Three-Dimensional Microtissue Assay for High-Throughput

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Three-Dimensional Microtissue Assay for High-Throughput Cytotoxicity of Nanoparticles Yang Luo,†,∥ Chaoming Wang,†,§ Mainul Hossain,†,‡ Yong Qiao,† Liyuan Ma,† Jincui An,† and Ming Su*,†,‡,§ †

NanoScience Technology Center, ‡School of Electrical Engineering and Computer Science, and §Department of Mechanical, Materials and Aerospace Engineering, University of Central Florida, Orlando, Florida 32826, United States ∥ Department of Laboratory Medicine, Southwest Hospital, Third Military Medical University, Chongqing 400038, China S Supporting Information *

ABSTRACT: Traditional in vitro nanotoxicity researches are conducted on cultured two-dimensional (2D) monolayer cells and thereby cannot reflect organism response to nanoparticle toxicities at tissue levels. This paper describes a new, highthroughput approach to test in vitro nanotoxicity in threedimensional (3D) microtissue array, where microtissues are formed by seeding cells in nonsticky microwells, and cells are allowed to aggregate and grow into microtissues with defined size and shape. Nanoparticles attach and diffuse into microtissues gradually, causing radial cytotoxicity among cells, with more cells being killed on the outer layers of the microtissue than inside. Three classical toxicity assays [3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide] (MTT), glucose-6-phosphate dehydrogenase (G6DP), and calcein AM and ethidium homodimer (calcein AM/EthD-1)] have been adopted to verify the feasibility of the proposed approach. Results show that the nanotoxicities derived from this method are significantly lower than that from traditional 2D cultured monolayer cells (p < 0.05). Equipped with a microplate reader or a microscope, the nanotoxicity assay could be completed automatically without transferring the microtissue, ensuring the reliability of toxicity assay. The proposed approach provides a new strategy for high-throughput, simple, and accurate evaluation of nanoparticle toxicities by combining 3D microtissue array with a panel of classical toxicity assays.

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lactate or alanine level, and drug response. Thus cell−cell interactions are substantially decreased in 2D cell culture, resulting in an inaccurate prediction of in vivo toxicity upon nanoparticle exposure. Moreover, a key physiological process of transporting nanoparticles through cell layers is also missing in the traditional 2D cell model. These factors have caused a large discrepancy in nanotoxicity results from 2D monolayer cell cultures and in vivo animal models. For instance, considerable literature has reported that quantum dots,14 fullerenes,15 and carbon nanotubes16 are highly or moderately toxic in 2D assay, while their animal counterpart models have shown much lower side effects.17−19 Avascular three-dimensional (3D) microtissue culture systems have been widely used to evaluate toxicities of drug candidates such as anticancer drugs.20,21 Three-dimensional micro- or millimeter tissues can develop morphological and physiological characteristics similar to those of in vivo grown tissues.22,23 Thereby, 3D tumor microtissue-based toxicity assays fill the knowledge gap between in vitro 2D cell assay and in vivo animal testing.24,25 Three-dimensional tumor microtissue could also acquire clinically relevant multicellular resistance to apoptosis-inducing drugs that mimic chemoresistance of solid tumors.25 A variety of methods have been

anoparticles have shown great potential for biomedical applications including drug delivery, bioimaging, chemotherapy, radiation therapy, and tissue engineering.1−5 Nanotoxicity should be clearly addressed prior to their in vivo applications due to their small sizes, highly reactive surfaces, and intimate contact with cellular components such as cell membranes, organelles, proteins, and DNA.6−9 Many methods have been developed to study nanotoxicity, including in vitro two-dimensional (2D) cell culture and in vivo animal models. Although they provide accurate nanotoxicity information, animal-based in vivo nanotoxicity assays are time-consuming and very expensive and thus are often limited to cases where toxicity has already been studied in vitro and need to be validated further. In vitro nanotoxicity assays have drawn great attention by providing fundamental information on in vitro nanotoxicity and can be done at high efficiency and low cost. With the goal of faster, more controlled, and low-cost assays, various high-throughput in vitro platforms for cytotoxicity assays have been developed such as microplate and lab-on-achip microfluidic techniques.10,11 Although these approaches demonstrate high efficiency in cytotoxicity assays by exhibiting real-time control of fluid flow and multiplexing capabilities, most in vitro nanotoxicity studies are conducted with 2D cultured monolayer cells that grow on polymer or glass surfaces.12,13 In such unnatural environments, many cells develop phenotypes that are different from those grown in real tissues in terms of morphology, metabolic characteristics, © XXXX American Chemical Society

Received: May 4, 2012 Accepted: June 29, 2012

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developed to make 3D microtissues such as surface patterning, hanging drop, micromicrotissue chips, liquid-overlay methods, spinning, microfluidics, and printing.26−29 Each method has its own advantages and limitations. For instance, spinner flask technique can be used for large-scale culturing and maintaining microtissues under optimal conditions over a long time. However, it is necessary to transfer the microtissues to stationary culture systems before running a toxicity assay. Such transfer could change microtissue morphology or damage its structure, making accurate microtissue monitoring impossible. Hanging drop techniques can form relatively uniform microtissue and are cost-effective, but a transfer step is also required before the toxicity assay, which is cumbersome and is counterproductive to large-scale screening; also, nonadhesive dishes result in formation of multiple microtissues with a broad size range, which require further sorting and are not suitable for screening. Size-defined 3D microtissues can be produced by some gravity-enforced assembly methods, but cellular organelles within the microtissue are potentially damaged due to shear stress.30,31 To our knowledge, there is no existing highthroughput nanotoxicity study with 3D microtissue array, mainly due to the lack of technique to make a size-uniform and dense 3D microtissues array. The 96-well microplate-based strategy is a good candidate for high-throughput toxicity assays in a single plate, because it is simple and cheap to make sizeuniform microtissues, and there is no need to transfer microtissues for further nanotoxicity assay.30,31 This study constructs a novel 3D microtissue array approach for high-throughput nanotoxicity assay of several nanoparticles (bismuth nanoparticles, iron oxide, and quantum dots). Bismuth nanoparticles are analyzed because they are widely used as contrast agents in X-ray imaging,32 active components in chemotherapy,33 and radiosensitizers in radiation therapy due to their superior ability to absorb X-rays (Z = 83) and good biocompatibility. Although bismuth-derived medicines are known to have low toxicities when administrated for Helicobacter pylori eradiation and tumor treatment,34 the toxicity of bismuth nanoparticles on mammalian cells remains unknown. Quantum dots and iron oxide nanoparticles are chosen as the most and the least toxic controls because their nanotoxicities are well recognized. HeLa or MG-63 cells are adopted to form the 3D microtissue in this study because they are from different tissue origins and they are easy to grow into tumorlike structures for drug screening. In this method, cell suspensions are seeded into microwells of agarose-coated microplate, where the cells aggregate into a microtissue and the microtissue keeps growing with time. After treatment of microtissues with bismuth nanoparticles for 24 h, three classical toxicity assays [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] (MTT), calcein AM and ethidium homodimer (calcein AM/EthD-1), and glucose-6-phosphate dehydrogenase (G6DP)] are adopted to test nanotoxicity via the proposed 3D platform (Figure 1). The results are compared with those derived from 2D cultured monolayer cells. Results show that 3D microtissue-derived nanotoxicity is significantly lower than 2D cultured monolayer cell-derived nanotoxicity, mainly due to the protective nature of 3D microtissues. This proposed high-throughput method would allow easy and accurate evaluation of nanoparticle toxicity via a panel of classical toxicity assays without the need for physically transferring the microtissues.

Figure 1. Schematic illustration of development process of 3D cell microtissues: (A) Seeded cells; (B) aggregated cells; (C) formed microtissues; (D) nanoparticle-treated microtissues; (E) toxicity assay of microtissues.

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RESULTS AND DISCUSSION Three-dimensional microtissues of HeLa or MG-63 cells have been produced in 96-well microplates coated by agarose gel at a seeding density of ∼2 × 105 cells/mL, which will grow into a microtissue of ∼2 mm in diameter within several days.35 Figure 2A−C shows optical images of a HeLa microtissue collected

Figure 2. HeLa microtissue growth with time: (A−C) optical images of one HeLa microtissue after culture for (A) 15 min, (B) 1day, and (C) 4 days; (D) size of HeLa cell microtissue as a function of time; and (E, F) typical fluorescent images of HeLa microtissue in a calcein AM/EthD-1 assay after (E) 4 days and (F) 6 days of culture.

after HeLa cells were seeded for certain time, where the suspended cells start to aggregate in 15 min (Figure 2A). As time goes on, the aggregate becomes denser than the suspended cells and an initial aggregated cell microtissue forms on the first day (Figure 2B). Then the size of microtissue keeps increasing, and the diameter of microtissue becomes larger. Figure 2C shows an image with clear edge observed after culturing for 4 days. The size of microtissue increases to 2.2 mm by day 7 and remains stable until day 12. Figure 2D shows the size of microtissue increases as a function of time, where 4− 5 days of culture can produce uniform HeLa microtissues with an average size of ∼2.0 mm, at the initial seeding concentration of 2 × 105 cells/mL. It is observed that the microtissue grows in a typical exponential solid tumor growth pattern during the first several days, followed by a plateau, with little or no growth in the coming days. This is consistent with the results reported previously.36 Considering that cellular growth status has a significant influence on the following cytotoxicity assays, we evaluate the viability of microtissue with calcein AM/EthD-1 dual-fluorescent assay. In a typical calcein AM/EthD-1 assay, the live cells are stained green by calcein AM and the dead cells are stained red by EthD-1. Figure 2E shows that 99% cells are alive at day 4, and Figure 2F shows that a longer culturing time B

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Figure 3. Calcein AM/EthD-1 results from 3D microtissues and 2D monolayer cells after exposure to surface-modified bismuth nanoparticles. (A1− D1) Schematic illustrations of (A1) bare, (B1) amine-group-terminated, (C1) PEG-terminated, and (D1) silica-encapsulated bismuth nanoparticles. (A2−D2) Representative fluorescent images of HeLa microtissues in calcein AM/EthD-1 assay after incubation with 200 μg/mL (A2) Bi, (B2) BiNH2, (C2) Bi-PEG, and (D2) Bi@SiO2 for 24 h. (A3−D3) Cell viabilities of 3D HeLa microtissues (black bars) and 2D monolayer cells (white bars) after being treated by (A3) Bi, (B3) Bi-NH2, (C3) Bi-PEG, and (D3) Bi@SiO2 for 24 h. (A4−D4) Cell viabilities of 3D MG-63 microtissues (black bars) and 2D monolayer cells (white bars) after being treated with (A4) Bi, (B4) Bi-NH2, (C4) Bi-PEG, and (D4) Bi@SiO2 for 24 h. All surface modifications were conducted on bare bismuth nanoparticles with 20 nm diameter. Data are shown as mean ± SEM (n = 5).

shows the characteristic Lα1 and Lβ1 peaks of bismuth at 10.82 and 13.02 keV, respectively (see Supporting Information). The surface of bismuth nanoparticles is slightly oxidized by heating at 120 °C to be modified with silane derivatives. Bare bismuth nanoparticles (Bi, Figure 3A1) are modified to have an amine functional group (Bi-NH2, Figure 3B1), PEG (Bi-PEG, Figure 3C1), and silica shell (Bi@SiO2, Figure 3D1), respectively. The microtissues are exposed to these nanoparticles at final concentrations of 2, 20, and 200 μg/mL, respectively. After staining of microtissues with calcein AM/EthD-1 dualfluorescence stain, live (green) and dead (red) cells can be clearly distinguished in the fluorescence images of microtissues. Figure 3A2−D2 show that Bi- and Bi-NH2-treated HeLa microtissues have more dead cells than Bi-PEG- or Bi@SiO2treated HeLa microtissues, indicating that Bi and Bi-NH2 are more toxic than Bi-PEG and Bi@SiO2 on microtissues. Figure 3A3−D3 shows that bismuth viabilities derived from both 3D and 2D cells decrease as bismuth nanoparticle concentration increases from 2 to 200 μg/mL. Low concentration of all the bismuth nanoparticles (2 μg/mL) shows no significant toxicities when compared with untreated control (p > 0.05), whereas 200 μg/mL Bi, Bi-NH2, Bi-PEG, and Bi@SiO2 kill 29.6%, 31.8%, 14%, and 17.9% of the HeLa cells in 3D microtissues, respectively (Figure 3A3−D3). Meanwhile, these bismuth nanoparticles killed 39.4%, 42.5%, 36.5%, and 47.9% of the 2D HeLa cells, respectively. These results show the 3D-derived bismuth nanotoxicity is in the order Bi-NH2 > Bi > Bi@SiO2 > Bi-PEG. In addition, the nanotoxicity derived from 3D microtissue is significantly lower than from 2D monolayer cells (p < 0.05). Bismuth nano-

(>6 days) usually leads to a significant increase in dead cell number located in the center of microtissues, due to shortage of oxygen and nutrition.20,36 To avoid the interference of dead cells with the background, all the toxicity tests are performed on day 4. The size of microtissue can be tuned by changing the seeding density of cells. HeLa microtissues with diameters of 1.5 ± 0.11, 2.1 ± 0.16, and 2.5 ± 0.18 mm, have been made at seeding densities of 5 × 104, 1 × 105, and 2 × 105 cells/mL, respectively. The growth rates of cells in 3D microtissue are studied by measuring the total DNA content of the cells in the microtissue, which is compared to those from 2D cultured monolayer cells. Results show that for 2 × 105 cells/mL seeding density, the division rates of 2D cells are higher than those of the 3D microtissues. The growth of 3D microtissues is more like in vivo cell growth within human tissues, and this phenomenon can be explained by the shortage of nutrient supply and down-regulation of various growth factors as in solid tumors.37 To make the comparison of toxicity assay between 2D and 3D cultured cells more accurate, the same cell numbers are subjected to the nanoparticle treatment. Because the 2D monolayer cells grow faster than 3D microtissue, the 2D monolayer cells are seeded at a lower concentration (5 × 104 cells/mL) than 3D microtissue culture (2 × 105 cells/mL), ensuring they are able to grow into similar cell numbers after 4−5 days of culture.36 Bismuth nanoparticles are made by thermal decomposition of bismuth acetate. Transmission electron microscopy (TEM) image shows that nanoparticles have an average diameter of ∼20 nm (Figure 6A). X-ray fluorescence (XRF) spectrum C

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toxicities were also tested with MG-63 microtissues. As expected, Figure 3A4−D4 shows that the nanotoxicity on 3D MG-63 microtissues is significantly lower than on 2D monolayer MG-63 cells at the same nanoparticle concentration (p < 0.05). In addition, the nanotoxicities derived from both 2D monolayer MG-63 cells and 3D MG-63 microtissues show the same order as those from HeLa cells (Bi-NH2 > Bi > Bi@SiO2 > Bi-PEG). These results clearly show that both 3D microtissues (HeLa and MG-63) are more resistant to nanotoxicity than 2D monolayer cells, although a surface modification-dependent difference is observed. The reason that 3D microtissues show higher viabilities can be explained by a protective effect of 3D microstructures. Cells at the external surface of microtissues are more susceptible to nanotoxicity than inner cells. This has been verified by the nonuniform distribution of dead cells (stained red) in microtissues (Figure 3C2,D2), where more dead cells (red) are located in the external area of microtissue while most of the inner cells are alive. Several factors may contribute to the protective effect in 3D microtissue: (1) reduced surface area of cells exposed to nanoparticles, (2) nanoparticle concentration profile along radial direction of microtissue, and (3) microenvironments in the microtissue. First, the surface area of cells exposed to the nanoparticles is decreased in microtissue. For a total of N cells each with radius r, the total area of cells in a monolayer is 2Nπr2, if it is assumed that monolayer cells are 100% confluent. If these cells aggregate to a spheroid with a radius of R, the surface area will be 4πR2. Assuming the shape and size of cells do not change upon aggregation and cells in microtissue can be taken as uniform spheres, then R = 3 N r. The ratio of microtissue surface area exposed to solution and total area of cells is 2/3 N . For a total of 1000 cells, the diameter of microtissue is 10 times larger than that of a cell; and the surface area of microtissue is 20% that of a cell monolayer. When nanoparticles are introduced in the solution containing microtissue, the nanoparticles diffuse rapidly in solution and diffuse into microtissue through the gap between cells. Because nanoparticles attach on the cell surface at certain rate, the number of nanoparticles across the radial direction of microtissue is gradually reduced. Finally, cells in the outer layer of the microtissue attach more nanoparticles than those in the center, and those in the center are less damaged due to low concentration of nanoparticles, whereas in 2D culture, each cell is evenly exposed to the same amount of nanoparticles, and thus damage to each cell is the same. Our experiment is identical to the previous reports that nanotoxicities of CdTe and CTAB-Au nanoparticles on the microtissue are substantially lower than on the 2D culture.24 Second, the diffusion process of nanoparticles penetrating into the microtissues is studied to help explain the protective effect. We estimated the diffusion efficiency of surface-modified bismuth nanoparticles by continuously recording the decrease in nanoparticle concentration in the solution. Figure 4A shows that the nanoparticle concentration decreases when incubation time increases, indicating the nanoparticles diffused into the microtissue in an exponential manner. The nanoparticles inside the microtissue become saturated ∼1 h later and no more nanoparticles can diffuse in. It is obvious that the diffusing velocity and the amount of nanoparticles that diffuse in the microtissue are dependent on surface modification. Bi-NH2 nanoparticle in the solution decreases fastest among the nanoparticles (Bi, Bi-NH2, Bi@SiO2, and Bi-PEG) in a

Figure 4. (A) Diffusion of bismuth nanoparticles into the microtissues. Red, black, green, and blue lines are exponential fits of the amount of Bi, Bi-NH2, Bi@SiO2, and Bi-PEG nanoparticles in the solution, respectively. Data are shown as mean ± SEM (n = 5). (B−D) Confocal images of microtissues treated for 1 h with 200 μg/mL 560 nm (B) CdSe/ZnS-COOH, (C) CdSe/ZnS-NH2, and (D) CdSe/ ZnS-PEG.

given time, indicating that the microtissue absorbs Bi-NH2 nanoparticles with the highest rate. On the contrary, the microtissue absorbs Bi-PEG at the lowest rate with only ∼15% nanoparticles decrease finally. The differences in absorption mainly depend on the interaction between nanoparticles and microtissues, either dominated by adhesion and phagocytosis or by diffusion. To differentiate individual mechanisms of nanoparticles reacting with microtissue, quantum dots are adopted to test the diffusion process. Figure 4B shows that CdSe/ZnS-COOH distributes relatively even within microtissue. However, a strong red fluorescent ring can be observed for CdSe/ZnS-NH2 (Figure 4C) and only a thin red fluorescent circle at the outer layer of microtissue can be seen for CdSe/ ZnS-PEG (Figure 4D). These results indicate cationic Bi-NH2 nanoparticle is more easily entrapped by outer layer cells (with negative surface charge); whereas the weak fluorescent ring on the outer layer of Bi-PEG-treated microtissue is mainly due to less cell-reactive properties of PEG. Third, microtissue microenvironment is another important factor affecting the toxicity results. The phenotype and function of individual cells are highly dependent on sophisticated interactions with 3D organized extracellular matrix (ECM) proteins and neighboring cells. ECM proteins may play important roles in the toxicity tolerance in 3D microtissue culture because they form a natural barrier that limits the diffusion of the nanoparticles. It has been reported that the surface of HepG2 microtissue is covered by a well-developed ECM layer, which is common for all tissues and can reduce the penetration of methotrexate.24 Moreover, the cellular functions are well maintained in microtissue cultures, and it is possible that cells in 3D microtissue will have enhanced repair ability or damage tolerance compared to those in 2D culture. Similar phenomena have been observed in hepatocyte microtissue.38 To verify the feasibility of this proposed nanotoxicity assay platform, two classical toxicity assays (MTT and G6DP) are applied, together with a dual-fluorescence live/dead assay. These cytotoxicity assays are most popular in nanotoxicity research because cell membranes are susceptible to nanotoxicity. Among them, MTT tests detect integrity of the cell D

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more easily encapsulated by lysosomes upon entering cells through endocytosis than Bi. Lysosome encapsulation reduces the contact between bismuth nanoparticles and mitochondria, decreasing the damage to mitochondria.8,39 In the case of BiPEG, it does not easily attach on the cell due to nonsticky property of PEG, thus reducing damage to mitochondria.40−43 To compare nanotoxicity of bismuth nanoparticles (Figure 6A) on 3D microtissues, nanotoxicities of nanoparticles (CdSe/

membrane and the activity of enzyme in mitochondria, reflecting the extent of damage on mitochondria function; the G6PD assay detects levels of G6PD leaking from damaged membranes; and EthD-1 stain detects cell membrane integrity while calcein AM detects intracellular esterase activities. MTT assay results (Figure 5A1−D1) show that viabilities of

Figure 5. Nanotoxicity results from MTT and G6PD assays. (A1− D1)MTT results for 3D microtissues after treatment with (A1) Bi, (B1) Bi-NH2, (C1) Bi-PEG, and (D1) Bi@SiO2. (A2−D2) G6PD results for 3D microtissues after treatment with (A2) Bi, (B2) Bi-NH2, (C2) Bi-PEG, and (D2) Bi@SiO2. Black bars represents HeLa microtissue, and white basr represents MG-63 microtissue. (*) Statistically significant difference from untreated microtissues (p < 0.05). Data are shown as mean ± SEM (n = 5).

Figure 6. Comparison of toxicity results from three nanoparticles. (A− C) TEM images of (A) bismuth nanoparticles, (B) iron oxide nanoparticles, and (C) CdSe/ZnS quantum dots. (D, E) Nanotoxicities of different nanoparticles from calcein AM/EthD-1 assay on (D) HeLa microtissues and (E) MG-63 microtissues. (*) Statistically significant difference when compared with untreated controls (p < 0.05); (#) significant difference when compared with CdSe/ZnSCOOH (p < 0.05). (**) significant difference when compared with Fe3O4-COOH (p < 0.05). Data are shown as mean ± SEM (n = 5).

microtissues decrease as nanoparticle concentrations increase, showing concentration-dependent nanotoxicity (both HeLa and MG-63). At low nanoparticle concentration (2 μg/mL), no significant killing is observed on HeLa and MG-63 microtissues when compared to untreated control (p > 0.05), indicating nontoxicity or extremely low nanotoxicity at low concentration. The viabilities of microtissues are significantly decreased when high concentrations of nanoparticles (200 μg/mL) are used, in comparison with untreated control (p < 0.05). At high nanoparticle concentration (200 μg/mL), Bi, Bi-NH2, BiPEG, and Bi@SiO2 nanoparticles kill 27%, 34%, 13%, and 16% of HeLa cells in microtissue, respectively. The G6PD assay results show that these nanoparticles kill 29%, 34%, 19% and 22% HeLa cells in 3D microtissue, respectively (Figure 5A2− D2). These results show that cytotoxicities of these surfacemodified nanoparticles are in the order Bi-NH2 > Bi > Bi@SiO2 > Bi-PEG, which is identical to the order of results from calcein AM/EthD-1 dual-fluorescent assay, indicating the consistency among the toxicity assays. Although the toxicities from G6PD assay for Bi-NH2 and bismuth nanoparticles are similar to those from MTT assay, the toxicities from G6PD assay for Bi-PEG and Bi@SiO2 are 6% higher than those from MTT assay. This is probably due to different principles of these toxicity assays: G6PD leaking assay detects membrane integrity, whereas the MTT result indicates metabolic activity. Thus the discrepancy between results from G6PD and MTT could be explained by different pathways affecting cells. In our experiments, toxicities from G6PD assay are identical to those from MTT for Bi-NH2 and bismuth nanoparticles, indicating these nanoparticles can affect cellular metabolic activity as well as plasma membrane integrity. However, toxicities from G6PD assay are higher than those from MTT for Bi-PEG and Bi@SiO2, indicating that cell metabolic activity is less affected compared to plasma membrane. Bi@SiO2 has less impact on the metabolic activity, probably because the size of Bi@SiO2 (50 nm) is larger than that of bismuth nanoparticle (20 nm), and thus Bi@SiO2 is

ZnS and Fe3O4) with different surface modifications (NH2 or COOH groups) are tested (Figure 6B,C). For different nanoparticles modified with COOH, calcein AM/EthD-1 results show the toxicity of CdSe/ZnS is significantly higher than that of Bi@SiO2 and Fe3O4 (p < 0.05). Figure 6D shows that CdSe/ZnS-COOH kills 32.3% of HeLa cells in microtissues, followed by Bi@SiO2-COOH (kills 10.3%), and Fe3O4COOH (kills 6.4%), at 200 μg/mL nanoparticle concentration. For nanoparticles of the same core, Bi@SiO2-NH2 is 4.3% more toxic than bare Bi@SiO2-COOH, and Fe3O4-NH2 is 3.1% more toxic than Fe3O4-COOH (p < 0.05), suggesting NH2 may give higher toxicity due to the mediation of endocytosis. Meanwhile, Figure 6E demonstrates that MG-63 microtissue responds similarly to HeLa microtissue to nanotoxicity, showing that CdSe/ZnS is the most toxic species in three types of nanoparticles (p < 0.05), which is consistent with the results from previous reports.44−47 In addition, Fe3O4-NH2 nanoparticles kill more HeLa microtissues than Fe3O4-COOH nanoparticles, while Fe3O4-NH2 nanoparticles kill less MG-63 microtissues than Fe3O4-COOH nanoparticles at the same concentration. These results indicate the toxicity of different surface-modified nanoparticles may also depend on the cell types that comprise the microtissues. Future research will focus on elucidating the mechanisms of protective effects in 3D microtissue and identifying the differences derived from different cell types.

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CONCLUSION A 3D microtissue array has been made and used to study nanotoxicity of bismuth nanoparticles. Nanotoxicities from 3D microtissue are lower than those from corresponding 2D E

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was collected at 40 kV and 100 μA after 2 min exposure for bismuth nanoparticles. Surface Modifications. To conjugate with poly(ethylene glycol) (PEG), 1 mg of bismuth nanoparticle and 20 μL of PEG silane were added in 180 μL of toluene. After reaction for 3 h, the modified nanoparticles were collected by centrifugation and unreacted PEG silane was removed by washing with toluene. Amine-terminated bismuth or iron oxide (Fe3O4) nanoparticles were obtained by incubating them with 5% APTES in toluene for 2 h, followed by centrifugation and washing with toluene to remove extra silane. Carboxylterminated CdSe/ZnS quantum dots (CdSe/ZnS-COOH) were made by mixing 35 μL of MAA with 1 mg of CdSe/ ZnS in 500 μL of chloroform under sonication for 2 h. The unreacted MAA was removed by dialysis against deionized (DI) water with a 3.5 kDa cutoff membrane for 12 h. The stock solutions of bare bismuth nanoparticle (Bi) or silicaencapsulated bismuth (Bi@SiO2); amine-terminated bismuth (Bi-NH2), Fe3O4 (Fe3O4-NH2), or silica-coated bismuth (Bi@ SiO2-NH2); carboxyl-terminated Fe3O4 (Fe3O4-COOH), silicaencapsulated bismuth (Bi@SiO2−COOH), or CdSe/ZnS (CdSe/ZnS-COOH); and PEG silane-modified bismuth (BiPEG) nanoparticles were dispersed in sterilized phosphatebuffered saline (pH = 7.4) and ready to use. Three-Dimensional Cell Culture and Nanoparticle Treatment. HeLa and MG-63 cell lines were used to study nanoparticle cytotoxicity in 3D microtissue and 2D monolayer cells. The 3D microtissue was cultured as follows: 50 μL of 2% agarose gel was introduced into each well of a flat-bottom 96well microplate. The microplate was gently shaken to spread gel over the bottom of well evenly, and any bubbles were pipetted out. After 10 min, a solid nonadhesive agarose gel was formed in each well of the microplate. All HeLa and MG-63 cells were grown in RPMI 1640 medium supplemented with penicillin (100 units/mL), streptomycin (100 μg/mL), and 10% fetal bovine serum (FBS). HeLa and MG-63 cells were cultured according to the protocol from ATCC. Briefly, 200 μL of cell suspension was seeded into each well with a final concentration of 2 × 105 cells/mL, and then the plate was cultured in an incubator with 5% CO2 at 37 °C for some time to allow 3D cell microtissue to form. The cells tend to aggregate and form a round microtissue on a nonadhesive gel surface. Total DNA content of the microtissue was quantified by PicoGreen dsDNA assay. Once microtissue reached a diameter of ∼2 mm, the medium was exchanged and nanoparticle suspension was added (Bi, Bi-NH2, Bi@SiO2, or Bi-PEG) in growth medium at final concentrations of 2, 20, and 200 μg/mL. After 24 h, the medium was removed and all wells were washed with PBS prior to any toxicity assays. For diffusion assay, the microtissues are treated with 200 μg/ mL bismuth nanoparticles (Bi, Bi-NH2, Bi@SiO2, or Bi-PEG) or quantum dots (CdSe/ZnS-COOH, CdSe/ZnS-NH2, or CdSe/ZnS-PEG). The solution was pipetted out at 10 min intervals for nanoparticle concentration measurement (XRF or fluorescence). For 2D cell culture, the same cell suspensions were prepared and 200 μL of HeLa or MG-63 cell suspension were seeded with final concentrations of 1 × 105 cells/mL in each well and cultured in an incubator with 5% CO2 at 37 °C. After the monolayer of cells become 80% confluent, the cells were incubated with nanoparticles (Bi, Bi-NH2, Bi@SiO2, BiPEG, Bi@SiO2-NH2, CdSe/ZnS-COOH, Fe3O4-COOH, or Fe3O4-NH2) in fresh medium at final concentrations of 2, 20, and 200 μg/mL. After 24 h, the medium in each well was

monolayer cells due to the protective effect of surrounding cells on the internal cells. Because 3D microtissues are closer to in vivo natural tissues morphologically and physiologically, the nanotoxicity derived from 3D microtissue can be more accurate than that from 2D monolayer cells. Most of the traditional toxicity assay methods derived from 2D monolayer cell culture can be applied in this proposed 3D high-throughput nanotoxicity approach. Based on 3D microtissues, the nanotoxicity of bismuth nanoparticle is found to be similar to that of iron oxide nanoparticles and is lower than that of CdSe/ZnS quantum dots. The nanotoxicity assay based on 3D microtissue can increase precision of therapeutic readouts, can enable drug screening at tissue level with low cost, and can be an alternative to existing 2D monolayer cell assays. The high-throughput 96microwell platform will allow highly efficient nanotoxicity assays of future nanomedicines.

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EXPERIMENTAL SECTION Materials. The kits for Vybrant MTT cell proliferation, Vybrant cytotoxicity, live/dead assays, and Quant-iT PicoGreen dsDNA assay were from Invitrogen (Carlsbad, CA). Poly(ethylene glycol)- (PEG-) terminated silane (PEG-silane, 472− 604 g/mol) was from Gelest (Tullytown, PA). Bi (CH3COO)3, BiCl3, NaBH4, RPMI 1640 culture medium, penicillin, streptomycin, fetal bovine serum (FBS), and Dulbecco’s phosphate-buffered saline (D-PBS) were from Sigma−Aldrich (St. Louis, MO). Anhydrous dimethyl sulfoxide (DMSO), toluene, poly(vinylpyrrolidone) (PVP), 3-aminopropyltriethoxysilane (APTES), iron oleate, tetraethyl orthosilicate (TEOS), octadecene, diphenyl ether, sodium oleate, and methacrylic acid (MAA) were from VWR (West Chester, PA). Ultrapure water (18.2 MΩ·cm−1) from Nanopure System (Barnstead, Kirkland, WA) was used. Synergy HT multimode microplate reader from Biotek (Winooski, VT) was used for absorbance and fluorescence measurements. HeLa (CLL-2) and MG-63 (CRL-1427) cell lines were from American Type Culture Collection (ATCC, Manassas, VA). Inverted optical microscopy from Milesco scientific (Accu-Scope 3032, Princeton, MN) was used to observe cultured cells. Fluorescence images of microtissue were taken on a Zeiss Axioskop 2 mot plus confocal microscope and an inverted fluorescence microscope from Olympus (GX51). All chemicals used in this study were analytical-grade and were used without further purification. Nanoparticle Synthesis and Characterization. Bismuth, core−shell structural zinc sulfate-coated cadmium selenide (CdSe/ZnS) and iron oxide (Fe3O4) nanoparticles were made by chemical reduction or thermally decomposition methods. Silica-coated bismuth nanoparticles were made as follows: bismuth nanoparticles were ultrasonically redispersed in a mixture that contains 240 mL of ethanol and 60 mL of water, followed by adding 4 mL of TEOS under vigorous stirring. After reaction for 1 h, the nanoparticles were separated by centrifugation and washed with deionized water and anhydrous ethanol prior to vacuum drying at 50 °C overnight. The morphology of bismuth nanoparticles was imaged by a JEOL 1011 transmission electron microscope (TEM) operated at 100 kV. An X-ray spectrometer (Amptek X-123) with Si-PIN photodiode was used to analyze X-ray fluorescence (XRF) emissions of the nanoparticles in transmission mode. A 25 μm thick silver and 250 μm thick aluminum filter were used to reduce background and improve signal-to-noise ratio in the low-energy (0−15 keV) region of the spectrum. XRF spectrum F

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ACKNOWLEDGMENTS This project has been supported by a research grant (0828466), a CAREER award from National Science Foundation and a Concept Award (W81XWH-10-1-0961) from Lung Cancer Research Program of Department of Defense, a grant from the National Natural Science Foundation of China (30900348), and a fund for the Transformation of Scientific and Technological Achievements of the Third Military Medical University (2010XZH08).

removed and all wells were washed with PBS prior to any toxicity assays. Cytotoxicity Assays. G6PD (glucose-6-phosphate dehydrogenase), calcein AM/EthD-1 (calcein AM and ethidium homodimer), and MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] assays were done by following the standard protocol from suppliers. For MTT assay, the medium in each well was removed and replaced with 100 μL of fresh medium. Ten microliters of 12 mM MTT stock solution was then added into each well and into a negative control (100 μL of medium without nanoparticles). After incubation at 37 °C for 4 h, 100 μL of SDS−HCl solution was added in each well and mixed thoroughly via pipet. After incubation at 37 °C for 6 h, each sample was mixed well with a pipet and the optical absorbance at 570 nm was recorded. For G6PD assay, 50 μL of 2× resazurin/reaction mixture was added to each well, as well as live and fully lysed control cells. The cells were lysed by adding 1 μL of 100× cell lysis buffer into the well. All samples were assayed in six duplicates. The microplate was incubated at 37 °C for 30 min prior to measurement of fluorescence intensity at 580 nm with 530 nm excitation. For calcein AM/ EthD-1 assay, 100 μL of D-PBS was added in each well to wash cells to dilute serum-containing esterase. Then 100 μL of dualfluorescence calcein AM/EthD-1 assay reagent was added into each well, and the plate was incubated for 30 min at room temperature before fluorescence intensity was measured. A cellfree control was used to measure background fluorescence, which was subtracted before calculation. The percentages of live and dead cells were calculated by the equation provide by the supplier. CdSe/ZnS was adopted to study the diffusion of nanoparticles into the microtissues. A final concentration of 50 μM CdSe/ZnS-COOH was introduced into each microwell containing a microtissue in 200 μL of D-PBS buffer. Then 2 μL of the suspension was taken out at 5-min intervals for fluorescence intensity measurements. Statistical Analysis. Five independent duplicates were performed for each test, and the means were obtained for statistical analysis. Error bars represent the standard error of the means. Statistical analyses were performed with SPSS 16.0 (SPSS Inc., Chicago, IL). One-way analysis of variance (ANOVA) and least significant difference (LSD) tests were applied to compare the viability of microtissues after different nanoparticle treatment. Comparisons between two groups were based on t-test for two independent samples. P < 0.05 was considered statistically significant.

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ABBREVIATIONS 2D two-dimensional 3D three-dimensional Bi bismuth nanoparticle Bi@SiO2 silica encapsulated bismuth Bi@SiO2-NH2 amine-terminated silica-coated bismuth Bi-NH2 amine-terminated bismuth Bi-PEG PEG silane-modified bismuth Fe3O4 iron oxide Fe3O4-NH2 amine-terminated iron oxide Fe3O4-COOH carboxyl-terminated iron oxide CdSe/ZnS-COOH carboxyl-terminated cadmium selenide/ zinc sulfide PEG poly(ethylene glycol) XRF X-ray fluorescence

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ASSOCIATED CONTENT

S Supporting Information *

One figure showing the XRF spectrum obtained at 40 kV and 100 μA after 10 min exposure for bismuth thin film (1 mm2 and 10 nm thick) using a 400 μm collimator, with the net count for bismuth L peak at 10.86 keV as functions of collimator size and exposure times. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail to:[email protected]. Notes

The authors declare no competing financial interest. G

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