs to induce high levels of TNF-α in our studies may be due to differing responses observed between cell populations studied (i.e., PBMC vs. alveolar macrophages), or reflect the longer NP treatment exposure period (i.e., 38 h vs. 24 h). Although, to the best of our knowledge, no published studies have demonstrated the ability of ZnO NPs to induce TNF-α or IL-6 production in purified primary cell cultures or in toxicological evaluations, it is interesting to note that inhalation of ultrafine ZnO particles in occupational settings can increase the expression of these cytokines, which is symptomatically recognized as metal fume fever in welders (e.g., fatigue, fever, chills, myalgias, cough, and leukocytosis) .
The ability of ZnO NPs to induce IL-12, IFN-γ, and TNF-α at NP concentrations below those causing appreciable cytotoxicity indicates immunomodulatory effects that may function to bias the immune response toward Th1-mediated immunity. It is the cytokine profile that directs the development and differentiation of T helper cells into the two different subsets, called type 1 (Th1) and type 2 (Th2) [45, 46]. Th1 cells are recognized to play an essential role in promoting innate and cell-mediated immunity, while Th2 cells promote antibody-based humoral responses . Relevant to our findings, IL-12 and IFN-γ play critical roles in Th1 development, and help set-up a perpetuating loop whereby more Th1 development is favored, Th2 development is suppressed, and the cytotoxicity activity of both NK cells and T cytotoxic cells against cancerous cells, virally infected cells, or intracellular pathogens is enhanced . Thus, our findings indicate that careful titration of ZnO NP-based therapeutic interventions may be successful in elevating a group of cytokines important for eliciting a Th1-mediated immune response with effective anti-cancer actions. These results, combined with our previous observations demonstrating that immortalized hematopoietic cancer cells are preferentially killed (~33-fold) by ZnO NPs compared to normal cells of identical lineage , suggest that ZnO NPs may function via a two-fold mechanism to eliminate cancer cells by direct and preferential cytotoxic actions, and by enhancing the type of immunity most effective at eliciting an in vivo anti-cancer response.
The ability of ZnO NPs to induce TNF-α may also help to promote Th1 differentiation [45, 47] as well as functioning as a regulator of acute inflammation . Notably, this cytokine received its name based on its potent in vitro and in vivo anti-tumor activity. However, high level and/or chronic exposure to TNF-α has been shown to produce serious detrimental effects on the host, including septic shock or symptoms associated with autoimmune disease . Our results demonstrate significant dose-dependent increases in TNF-α over a somewhat narrow range of ZnO NP concentrations. The magnitude of TNF-α induction, as well as other proinflammatory cytokines, and their local–regional delivery to tumor sites or other desired areas, will undoubtedly be important parameters when considering ZnO NP for biomedical purposes to achieve the desired therapeutic response without eliciting potential systemic damaging effects from these cytokines.
Results from these studies demonstrate that ZnO NPs induce toxicity in a cell-type specific manner that is dependent on the degree of NP-cellular membrane association, phagocytic ability, and inherent cellular capacities for ROS production. Monocytic cells displayed the greatest susceptibility and intracellular ROS production following NP exposure, followed by NK cells, followed by lymphocytes, which displayed the most resistance. Studies employing ROS quenchers implicate ROS formation as a major mechanism of ZnO NP-induced toxicity, and demonstrate that the generation of ROS and the cytotoxic profile occurs in a NP size-dependent manner, with smaller NPs displaying the greatest effect. The variable responses of immune cells described in this study underscore the need for careful in vitro evaluation across a spectrum of relevant cell types, followed by appropriate in vivo studies to provide useful and potentially predictive screening data regarding the relative toxicity effects of metal-oxide NPs. These factors are also important considerations for the potential incorporation of ZnO NPs into novel nanotechnology-based biological applications.
Our results demonstrating that ZnO NPs can induce the expression of immunoregulatory cytokines is also a relevant consideration for potential use in biomedical applications, as many current treatments for human disease function by manipulating and controlling components of the immune response . To the best of our knowledge, our results appear to be the first to document the ability of ZnO NPs to increase the expression of IFN-γ, TNF-α, and IL-12 in primary immune human cells, and at NP concentrations below those causing appreciable cell death. These findings suggest that ZnO NPs, when used at appropriate concentrations, could directly enhance tumor cell killing through the production of TNF-α, and could also facilitate effective anti-cancer actions by eliciting a cytokine profile crucial for directing the development of Th1-mediated immunity. The in vivo therapeutic window for ZnO NP to increase proinflammatory cytokines indicates that parameters controlling ZnO NP toxicity, such as particle size, concentration, and biodistribution will need to be carefully controlled when considering metal-oxide NPs for use in biological applications, especially given the recognized relationship of chronic inflammatory processes and tumorigenesis. Future studies are needed to investigate the effects of ZnO NPs on additional cytokines and inflammatory mediators, as well as mechanisms of NP cellular uptake and ROS formation.
This research was supported in part by the Mountain States Tumor and Medical Research Institute, Boise, ID, NSF grants (DMR-0449639, DMR-0605652, 0840227, MRI 0821233, and MRI 0521315), NIH (1R15 AI06277-01A1), DOD (W911NF-09-1-0051), and DoE-EPSCoR (DE-FG02-04ER46142).