Bioconjugate Chem. 2004, 15, 897−900
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Toxicity of Gold Nanoparticles Functionalized with Cationic and Anionic Side Chains Catherine M. Goodman, Catherine D. McCusker, Tuna Yilmaz, and Vincent M. Rotello* Department of Chemistry, University of Massachusetts, Amherst, Massachusetts 01003. Received March 2, 2004
The structure and properties of gold nanoparticles make them useful for a wide array of biological application. Toxicity, however, has been observed at high concentrations using these systems. MTT, hemolysis, and bacterial viability assays were used to explore differential toxicity among the cell types used, using 2 nm core particles. These studies show that cationic particles are moderately toxic, whereas anionic particles are quite nontoxic. Concentration-dependent lysis mediated by initial electrostatic binding was observed in dye release studies using lipid vesicles, providing the probable mechanism for observed toxicity with the cationic MMPCs.
The development of alternate drug delivery agents and drug scaffolds is an ongoing task in medicinal chemistry (1-3). The recent burst of research involving gold nanoparticles as transfection vectors (4-6), DNA-binding agents (7-9), protein inhibitors (10), and spectroscopic markers (11-14) demonstrates the versatility of these systems in biological applications. Gold particles display several features that make them well suited for biomedical applications, including straightforward synthesis (7), stability (15), and the facile ability to incorporate secondary tags such as peptides targeted to specific cell types to afford selectivity (16-18). These nanoparticle systems, however, have not been well evaluated to determine their interactions with cells beyond the designated functions. Recently, we have shown that cationically functionalized mixed monolayer protected gold clusters (MMPCs) are capable of mediating DNA translocation across the cell membrane in mammalian cells at levels much higher than polyethyleneimine (PEI), a widely used transfection vector (4). At higher nanoparticle concentrations, however, transfection efficiency decreases due to cytotoxicity. While many available drugs display toxicity at sufficiently elevated concentrations (19-20), the toxicity of the nanoparticles was observed in this case at concentrations only 2-fold in excess of that found to have maximal transfection activity. To provide insight into the origins of toxicity in MMPC systems, we fabricated cationic nanoparticle 1 and anionic particle 2. (Figure 1). The quaternary ammoniumfunctionalized nanoparticle (MMPC 1) is composed of the same monolayer components as those demonstrated to cause cytotoxicity in the mammalian cell cultures (4). The carboxylate-substituted nanoparticle (MMPC 2) does not bind DNA (7) and was therefore not tested in the previous report. The negative substituents on the surface of MMPC 2 are not anticipated to interact with the cell membrane, given the overall negative charge of the lipid bilayer (24). The structure of the gold core and alkane thiol components, however, is identical to that of MMPC 1 and so should provide information relevant to the toxicity due to the particle structure and charge. We utilized cells Cos-1 cells, red blood cells, and bacterial cultures (Escherichia coli) to probe the effect * To whom correspondence should be addressed. E-mail:
[email protected].
Figure 1. Schematic of the structures of MMPCs 1 and 2. Each monolayer consists of approximately 70 charged thiols and 30 unsubstituted thiols.
of the nanoparticles on cell viability. Further insight into the origin of toxicity was obtained using dye release studies using lipid vesicles. Dye release from these vesicles has been strongly correlated with the ability of amphiphilic polymers to lyse bacterial cells (25), providing an indication of whether membrane disruption is a significant factor in the mechanism of toxicity. Taken together, these studies provide insight into the origins of the toxicity observed, allowing design of new particles with an improved toxicity profile. MATERIALS AND METHODS
Colorimetric (MTT) kit for cell survival and proliferation was purchased from Chemicon International. Dulbecco’s Modified Eagle Medium, L-glutamine, fetal bovine serum, trypsin, Trypan Blue stain (0.4%), and gentamicin
10.1021/bc049951i CCC: $27.50 © 2004 American Chemical Society Published on Web 06/05/2004
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were purchased from Gibco. Tissue culture plates were purchased from Costar. Cos-1 cells purchased from ATCC. Acetic acid, ampicillin sodium salt, and sodium phosphate were purchased from EM. LB broth (Miller) and bacto-agar were purchased from DIFCO. Calcein, rhodamine 6G, 11-mercaptoundecanoic acid, and Triton X-100 were purchased from Sigma-Aldrich. Sodium chloride was purchased from Mallinckrodt. L-R-Stearoyloleoyl-phosphotidylcholine (SOPC) and L-R-stearoyl-oleoyl-phosphotidylserine (SOPS) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL). Ampicillin was dissolved in double distilled H2O to yield stock solutions of 50 mg/mL, and aliquots were stored at -20 °C. All other materials were used as received. Gold nanoparticles were fabricated as previously described (7). Cos-1 Viability Assay. Cos-1 cells were split to approximately 80% confluency in a 96-well culture plate the day preceding the assay. Cells were incubated at 37 °C in 7.5% CO2 for 1, 2.5, 6, or 24 h with either MMPC 1 or 2 at 0.38, 0.75, 1.5, or 3 µM. Positive controls contained no nanoparticles. Following incubation periods, the media and nanoparticles were aspirated from the wells, cells were washed 2× with PBS, and fresh media was added. Following the MTT kit protocol, viability was measured by enzymatic reduction of the yellow tetrazolium MTT to a purple formazan, as measured using an EL808IU microplate reader (Biotek Instruments, Inc.) for absorbance readings (26). Mortality studies using Trypan Blue staining were performed using a modified procedure from the described above. In this case, the fresh media to harvest the cells was replaced by 20 µL of trypsin (0.25%), and 20 µL of Trypan Blue was added 5 min before counting cells on a hemocytometer. Top Agar Assays. This procedure was adapted from a bacteriophage λ assay (27). BL21 (dE3) E. coli cells, transformed with a Pet16b vector encoding for ampicillin resistance and cellular retinoic acid binding protein I (28), were grown for 4-20 h in LB media with 0.1 mg/mL ampicillin. The OD of the cells at 600 nm was then measured and the relative concentration diluted to 0.5 × 10-4 absorbance using LB media or double distilled H2O. A volume of 20 µL of the resultant cell solution was diluted with 0.55 mL of a LB-agar mix (25 g/L LB, 7.5 g/L agar) and 1-20 µL 833 µM MMPC solution and poured onto 3.5 cm LB-agar plates (25 g/L LB, 15 g/L agar, 0.1 mg/mL ampicillin). The plates were incubated overnight at 37 °C, and all resultant colonies were counted. The growth of the treated cells is reported relative to control plates. Hemolysis Assay. This method was modified from a published procedure (29). In the studies, 40 µL of red blood cells from a healthy human donor were diluted with 10 mL of 10 mM Tris 150 mM NaCl buffer (pH 7) filtered with a 0.2 µm Acrodisc syringe filter (Pall Corporation, Ann Arbor, MI). The cell solution was centrifuged at 3000 rpm for 5 min, and the supernatant was poured off. Cells were washed until supernatant was clear, and the packed cells were resuspended in a final volume of 10 mL of Tris buffer. A volume of 80 µL of the resultant solution was incubated with 20 µL of MMPC solution (stocks ranging from 0.27 to 833 µM) with shaking for 30 min at 37 °C in a 96-well plate. Sample solutions were then diluted as necessary to allow quantification on an EL808IU microplate reader (Biotek Instruments, Inc.) at 405 nm with 5 s shaking prior to reading the sample. Nanoparticle absorbance was subtracted out according to the absorbance of the nanoparticles in buffer, and the resultant intensity due to detection of heme in solution was used to determine LC50 values.
Goodman et al. Table 1. LC50 Values of MMPC 1 and 2 in Mammalian Cells and E. coli Cos-1 red blood cells E. coli
MMPC 1, µM
MMPC 2, µM
1.0 ( 0.5a 1.1 ( 0.1 3.1 ( 0.6
>7.37b 72 ( 18 >28c
a LC 50 value for MMPC 1 observed after 1 h of nanoparticle incubation. b Cells were 100% viable after 24 h of incubation with MMPC 2. Higher concentrations of nanoparticles could not be completely washed from wells, and interfered with absorbance readings. c Higher concentrations could not be tested due to decreasing visibility of the colonies on the nanoparticle-doped agar.
Vesicle Leakage Assay. This assay was adapted from a previous report (25). Vesicles were prepared by evaporating the solvent from SOPC (0.55 mL 25 mg/mL solution in CHCl3) or a mixture of SOPC/SOPS (0.5 mL 25 mg/mL:0.15 mL 10 mg/mL solution in CHCl3, resulting in an 8.5:1 mole ratio) with a stream of N2 (g). The dried lipids were resolubilized in 40 mM sodium phosphate buffer containing 100 mM calcein (further adjusted to pH 7 using NaOH and HCl) or 6.42 mM rhodamine. Three freeze-thaw cycles were then performed using a dry ice/ acetone bath. The vesicles were then either sonicated gently two times for 10 min each using a 150T aquasonic sonicator (VWR), followed by an additional freeze-thaw cycle, or extruded using a 1 µm polycarbonate membrane (Avanti Polar Lipids, Inc) (30, 31). Free fluorophore was removed by elution through a Sephadex 25 column with 90 mM sodium chloride and 10 mM sodium phosphate buffer adjusted to pH 7 or 7.4. Samples were prepared by diluting 0.2 or 0.3 mL of the collected vesicles to 2 mL using the sodium chloride buffer. The baseline fluorescence was corrected according to calibration of the nanoparticles with the fluorophore only or with Tritonlysed vesicles to control for light absorption by the particles. Components were added as aliquots of 1-10 µL of 0.005-10 mg/mL solutions, and the increasing fluorescence from vesicle lysis was monitored for 300 s at 515 nm (λex ) 490 nm) for calcein solutions and 554 nm (λex ) 524 nm) for rhodamine. The emission slit width was 1.5 or 3 nm. Complete leakage was caused by the addition of 50 µL 0.2% Triton X-100 in DMSO; the fluorescence intensity measured as a result was fixed at 100% vesicle lysis. RESULTS AND DISCUSSION
MMPCs 1 and 2 were tested for toxicity in red blood cells, Cos-1 cells, and bacteria (E. coli). LC50 values determined in each case are shown in Table 1. Toxicity profiles for the two mammalian cells are virtually indistinguishable, while a 2- to 3-fold increase is required as a lethal concentration for the bacterial cultures. This modest increase may be due to the nature of the bacterial assay: use of a low-density cross-linked agar matrix will decrease the mobility of both the cells and nanoparticles, potentially altering slightly the manner in which the two components interact. Alternatively, the difference observed may be due to the increased protection provided by the outer membrane and cell wall surrounding the Gram-negative E. coli (24), requiring a higher nanoparticle concentration to fully rupture the bacteria. The small variation in LC50 values observed suggests that the mechanism of toxicity is similar for the three cell types. This implies that the nanoparticles interact with the cells in a passive process, as the receptors and other related molecules active in energy-dependent processes are likely to be different for the separate species.
Toxicity of Gold Nanoparticles
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Figure 2. Comparison of MMPC 1 and 2 in disrupting vesicles with an overall negative (SOPC/SOPS; panel a) or neutral (SOPC only; panel b) charge. Nanoparticle concentration is 220 nM. Curves are meant to lead the eye.
Figure 3. Concentration dependence of vesicle rupture by MMPC 1. Higher concentrations could not be tested due to light absorption by the nanoparticles. Curves are meant to lead the eye.
Furthermore, the endocytic pathway is actively utilized by Cos-1 cells, but not erythrocytes, suggesting that toxicity is not caused by cellular uptake via endocytosis.1 One mechanism anticipated to be similar in comparing the eukaryotic and prokaryotic samples is membrane adhesion or cell lysis by the nanoparticles. Certainly the specific lipids in the bilayer for each cell type are different in identity or percent composition, but all cell types feature an overall negative charge with both highly charged and amphiphilic substituents (24). MMPC 1 would be drawn to the negative membrane, whereas MMPC 2 would not be attracted as strongly. Once bound to the cell, the amphiphilicity of the mixed monolayer of MMPC 1 could induce a variety of further interactions. To evaluate this potential mode of action, we performed a vesicle-disruption assay, in which a fluorescent dye is released from vesicles by molecules capable of interrupting the lipid bilayer. An increase in fluorescence in this assay is typically interpreted as an indication that a designed molecule may be antibacterial due to membrane leakage, as the bilayer is representative of the bacterial cell membrane (25). Two vesicle systems were examined, 1 We cannot rule out the possibility of these more complex pathways of internalization when nanoparticles are in use as drug vectors.
one composed of phosphotidylcholine (SOPC) and phosphotidylserine (SOPS) (net negative charge), and a second preparation of SOPC only (no net charge). MMPC 1 lysed the SOPC/SOPS vesicles more efficiently than MMPC 2 due to the electrostatic complementarity of the cationic substituents and the SOPS lipids (Figure 2a). In the case of the neutral SOPC vesicles, MMPC 2 showed higher levels of fluorophore release than MMPC 1, although the intensity was less than that of MMPC 1 with the negatively charged vesicles (Figure 2b). This result demonstrates that the specific charge pairing of the nanoparticles and lipid bilayers mediates membrane interaction and lysis to release the fluorophore. This process is concentration dependent, as demonstrated by the increased lysis of the negative SOPC/SOPS vesicles observed with increasing concentrations of MMPC 1 (Figure 3). The reason for the decrease in lysis rate at higher MMPC concentrations is unclear and is currently under investigation. In summary, we have demonstrated that the toxicity of the gold nanoparticles is related to their interactions with the cell membrane, a feature initially mediated by their strong electrostatic attraction to the negatively charged bilayer. We are currently designing nanoparticles with different properties to ameliorate the observed toxicity and will report our findings in due course. ACKNOWLEDGMENT
We thank Joseph Simard for synthesis of the nanoparticles, Jennifer Habink for the generous donation of the Escherichia coli cell line, Renuka Sivendran for advice concerning the top-agar assay, Lachelle Arnt for assistance with the hemolysis assay, and Nandini Chari and Firat Ilker for assistance with the vesicle assay (Univeristy of Massachusetts). This research was funded by NIH (GM 62998) C.M.G. acknowledges NIH ChemistryBiology Interface Training Grant GM 08515. LITERATURE CITED (1) Hruby, V. J. (2002) Designing peptide receptor agonists and antagonists. Nat. Rev. Drug Discovery 1, 847-858. (2) Opalinska, J. B., and Gewirtz, A. M. (2002) Nucleic-acid therapeutics: basic principles and recent applications. Nat. Rev. Drug Discovery 1, 503-14. (3) LaVan, D. A., Lynn, D. M., and Langer, R. (2002) Moving smaller in drug discovery and delivery. Nat. Rev. Drug Discovery 1, 77-84. (4) Sandhu, K. K., McIntosh, C. M., Simard, J. M., Smith, S. W., and Rotello, V. M. (2002) Gold nanoparticle-mediated transfection of mammalian cells. Bioconjugate Chem. 13, 3-6.
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