Cytotoxicity of quantum dots and graphene oxide to erythroid cells and macrophages
© Qu et al.; licensee Springer. 2013
Received: 16 March 2013
Accepted: 16 April 2013
Published: 30 April 2013
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© Qu et al.; licensee Springer. 2013
Received: 16 March 2013
Accepted: 16 April 2013
Published: 30 April 2013
Great concerns have been raised about the exposure and possible adverse influence of nanomaterials due to their wide applications in a variety of fields, such as biomedicine and daily lives. The blood circulation system and blood cells form an important barrier against invaders, including nanomaterials. However, studies of the biological effects of nanomaterials on blood cells have been limited and without clear conclusions thus far. In the current study, the biological influence of quantum dots (QDs) with various surface coating on erythroid cells and graphene oxide (GO) on macrophages was closely investigated. We found that QDs posed great damage to macrophages through intracellular accumulation of QDs coupled with reactive oxygen species generation, particularly for QDs coated with PEG-NH2. QD modified with polyethylene glycol-conjugated amine particles exerted robust inhibition on cell proliferation of J744A.1 macrophages, irrespective of apoptosis. Additionally, to the best of our knowledge, our study is the first to have demonstrated that GO could provoke apoptosis of erythroid cells through oxidative stress in E14.5 fetal liver erythroid cells and in vivo administration of GO-diminished erythroid population in spleen, associated with disordered erythropoiesis in mice.
Of the popular nanomaterials, quantum dots (QDs) and graphene have promising applications in various fields; however, the cytotoxicty of these nanomaterials is also largely concerned [1, 2]. To date, a few studies have revealed that QDs and graphene posed harm to a spectrum of organisms and cells [3–6]. Blood cells are a large group of cells that play critical roles in many physiological and pathological processes. Of the blood cells, erythrocytes are responsible for carrying oxygen, carbon dioxide, and other wastes; whereas, macrophages are part of the immune system responsible for inflammation and the clearance of pathogens . Erythropoiesis is a highly dynamic process that produces numerous new red blood cells (RBCs), which requires a large amount of iron [8, 9]. Senescent erythrocytes undergo phagocytosis by macrophages, and iron is released into the circulation for erythropoiesis upon erythropoietic demand . Thus, erythrocytes and macrophages are essentially involved in governing the balance of erythropoiesis and iron recycling in the body. Thus far, limited work has been performed in blood cells in evaluating the biosafety of QDs and graphene.
Previous studies have documented that QDs could transport through the plasma membrane of RBCs, exerting potential impairment on the survival or function of RBCs . Our own studies have demonstrated that QDs engulfed by macrophages in spleen could cause impairment to macrophages, which triggered the accumulation of aged RBCs in spleen with splenomegaly . A few other studies have also suggested that graphene or graphene oxide (GO) might impose toxicity to RBCs through hemolysis and incur cell death and cytoskeleton destruction to macrophages [13–16]. To date, the cytotoxicity and related mechanisms of QDs and graphene still remain inconclusive for blood cells due to limited data. To this end, in the current study, we embarked on the cytotoxicity of QDs with different surface modifications to macrophages and GO to erythroid cells. Overall, we demonstrated significant adverse effects of QDs on macrophages and GO on erythrocytes.
QDs with the same core Cd/Te coated with Sn/S and the same diameter (approximately 4 nm) modified with polyethylene glycol (PEG) (QD-PEG), PEG-conjugated amine (QD-PEG-NH2), or PEG-conjugated carboxyl groups (QD-PEG-COOH) were purchased from Wuhan Jiayuan Quantum Dots Co., Ltd. (Wuhan, China) [12, 17]. The evaluation of the fluorescence spectrum indicated that the maximum emission wavelength for QDs used here was around 605 nm, indicative of red light. GO was synthesized using the Hummers method with minor revisions as previously described . The size of GO was 300 to 1,000 nm, and the thickness was approximately 1 nm . GO suspension was stable for at least 1 month. GO suspension was diluted in phosphate buffered saline (PBS) for the following experiments.
Regarding the GO administration in vivo, 6-week-old BALB/C male mice were intraperitoneally injected with 200 μl GO suspension at a concentration of 1 mg/ml (10 mg/kg body weight) every 3 days for 3 weeks. Control mice received PBS only. Twenty four h after the final administration, blood was collected via the heart, and complete blood count (CBC) analysis was carried out using a whole blood analyzer at Peking University Health Center. After the mice were sacrificed, organs were collected.
After perfusion with saline, livers were perfused with 0.05% collagenase and then minced and resuspended in 0.05 g/ml collagenase type IV (Sigma-Aldrich, St. Louis, MO, USA) in Hank's balanced salt buffer . The samples were then incubated in the solution without either cadmium or magnesium for enzymatic digestion at 37°C for 30 min. The digested samples were passed through 70 μm filters. The cells were resuspended in PBS and then incubated with fluorescein isothiocyanate (FITC)-conjugated anti-F4/80 mAb (eBioscience Inc., San Diego, CA, USA) for the selection of macrophage population. Phycoerythrin (PE)-conjugated anti-Ter119 mAb (BD Pharmingen, Franklin Lakes, NJ, USA) was applied to cell suspension for erythroid cell selection. After washing, the cells were analyzed on a fluorescence-activated cell sorting (FACS) Calibur™ (BD Biosciences, San Jose, CA, USA). Splenocytes were similarly prepared from the spleen for FACS analysis.
Mouse J774A.1 (purchased from the Shanghai Cell Bank of Type Culture Collection of the Chinese Academy of Sciences, Shanghai, China) were cultured in DMEM (Hyclone, Thermo Fisher Scientific, Waltham, MA, USA), supplemented with 10% fetal bovine serum (Gibco, Carlsbad, CA, USA) and 100 U/ml penicillin/streptomycin (Gibco). E14.5 fetal liver cells were isolated and cultured as described .
Regarding the assessment of intracellular cadmium mass, J774A.1 cells cultured in 10-cm plates were exposed to QDs for 24 h. Thereafter, the cells were collected and washed with PBS for three times, and cells were digested with HNO3 and H2O2 (3:2, v/v) by microwave-assisted extraction. After the removal of acid, the digested samples were diluted to 5 ml, and Cd mass was assessed using inductively coupled plasma mass spectrometry (ICP-MS) (Agilent 7500, Santa Clara, CA, USA) according to the protocol as previously described . A series of cadmium standard solutions (10, 5, 2, 1, 0.5, 0.2, and 0 ng/g) were prepared to conduct a standard curve for the calibration of Cd concentration.
Cell proliferation was evaluated by the BrdU incorporation assay (Roche, Penzberg, Germany). Briefly, the cells were seeded in 96-well plates with 5.0 × 104 cells per well in 100 μl. The cells were starved in 1% FBS serum medium overnight. The cells were then treated with 47 μg/ml QDs for 48 h, and cell growth was examined according to the instructions provided by the manufacturer.
After exposure to 47 μg/ml QDs for 24 h, the cells were fixed by formaldehyde, followed by a wash with 1% Triton X-100 in PBS. FITC-conjugated phalloidin (Molecular Probes, Invitrogen Corporation, Grand Island, NY, USA) was used to stain filamentous actin (F-actin), and nuclei were counterstained with 4',6-diamidino-2-phenylindole (DAPI) (blue) (Molecular Probes). Laser scanning confocal microscopy was performed to image cells as previously described .
After preincubation with 10 μM 2'-7'-Dichlorodihydrofluorescein diacetate (DCFH-DA) (Sigma-Aldrich) for 30 min, the J774A.1 cells seeded in 24 well-plate (1.0 × 105 per well) were treated with QDs at 47 μg/ml for 6 h. After treatment, the emission spectra of dichlorodihydrofluorescein (DCF) fluorescence at 525 nM were measured using FACS Calibur™ (BD Biosciences). The E14.5 fetal cells were similarly cultured and preincubated with DCFH-DA. Thereafter, the cells were washed with PBS, and treated with 10, 20, 40, and 80 μg/ml GO for 15 min, 0.5 h, 1 h, and 6 h, respectively, followed by DCF fluorescence determination.
For apoptosis analysis of erythroid cells from spleen, splenic cell suspension was co-stained with PE-conjugated anti-Ter119 Ab, FITC-conjugated Annexin V and 7-amino-actinomycin D (7AAD). The cell death of erythroid cells was determined with the channels of Annexin V fluorescence and 7AAD fluorescence by gating Ter119+ cells. With respect to J774A.1 cells, after exposure to QDs for 24 h, the cells were subject to FITC-conjugated Annexin V and propidium iodide (PI) staining. Apoptotic and necrotic cells were assessed by FACS as described previously . The E14.5 fetal liver cells were treated with 20 μg/ml GO for 18 h, and cell death was then similarly examined.
One-way analysis of variance (ANOVA) was employed to assess the mean difference among the groups compared to control. The difference between the two groups was analyzed with two-tailed Student's t test. All experimental data were shown in mean ± SD. P < 0.05 was considered to be statistically significant.
All animal care and surgical procedures were approved by the Animal Ethics Committee at the Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences.
Our recent study suggested that sodium arsenite induced substantial oxidative stress (ROS synthesis), resulting in apoptosis on erythroid cells . We therefore assessed the intracellular ROS level in fetal liver cells after GO treatment. As shown in Figure 5B, the DCF fluorescent intensity was greatly enhanced in fetal liver cells treated with GO at various concentrations for only 15 min (Figure 5B, P < 0.001). The clear shift of DCF fluorescent peak continued at 0.5, 1, and 6 h (Figure 5B, P < 0.001). These results together suggested that GO-induced apoptosis in erythroid cells was likely dependent on ROS-mediated oxidative stress, similar to the mechanism responsible for arsenic-stimulated apoptosis in erythroid cells . Additionally, GO treatment was here determined to cause cell death in erythroid cells via apopotosis, similar to a study demonstrating that graphene stimulated ROS generation and induced cell death via apoptosis in PC12 cells, a cell line derived from a pheochromocytoma of the rat adrenal medulla .
The blood circulation system is an important barrier against invaders, including nanomaterials under biomedical applications or environmental absorption. The blood cells are primarily responsible for governing their trafficking and systemic translocation. Since RBCs are the most abundant cell population in peripheral blood (4.1 to 5.9 × 106/ml RBCs vs. 4.4 to 11.3 × 106/ml white blood cells in humans), these cells presumably have a much greater probability of exposure to nanomaterials in the circulation after administration, with possible adverse effects such as hemolysis [33–35]. For clearance of nanomaterials from the circulation, the macrophages are responsible for recognizing and ingesting these particles . Therefore, the nanomaterials transporting in the circulation or deposited within macrophages could cause harm to these cells as well as to the immune system. To date, studies on toxicity of QDs and GO to RBCs or macrophages have been limited and without conclusive answers, and this certainly warrants detailed investigation.
Our combined results demonstrated that QDs could be readily engulfed by macrophages and provoked intracellular ROS generation. Particularly, QDs coated with PEG-NH2 had a greater capability for entering the cells and revealed a robust ability to repress the proliferation of J774A.1 cells. This indicated that surface modification could be optimized to ensure the function and the safety of QDs as well. Meanwhile, to the best of our knowledge, the biological impact of graphene on erythroid progenitor cells has not been previously reported. Our study is the first to demonstrate that GO could provoke apoptosis of erythroid cells in vitro and in vivo. These data suggested that GO could likely possess the potential to disrupt the concerted balance of erythropoiesis in mammalians including humans. Thus, the adverse effects of GO on RBCs warranted further detail investigation, especially for humans under biomedical and environmental exposure.
This work was supported by grants from the National Basic Research Program of China (2009CB421605), the National Natural Science Foundation of China (grant numbers: 21077128, 20921063, 21177151, 21207152), and from the program of ‘Hundreds Talents’ from the Chinese Academy of Sciences. We thank the laboratory members for their invaluable assistance with experiments and reagents.
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